_______________________________________________________________________
Therapeutic interventions for Duchenne Muscular Dystrophy
– studies in the mdx mouse.
Hannah Grace Crabb
BSc(Hons)
(nee Hannah Grace Radley)
This Thesis is presented for the degree of Doctor of Philosophy
School of Anatomy and Human Biology
The University of Western Australia
September 2010
_______________________________________________________________________
________________________________________________________________Abstract
Abstract
Duchenne Muscular Dystrophy (DMD) is a lethal X-linked muscle wasting disease
resulting from defects in the myofibre subsarcolemmal protein dystrophin. The lack of
functional dystrophin protein leads to myofibre membrane fragility, repeated cycles of
myofibre necrosis and regeneration, and the eventual replacement of skeletal muscle
by fatty and fibrous connective tissue. DMD is characterised by progressive muscle
weakness and wasting with a limited life expectancy of approximately 20 years in
humans. Myofibre necrosis is the fundamental destructive process in DMD although
the exact mechanism by which the absence of dystrophin leads to myofibre necrosis is
unknown; inflammation, oxidative stress, metabolic abnormality, mislocalisation of
nNOS and increased intracellular calcium are all heavily implicated in the process. The
mdx mouse is a very useful animal model for DMD to examine the potential
mechanisms by which the absence of dystrophin leads to myofibre necrosis and to test
potential therapeutic interventions to reduce the extent of dystropathology in vivo.
While cell or gene therapy to replace the defective dystrophin is the definitive
treatment option for DMD, there are many problems to overcome before clinical
application. Corticosteroids remain the standard pharmacological treatment to
maintain muscle mass and help prolong life, despite severe adverse side-effects
including obesity, increased incidence of bone fractures, behavioural changes and
cataracts.
The overall aim of this Thesis was to examine, using the mdx mouse, the potential
benefits of 3 therapeutic interventions in vivo: 1) an anti-inflammatory drug, cV1q a
mouse specific tumour necrosis factor (TNF) antibody, 2) an antioxidant drug, Nacetylcysteine (NAC) a scavenger of reactive oxygen species (ROS), and 3) dietary
interventions, specifically a high fat (16%) and high protein (50%) diet. These
interventions were tested in both sedentary and exercised dystrophic mdx mice of
various ages, and assessed both histologically and biochemically using numerous
techniques. All 3 therapeutic interventions showed some benefits in dystrophic mdx
mice, confirming the involvement of inflammation, oxidative stress and metabolic
abnormality in myofibre necrosis in dystrophic muscle.
i
________________________________________________________________Abstract
Comprehensive analyses of metabolism, including protein synthesis rate, energy
expenditure and body composition, were also conducted using both young and adult
mice to test the hypothesis that dystrophic mdx have an altered metabolism compared
with control C57Bl/10 mice. Adult mdx mice show striking differences in response to a
high fat diet (16%) in comparison to control C57Bl/10 mice, they are also very lean,
with significantly increased muscle mass and increased protein synthesis rates. In
addition dystrophic mdx mice exhibit continuous cycles of myofibre necrosis and
regeneration and significant myofibre hypertrophy up to 24 weeks of age. These are
all very energy expensive procedures and may explain why mdx mice (with the same
energy intake compared to C57Bl/10 mice) do not exhibit the same negative sideeffects of a high fat diet. However, at this stage, it is still unclear if the metabolic
differences between strains are an innate feature of dystrophic muscle or a result of
the continuous cycles of myofibre necrosis and regeneration, and the associated high
energy demands of growing myofibres in dystrophic muscle.
Also contained in this Thesis is a standardised 30 minute treadmill protocol and timecourse analysis of molecular and cellular changes after a single 30 minute treadmill
exercise session. It was concluded that a single 30 minute treadmill session is a
sufficient and conveniently fast screening test to evaluate some of the benefits of preclinical drugs in vivo, which has major applications to speed up pre-clinical drug
screening in mdx mice.
A high level of variation in the extent of dystropathology in mdx mice is identified (and
addressed) through-out this Thesis. This emphasises the need for establishing Standard
Operating Procedures to enable comparison of data between laboratory groups worldwide when designing and conducting pre-clinical in vivo therapeutic trials in mdx mice.
These novel observations are of considerable interest to the field of preclinical DMD
research because they demonstrate, for the first time, the potential in vivo benefits of
3 new therapeutic interventions, the major metabolic differences between dystrophic
mdx and control C57Bl/10 mice, and the possible use of a 30 minute treadmill exercise
session as a fast and reproducible in vivo screening test.
ii
________________________________________________________Acknowledgements
Acknowledgements
I would like to sincerely thank and acknowledge the following people for their help in
completing this PhD Thesis.
My supervisor, Professor Miranda Grounds, your guidance, support and immense
knowledge of muscle biology has been invaluable. Thank you for absolutely everything.
Dr Marta Fiorotto, Miss Jessica Terrill, Dr Peter Arthur and Dr Thea Shavlakadze; you
are amazingly talented scientists and it was a pleasure to collaborate with you. I have
learnt so much from working with you and sincerely hope we can continue to do so in
the future. I would also like to extend my gratitude to everyone in the School of
Anatomy and Human Biology, in particular the members of my laboratory group,
fellow students in room 236, Mr Greg Cozens for expert advice on molecular biology
and Mr Guy Ben-Ary for assistance with image acquisition and analysis. I am very
appreciative of the Australian Postgraduate Award (APA) which supported my PhD
studies and the UWA Grant for Research Student Training (GRST) which enabled me to
complete an internship at Baylor College of Medicine, Houston, USA.
On a personal note, I must ever so gratefully thank my family and friends for their
endless support and encouragement. Last, but definitely not least, I need to thank my
husband Ben for his never ending support and understanding – you can have your wife
back now!
Thank you.
iii
__________________________________________________________Explanatory Note
Explanatory Note.
The regulations of The University of Western Australia allow candidates for the Doctor
of Philosophy degree (PhD) to present their work as a series of manuscripts rather than
as a conventional full text Thesis. These may be review manuscripts, or papers that
have been published, submitted but not accepted, papers in preparation, or any
combination of these. However, these papers must be related to the same central
theme and be integrated by a general introduction and discussion.
This Thesis is comprised of 7 published papers, 2 submitted for publication and 2
papers in preparation for submission (see pages v-vi for a full list of publications). This
series of papers is brought together in the Thesis by an Abstract of the Thesis, an
Introduction which contains unpublished material and 3 published review papers
(Publications #1-3), and the general Aims and Hypotheses of the Thesis. These 3
components form the introductory part of the Thesis (Chapter 1). Methodology is
described in General material and methods (Chapter 2) and also within the methods
section specific to each manuscript. A published methods manuscript (Publication #4)
is also included. Thereafter the Thesis contains 5 results chapters (Chapters 3-7) each
consisting of one research paper (Publications #5-9) that addresses the aims outlined
in the introductory part, plus an additional Results chapter (Chapter 8) from other
collaborative studies conducted during candidature (Publications # 10 & 11). Finally a
General Discussion integrates the body of knowledge derived from the Thesis and
establishes the significance of the work.
For the 6 published manuscripts the re-print is included with the original published
page numbering, figures and tables. The remaining manuscripts (submitted and/or in
preparation) are formatted specifically for the intended journal, each manuscript has
both an individual page numbering system and referencing system with references
listed at the end of each manuscript. All other parts of the Thesis have a common page
numbering system (bottom right-hand-side of the page) and a common referencing
system with all references listed at the end of the Thesis (Chapter 10).
iv
_________________________________________________________List of Publications
Hannah Crabb - Publications included in this PhD Thesis.
Publications are listed in order of appearance in the Thesis. The journal impact factor
and citation count of each publication is also listed. On publications, my name has
changed from Radley to Radley-Crabb (to reflect my married status).
General Introduction:
1) Radley HG, DeLuca A, Lynch GS, Grounds MD. (2007). Duchenne Muscular
Dystrophy: focus on pharmaceutical and nutritional interventions. Medicine in Focus:
International Journal of Biochemistry & Cell Biology. 39: 469-477. (IF 4.01) (24)
2) Grounds MD, Radley HG, Gebski BL, Bogoyevitch MA, Shavlakadze T. (2008).
Implications of cross-talk between TNF and IGF-1 signalling in skeletal muscle. Clinical
and Experimental Pharmacology and Physiology. 35(7): 846-51. (IF 2.04) (10)
3) Grounds MD, Radley HG, Lynch GS, Nagaraju K, De Luca AM. (2008). Towards
developing standard operating procedures for pre-clinical testing in the mdx mouse
model of Duchenne muscular dystrophy. Neurobiology of Disease. 31:1-19. (IF 4.38)
(19)
Methods:
4) Grounds MD, Radley-Crabb HG, Nagaraju K, Aartsma-Rus A, van Putten M, Rüegg
MA, de La Porte S. Hematoxylin and Eosin staining for histology of dystrophic skeletal
muscle M.1.2_007. TREAT NMD Neuromuscular Network. Pre-clinical testing for DMD:
Endpoints in the mdx mouse. http://www.treat-nmd.eu/research/preclinical/SOPs/
Results:
5) Radley HG, Davies MD, Grounds MD. (2008). Reduced muscle necrosis and longterm benefits in dystrophic mdx mice after cV1q (blockade of TNF) treatment.
Neuromuscular Disorders. 18: 227-238. (IF 2.67) (12)
6) Radley-Crabb HG, Terrill J, Shavlakadze T, Tonkin J, Arthur PG, Grounds MD. A single
30 minute treadmill exercise session is suitable for ‘proof-of concept studies’ in adult
mdx mice: a comparison of the early consequences of two different treadmill
protocols. Re-submitted to Neuromuscular Disorders in November 2010.
7) Terrill J, Radley-Crabb HG, Shavlakadze T, Arthur PG, Grounds MD. The in vivo
benefits of N-acetylcysteine treatment in dystrophic mdx mice. Submitted to
International Journal of Biochemistry and Cell Biology – January 2011.
v
_________________________________________________________List of Publications
8) Radley-Crabb HG and Grounds MD. The different impacts of a high fat diet on
dystrophic mdx and control C57Bl/10 mice. In preparation.
9) Radley-Crabb HG, Marini J, Grounds MD, Fiorotto M. Major metabolic differences
between dystrophic mdx and control C57Bl/10 mice. In preparation.
10) Piers AT, Lavin T, Radley-Crabb HG, Bakker AT, Grounds MD, Pinniger G. Blockade
of TNF (using cV1q antibody) in vivo reduces contractile dysfunction of skeletal muscle
in response to eccentric exercise in dystrophic mdx and normal mice. Neuromuscular
Disorders – Nov 3 [Epub ahead of print].
11) Klyen BR, Armstrong JJ, Adie SG, Radley HG, Grounds MD, Sampson DD. (2008).
Three-dimensional optical coherence tomography of whole muscle autografts as a
precursor to morphological assessment of muscular dystrophy in mice. J. Biomedical
Optics. 13 (1):011003. (IF 3.08) (1)
vi
______________________________________________________________ Declaration
Declaration
I hereby declare that the work contained within this Thesis is my own. Published work
and/or work in preparation for publication which has been co-authored is clearly
labelled and my individual contribution to the co-authored work is clearly stated.
___________________________
Hannah Crabb
September 2010
vii
_______________________________________________________________ Contents
Contents
Title
Abstract
i
Acknowledgements
iii
Explanatory Note
iv
List of publications
v
Declaration
vii
Contents
viii
Chapter 1 – General Introduction
1
1.1 Skeletal muscle structure and function
2
1.1.1 Plasticity of skeletal muscle (hypertrophy, atrophy, regeneration)
5
1.1.2 Skeletal muscle and disease
9
1.2 Duchenne Muscular Dystrophy
9
1.2.1 Current and potential therapies for DMD
11
1.2.2 Dystrophin and the dystroglycan complex
12
1.2.3 The absence of dystrophin
13
1.2.4 Animal models for Duchenne Muscular Dystrophy
14
1.3 The mdx mouse
15
1.3.1 Advantages and limitations of the mdx mouse
19
1.3.2 Exercise and the mdx mouse
19
1.3.3 Development of ‘Standard Operating Procedures’.
22
1.4 Myofibre necrosis in dystrophic muscle
22
1.4.1 Inflammation and dystrophic muscle
23
1.4.2 Anti-inflammatory therapies and dystrophic muscle
27
1.4.3 Reactive oxygen species and oxidative stress
29
1.4.4 Oxidative stress and dystrophic muscle
30
1.4.5 Anti-oxidant therapies and dystrophic muscle
31
1.5 Skeletal muscle metabolism
33
1.5.1 Protein turnover in skeletal muscle
34
1.5.2 Metabolic alterations in dystrophic muscle
34
1.5.3 Nutritional therapies and dystrophic muscle
37
1.6 Aims and hypotheses
39
viii
_______________________________________________________________ Contents
Publication 1: Radley HG, DeLuca A, Lynch GS, Grounds MD. (2007).
Duchenne Muscular Dystrophy: focus on pharmaceutical and nutritional
interventions. Medicine in Focus: International Journal of Biochemistry & Cell
Biology. 39: 469-477.
41
Publication 2: Grounds MD, Radley HG, Gebski BL, Bogoyevitch MA,
Shavlakadze T. (2008). Implications of cross-talk between TNF and IGF-1
signalling in skeletal muscle. Clinical and Experimental Pharmacology and
Physiology. 35(7): 846-51.
51
Publication 3: Grounds MD, Radley HG, Lynch GS, Nagaraju K, De Luca AM.
(2008). Towards developing standard operating procedures for pre-clinical
testing in the mdx mouse model of Duchenne muscular dystrophy.
Neurobiology of Disease. 31:1-19.
58
Chapter 2 – General materials and methods
75
2.1 In vivo animal protocols
76
2.2 Histology protocols
81
2.3 Molecular and biochemical protocols
83
2.4 Metabolism protocols
76
Publication 4: Grounds M.D, Radley-Crabb H.G, Nagaraju K, Aartsma-Rus A,
van Putten M, Rüegg M.A, de La Porte S. Hematoxylin and Eosin staining for
histology of dystrophic skeletal muscle M.1.2_007. TREAT NMD
Neuromuscular Network. Pre-clinical testing for DMD: Endpoints in the mdx
mouse. http://www.treat-nmd.eu/research/preclinical/SOPs/
89
Chapter 3 – Reduced muscle necrosis and long-term benefits in dystrophic
mice after cV1q (blockade of TNF) treatment.
101
Publication 5: Radley HG, Davies MD, Grounds MD. (2008). Reduced muscle
necrosis and long-term benefits in dystrophic mdx mice after cV1q (blockade
of TNF) treatment. Neuromuscular Disorders. 18: 227-238.
103
Chapter 4 – The early consequences of two different treadmill exercise
protocols on dystrophic mdx mice.
115
Publication 6: Radley-Crabb HG, Terrill J, Shavlakadze T, Tonkin J, Arthur PG,
Grounds MD. A single 30 minute treadmill exercise session is suitable for
‘proof-of concept studies’ in adult mdx mice: a comparison of the early
consequences of two different treadmill protocols. Re-submitted to
Neuromuscular Disorders in November 2010.
117
Chapter 5 – The benefits of N-acetylcysteine treatment in dystrophic mdx
mice.
158
Publication 7: Terrill J, Radley-Crabb HG, Shavlakadze T, Arthur PG, Grounds
160
ix
_______________________________________________________________ Contents
MD. The in vivo benefits of N-acetylcysteine treatment in dystrophic mdx
mice. Submitted to International Journal of Biochemistry and Cell Biology –
January 2011.
Chapter 6 – The different impacts of a high fat diet on dystrophic mdx and
control C57Bl/10 mice.
192
Publication 8: Radley-Crabb HG, Grounds MD. The different impacts of a high
fat diet on dystrophic mdx and control C57Bl/10 mice. In preparation.
193
Chapter 7 – A comparison of metabolism and protein synthesis rates in
young an adult dystrophic mdx and control C57Bl/10 mice.
238
Publication 9: Radley-Crabb HG, Marini J, Grounds MD, Fiorotto M. A
comparison of metabolism and protein synthesis rates in young an adult
dystrophic mdx and control C57Bl/10 mice. In preparation.
240
Chapter 8 – Additional work completed during candidature; not central to
the Thesis.
278
Publication 10: Piers AT, Lavin T, Radley-Crabb HG, Bakker AT, Grounds MD,
Pinniger G. Blockade of TNF (using cV1q antibody) in vivo reduces contractile
dysfunction of skeletal muscle in response to eccentric exercise in dystrophic
mdx and normal mice. Neuromuscular Disorders – Nov 3 [Epub ahead of
print].
280
Publication 11: Klyen BR, Armstrong JJ, Adie SG, Radley HG, Grounds MD,
Sampson DD. (2008). Three-dimensional optical coherence tomography of
whole muscle autografts as a precursor to morphological assessment of
muscular dystrophy in mice. J. Biomedical Optics. 13 (1):011003.
291
Chapter 9 – General Discussion
299
Chapter 10 - References
312
Appendix 1 – Additional dietary intervention studies.
AI
x
_____________________________________________________Chapter 1 - Introduction
Introduction:
Overview: The following is a comprehensive review of background literature pertinent
to the present study. The Introduction contains unpublished material and 3 published
review manuscripts. Details of each manuscript, the impact factor of the journal and
citation count (as of September 2010) along with the candidate’s individual
contribution to each manuscript are listed below. Re-prints of the 3 review
manuscripts are included at the end of the Introduction (Page 41-74).
1) Radley HG, DeLuca A, Lynch GS, Grounds MD. (2007). Duchenne Muscular
Dystrophy: focus on pharmaceutical and nutritional interventions. Medicine in Focus:
International Journal of Biochemistry & Cell Biology. 39: 469-477. (IF 4.01) (24).
Candidate completed approximately 85% of the work required for the manuscript.
2) Grounds MD, Radley HG, Gebski BL, Bogoyevitch MA, Shavlakadze T. (2008).
Implications of cross-talk between TNF and IGF-1 signalling in skeletal muscle. Clinical
and Experimental Pharmacology and Physiology. 35(7): 846-51. (IF 2.04) (10).
Candidate completed approximately 35% of the work required for the manuscript.
3) Grounds MD, Radley HG, Lynch GS, Nagaraju K, De Luca AM. (2008). Towards
developing standard operating procedures for pre-clinical testing in the mdx mouse
model of Duchenne muscular dystrophy. Neurobiology of Disease. 31:1-19. (IF 4.38)
(19).
Candidate completed approximately 40% of the work required for the manuscript.
1
_____________________________________________________Chapter 1 - Introduction
1.1 Skeletal muscle structure and function:
Skeletal muscle is the most abundant tissue in the human body comprising
approximately 40-50% of body mass; it is a highly specialised contractile tissue under
voluntary nervous control. The primary function of skeletal musculature is movement;
in addition to this muscles play a vital role in breathing, posture, heat regulation,
metabolism and reflexes. There are over 600 individual skeletal muscles in the human
body, individual skeletal muscles vary considerably in size, shape, and myofibre
arrangement; ranging from the extremely tiny stapedium of the middle ear, to very
large muscle masses such as the quadriceps of the thigh [Reviewed in (MacIntosh et al,
2006; Saladin, 2010)].
Despite extreme variation every skeletal muscle consists of both skeletal muscle cells
and connective tissue, and requires both a nerve and blood supply. Individual skeletal
muscles are enclosed by a tough connective tissue layer called the epimysium which
acts to give each muscle its shape. Bundles (fascicles) of skeletal muscle myofibres are
surrounded by perimysium, large blood vessels and nerves pass through perimysium.
Inside the fascicles, individual myofibres are each enclosed by a layer of endomysium
(Figure 1.1.1). The endomysium, approximately 60-120ηm in diameter, is a dense
network of extracellular proteins. Myofibres are long multi-nucleated muscle cells; the
cytoplasm of a myofibre is referred to as sarcoplasm and the plasma membrane as
sarcolemma. Located between the myofibre sarcolemma and the endomysium is the
specialised basement membrane rich in laminins and other proteins which play many
roles in signalling and scaffolding (Grounds et al, 2005c). Muscle satellite cells (undifferentiated mononuclear myogenic cells), are closely associated with the surface of
the myofibres, and represent a reserve of muscle precursor cells to facilitate growth
and muscle regeneration. Myofibres are peripherally nucleated and contain transverse
tubules, sarcoplasmic reticulum, mitochondria and myofibrils. Myofibrils are
specialised arrangements of contractile proteins organised into sarcomeres that are
responsible for myofibre contraction (Figure 1.1.2) [Reviewed in (Charge and Rudnicki,
2004; Grounds et al, 2005c; MacIntosh et al, 2006; Saladin, 2010)].
2
_____________________________________________________Chapter 1 - Introduction
Figure 1.1.1. Schematic gross structure of a skeletal muscle. Skeletal muscle is arranged in bundles. The
bundles are encased within three sheaths of connective tissue; epimysium covers the whole muscle,
perimysium covers the bundles of myofibres and the endomysium covers each individual myofibre.
th
Figure scanned from Saladin (2010) Anatomy and Physiology 5 Edition. Figure 10.1 ‘Connective tissue of
a Muscle’. Page 321.
3
_____________________________________________________Chapter 1 - Introduction
Figure 1.1.2. The internal organisation of an individual myofibre. Myofibres are composed of
sarcoplasma filled with striated myofibrils and are encased within a sarcolemma. Figure scanned from
th
Martini (2000) Fundamentals of Anatomy and Physiology 4 Edition. Figure 10.2 ‘Formation and
structure of a skeletal muscle fiber’. Page 280.
Skeletal muscle is often referred to as striated muscle as the myofibres appear
transversely striped or ‘striated’ when viewed longitudinally. This is due to the protein
arrangement within the myofibrils. Thick protein filaments (myosin) produce a dark A
band and thin protein filaments (actin, troponin and tropomyosin) produce a light I
band (Figure 1.1.2). The entire array of thick and thin filaments between the Z lines is
called a sarcomere. Shortening of the sarcomeres within a myofibril produces
shortening of the myofibril, shortening of the myofibre and ultimately contraction of
the skeletal muscle [Reviewed in (MacIntosh et al, 2006; Saladin, 2010)].
4
_____________________________________________________Chapter 1 - Introduction
1.1.1 Plasticity of skeletal muscle (hypertrophy, atrophy and regeneration):
The plasticity of adult skeletal muscle is outstanding, muscles are able to respond and
adapt to a wide variety of external stimuli including exercise, nutrition and
temperature, however skeletal muscles are also susceptible to injury after direct
trauma (e.g intense physical exercise, burns or laceration) or indirect causes such as
disease or genetic defects. The maintenance of adult skeletal musculature is facilitated
by the fine balance between protein synthesis and degradation and, the remarkable
ability of skeletal muscle to regenerate [Reviewed in (Charge and Rudnicki, 2004;
Karagounis and Hawley, 2010; Saini et al, 2009; Zierath and Hawley, 2004)].
Skeletal muscle hypertrophy
Skeletal muscle hypertrophy is an increase in the size and diameter (CSA - cross
sectional area) of individual myofibres. All postnatal muscle growth, including
hypertrophy, depends on the activation, proliferation and subsequent fusion of
satellite cells with existing myofibres. Muscle hypertrophy also requires protein
synthesis to exceed protein degradation. In adult skeletal muscle hypertrophy occurs
in response to resistance exercise, increased physiological loading or enhanced
nutritional uptake in vivo. Skeletal muscle adapts by increasing the size and amount of
contractile proteins within the sarcomeres of each myofibre, however due to the
fusion of satellite cells into hypertrophying myofibres the protein to DNA ratio of the
myofibre stays the same. This increase in contractile proteins leads to an increase in
the CSA of myofibres and associated force production, and also an alteration in protein
synthesis and degradation rates. Numerous factors and signals can also trigger muscle
hypertrophy, for example it is well known that Insulin-like growth factor-1 (IGF-1),
hormones and anabolic steroids have a crucial role in inducing skeletal muscle
hypertrophy in different situations [Reviewed in (Karagounis and Hawley, 2010; Saini
et al, 2009; Shavlakadze et al, 2010; Shavlakadze and Grounds, 2006)].
Skeletal muscle atrophy
When protein degradation outweighs protein synthesis skeletal muscles shrink;
atrophy is the reduction in CSA of individual myofibres, accompanied by a significant
5
_____________________________________________________Chapter 1 - Introduction
decrease in myofibrillar protein content, decreased force production, and lower
fatigue resistance. The biochemical and enzymatic responses to atrophy include the
reduced capacity to synthesize new protein and the up-regulation of pathways leading
to increased protein breakdown. Many factors (with different aetiologies) can lead to
muscle atrophy including aging, disuse (immobilisation), denervation, inflammatory
conditions associated with cancer (cachexia), burns and anorexia/starvation [Reviewed
in (Lynch et al, 2007; Saini et al, 2009; Shavlakadze and Grounds, 2003; Shavlakadze
and Grounds, 2006)].
Skeletal muscle necrosis and regeneration
Under normal conditions adult skeletal muscle is a stable tissue; however, when
damage to skeletal myofibres results in necrosis this initiates regeneration and the
formation of new muscle. In brief, the process of myofibre necrosis and regeneration is
as follows: 1) injury/damage to the sarcolemmal leading to myofibre necrosis, 2) resealing of the damaged ends of the myofibre, 3) infiltration of inflammatory cells and
phagocytosis
of
myofibre
debris,
4)
hyper-contraction
of
myofibrils,
5)
revascularisation of injured myofibres, 6) activation and proliferation of muscle
satellite cells, 7) fusion of myoblasts into myotubes and fusion of these with damaged
myofibres (Figure 1.1.3), 8) re-innervation, growth and maturation of regenerated
myofibres [Reviewed in (Charge and Rudnicki, 2004; Grounds, 1991)].
Figure 1.1.3. Regeneration of a damaged skeletal myofibre. Damage to skeletal muscle initiates a
reaction in which satellite cells are activated and become proliferating myoblasts. Myoblasts migrate (if
necessary) to the damaged site where they fuse into myotubes, which then fuse with the existing end of
damaged myofibres.
6
_____________________________________________________Chapter 1 - Introduction
The term ‘myofibre necrosis’ refers to the irreversible breakdown of the sarcolemma
and all sarcoplasmic structures, necrosis can occur through the whole length of a
myofibre or only part of the myofibre can degenerate. Partial degeneration of
myofibres (focal necrosis) is common in Duchenne Muscular Dystrophy (DMD). There
are numerous causes of myofibre necrosis: disease (polymyositis, muscular
dystrophies), eccentric exercise, ischemia, direct trauma (crush, cut, tear),
transplantation (grafting), thermal injury or biological toxins (venom, streptococcus A)
[Reviewed in (Engel and Franzini-Armstrong, 1994; MacIntosh et al, 2006; Wallace and
McNally, 2009)].
A transient inflammatory response is the body’s natural reaction to acute tissue injury
resulting from mechanical, physical or chemical damage. In skeletal muscle, the
inflammatory response classically consists of early vascular changes producing
oedema, the activation of resident leukocytes (mast cells and macrophages) and
infiltration of additional leukocytes e.g. neutrophils (within the hour after muscle
damage), release of pro-inflammatory cytokines and assistance with tissue
phagocytosis and myofibre regeneration [Reviewed in (MacIntosh et al, 2006; Saini et
al, 2009; Tidball, 1995)]. It is widely documented that excess (or dysregulated)
inflammation may exacerbate muscle damage in both Duchenne Muscular Dystrophy
(DMD) patients and the mdx mouse model of DMD; thus much pre-clinical research is
conducted into anti-inflammatory drugs as potential therapeutic interventions for
DMD, while waiting for gene/cell therapies to correct the fundamental gene defect
(Evans et al, 2009a; Radley et al, 2008; Radley et al, 2007).
(For more detailed information on ‘The role of inflammation in Duchenne Muscular Dystrophy’, see
sections 1.4 of this Introduction).
To minimise necrosis after injury, the damaged portion of the myofibre must be rapidly
sealed off from the nearby undamaged sarcoplasm. Within 3 hours after damage,
hyper-contraction of sarcomeric structures separates the undamaged viable parts of
myofibre from the necrotic area. By 9-12 hours the necrotic tissue section is totally
sealed off from the remaining viable parts of the damaged myofibre by formation of
new plasmalemma (Grounds, 1991; Papadimitriou et al, 1990).
7
_____________________________________________________Chapter 1 - Introduction
Myofibre necrosis can occur in the presence of a blood supply (e.g eccentric exercise)
or in conditions of ischemia. Ischemic myofibre necrosis and regeneration occurs in
transplanted whole or minced muscle grafts due to the loss of blood supply. In
transplants (grafts) of whole muscles, some myofibres at the periphery do not
degenerate due to diffusion of nutrients and gases from vessels in adjacent tissues
(skin & muscle), new muscle formation in the centre of the graft is normally excellent
suggesting a strong resistance to ischemia (Grounds, 1991; Grounds et al, 2005a;
Radley et al, 2008; White et al, 2000). However, in very large grafts the central zone
fails to re-vascularise, does not regenerate successfully and becomes fibrous, this may
be due to prolonged ischemia which can favour the proliferation of fibroblasts.
Revascularisation is an important event for ensuring successful new muscle formation
[Reviewed in (Grounds, 1991; MacIntosh et al, 2006)].
Tissue resident adult stem cells play a crucial role in the maintenance of many body
tissues and have been identified in bone marrow, brain, liver, intestines, heart and
skeletal muscle [Reviewed in (Boonen and Post, 2008)]. Satellite cells, the resident
stem cells of adult skeletal muscle, play a pivotal role in both postnatal growth and
regeneration of skeletal muscle. Skeletal myofibres are terminally differentiated cells
and cannot re-enter the mitotic cell cycle. Satellite cells are normally quiescent,
however damage to skeletal muscle results in activation of satellite cells (then called
myoblasts), they proliferate and migrate (if necessary) to the damaged site where they
differentiate and fuse with existing or damaged myofibres (Figure 1.1.3) [Reviewed in
(Boonen and Post, 2008; Buckingham and Montarras, 2008; Charge and Rudnicki,
2004; Grounds, 1991)].
The early events of muscle regeneration involving inflammation, re-vascularisation and
activation and fusion of satellite cells into new myotubes, does not require
innervation. However, reinnervation is a key factor in the restoration of muscle
function and successful formation of synapses and motor endplates which connect
muscles to the appropriate motorneurons completes the maturation of the newly
regenerated muscle [Reviewed in (Grounds, 1991)].
8
_____________________________________________________Chapter 1 - Introduction
When sarcolemmal damage is repaired and patched locally the myofibre avoids the
response of necrosis (and subsequent regeneration is not required) (Bansal et al, 2003;
Doherty and McNally, 2003). The process of skeletal muscle necrosis and regeneration
is central to the lethal muscle disease DMD, which is the focus of this Thesis.
(For more detailed information on ‘Duchenne Muscular Dystrophy’, see section 1.2 of this Introduction).
1.1.2 Skeletal muscle and disease:
There are numerous disorders and diseases that can heavily affect skeletal muscle. The
most common disorder is sarcopenia; the term used to describe the loss of skeletal
muscle mass and strength that inevitably occurs with aging (Evans, 2004; Saini et al,
2009; Shavlakadze and Grounds, 2003). Neuromuscular diseases such as Parkinson’s or
Multiple Sclerosis affect both skeletal muscle and/or their nervous control. Examples
of other disease that affect muscle are the congenital myopathies (Central core
disease, Nemaline myopathy and Minicore myopathy), mitochondrial myopathies,
metabolic disorders (Pompe disease, McArdle disease and glycogen storage disease),
inflammatory myopathies (dermatomyositis, polymyositis and inclusion body mitosis),
and the muscular dystrophies (Duchenne, Becker, Emery-Dreifuss, limb-girdle and
facioscapulohumeral) [Reviewed in (Hilton-Jones and Kissell, 2010)].
(For more detailed information of Duchenne Muscular Dystrophy, see section 1.2 of this Introduction).
1.2 Duchenne Muscular Dystrophy:
The muscular dystrophies are a set of inheritable muscle disorders, characterised by
progressive muscle weakness and wasting, and elevated blood serum creatine kinase
(CK) level. Based on the distribution of predominant muscle weakness seven major
forms of dystrophy can be defined; Duchenne muscular dystrophy, Becker muscular
dystrophy, Congenital muscular dystrophy, Emery-Dreifuss muscular dystrophy, Distal
muscular dystrophy, Facioscapulohumeral muscular dystrophy, Occulopharyngeal
muscular dystrophy and Limb-Girdle muscular dystrophy. Duchenne muscular
dystrophy (DMD) is one of the most severe and the most common, affecting
approximately 1/3500-6000 male births [Reviewed in (Bushby et al, 2010a; Emery,
2002; Manzur and Muntoni, 2009; Sinnreich, 2010)].
9
_____________________________________________________Chapter 1 - Introduction
DMD is caused by a mutation in the X-linked gene that encodes for the sarcolemmal
protein dystrophin, complete absence or impaired function of the dystrophin protein
results in DMD. The dystrophin gene the largest in the human genome known to date,
a feature which results in a very high spontaneous mutation rate. The gene is
approximately 3 million base pairs in length, containing 79 exons; the full-length
dystrophin protein is 427kDa in molecular mass [Reviewed in (Blake et al, 2002; Le
Rumeur et al, 2010; Sinnreich, 2010)]. The gene is located on the short arm of the X
chromosome (Xp21) thus the majority of DMD patients are male, however up to 10%
of female carriers also manifest clinical symptoms due to the relative proportion of Xchromosomes that are inactivated (Matthews et al, 1995; Wenger et al, 1992). The
majority of DMD patients inherit their dystrophin gene mutation from a carrier
mother, however; approximately 30% of DMD cases occur in a child with no positive
family history as the result of a de novo mutation (Manzur and Muntoni, 2009). Germ
line mosaicism also complicates the rate of de novo mutations, with an 8.6%
recurrence rate of a de novo mutation in the same healthy mother (Helderman-van
den Enden et al, 2009).
Approximately 70% of DMD patients have a gene mutation located in the mutation
‘hot-spot’ in the central genomic region (exons 45–53) of the dystrophin gene
[Reviewed in (Walmsley et al, 2010)]. The dystrophin deficient phenotype varies from
the severe Duchenne (DMD) phenotype to the milder allelic form known as Becker
Muscular Dystrophy (BMD). Mutations that result in the premature truncation of
dystrophin typically result in the severe Duchenne phenotype because no functional
dystrophin is produced; patients with the milder Becker phenotype have partial
deficiencies of dystrophin that still enable translation of a semi-functional dystrophin
protein [Reviewed in (Bushby et al, 2010a; Hoffman, 2001; Sinnreich, 2010)]. Most
dystrophin gene mutations involve deletions or duplications of whole exons; less often,
small point mutations are seen (Deburgrave et al, 2007).
Patients with DMD begin to show clinical symptoms (Gower’s manoeuvre and
hypertrophied calf muscles) around 3 years of age. The absence of dystrophin in DMD
patients is confirmed by immunohistochemical staining of a muscle biopsy, genetic
10
_____________________________________________________Chapter 1 - Introduction
testing is also performed to determine the exact gene mutation(s). In young boys,
walking is delayed and awkward and as the disease advances patients undergo a
progressive loss of muscle mass and function; most boys lose the ability to ambulate
and are wheelchair bound by 12 years of age. Loss of respiratory function usually
begins early in the second decade and progresses to respiratory failure in the mid to
late teens (Bushby et al, 2010a; Emery, 2002; Sinnreich, 2010). Many patients affected
with the disease also have cognitive impairment (Giliberto et al, 2004; Polakoff et al,
1998). Without intervention the average survival age for an individual with DMD is
approximately 19 years, with death occuring due to the overwhelming loss of muscle
mass, respiratory and/or cardiac complications. However, corticosteroid, cardiac,
respiratory and orthopaedic interventions have led to major improvements in quality
of life, health and longevity; and children who are diagnosed with DMD today, have a
potential life expectancy of up to 40 years [Reviewed in (Biggar, 2006; Bushby et al,
2010a; Toussaint et al, 2007)].
1.2.1 Current & potential therapies for Duchenne Muscular Dystrophy:
There is currently no cure or definitive treatment for DMD and the disease is fatal. The
existing treatment for DMD patients is multidisciplinary including psychosocial
management, palliative care and pharmacological treatment with corticosteroids
(Bushby et al, 2010a; Bushby et al, 2010b). Corticosteroids offer the only avaliable
method for preserving some muscle function; however their exact mechanism of
action in dystrophic muscle remains unknown and they are not effective in all patients.
Two corticosteroids, prednisone and deflazacort have been used extensively in DMD
patients and they appear to be equally effective in preserving skeletal muscle function.
Unfortunately both drugs are associated with adverse side-effects although these,
particularly weight gain, are less severe with deflazacort (Biggar et al, 2006; Campbell
and Jacob, 2003). Recent critical review of corticosteroid use in DMD patients conclude
that the use of steroids is limited due to detrimental side-effects but they are the best
pharmacological treatment option currently available (Angelini, 2007; Manzur et al,
2008). Cell or gene therapy to replace the defective dystrophin is the ideal scenario
and there have been many promising breakthroughs in the last 10 years, however
interpretation of efficacy is sometimes complicated by the presence of dystrophin
11
_____________________________________________________Chapter 1 - Introduction
positive revertant myofibres (Arechavala-Gomeza et al, 2010) and there are many
problems to overcome before clinical application of these new potential interventions
can become wide-spread (Bushby et al, 2009; Cossu and Sampaolesi, 2007; Grounds
and Davies, 2007; Guglieri and Bushby, 2010; Manzur and Muntoni, 2009; Nagaraju
and Willmann, 2009; Odom et al, 2007; Partridge, 2010; Tremblay and Skuk, 2008;
Wells, 2008).
Therapeutic approaches for DMD fall into three main strategies: (i) replacement of
dystrophin by cellular, genetic or molecular interventions; (ii) enhancement of muscle
regeneration and (iii) reduced muscle necrosis [Reviewed in (Bushby et al, 2009;
Manzur and Muntoni, 2009; Radley et al, 2007; Tidball and Wehling-Henricks, 2004a)].
The latter approach is the main focus of this thesis which examines pharmacological
treatment and dietary interventions as potential therapies to reduce muscle necrosis
in dystrophic mdx mice.
1.2.2 Dystrophin & the Dystroglycan complex:
Through the use of different promoters, dystrophin is located in skeletal, smooth and
cardiac muscle, neurons, Schwann cells and retinal synapses [Reviewed in (Blake et al,
2002; Hoffman, 2001; Sinnreich, 2010); however, skeletal muscle is by far the most
severely affected in DMD and is the focus of this thesis. In normal skeletal muscle,
dystrophin is located beneath the myofibre sarcolemma and forms part of the
mechanical link between the internal cytoskeleton and the external extracellular
matrix (ECM) of a myofibre. Dystrophin is part of the dystroglycan complex (DGC)
which spans through the sarcolemma (Figure 1.2.1). Dystrophin links proteins within
the cell sarcolemma to the cell cytoskeleton and plays a vital role in membrane
stability and muscle strength (Grounds et al, 2005c; Lapidos et al, 2004; Roberts,
2001). The DGC also plays a vital role in cell signalling (Rando, 2001; Wehling et al,
2001). Dystrophin binds to cytoplasmic f–actin and, within the DGC, to sarcolemmal βdystroglycan (βDAG), that binds to extracellular α-dystroglycan, that binds to laminin
α2 in the basement membrane, ultimately joining the myofibre sarcolemma to the
ECM (Grounds et al, 2005c).
12
_____________________________________________________Chapter 1 - Introduction
Figure 1.2.1. The dystroglycan complex. Dystrophin (shown in red) is absent in DMD leaving dystrophic
myofibre membranes fragile and susceptible to contraction induced damage. Adapted from Roberts et
al (2001) (Roberts, 2001).
1.2.3 The absence of dystrophin:
It is well established that impaired function or absence of the skeletal muscle protein
dystrophin, and numerous other dystrophin-associated glycoproteins, render
dystrophic myofibres susceptible to sarcolemma damage during mechanical
contraction (Gailly, 2002; Ohlendieck and Campbell, 1991; Petrof et al, 1993; Reed and
Bloch, 2005; Watchko et al, 2002). Dystrophic myofibres are particularly vulnerable to
damage after eccentric lengthening contractions (Brussee et al, 1997; Tegeler et al,
2010; Vilquin et al, 1998). The absence of functional dystrophin protein leads to
myofibre membrane fragility, and myofibre necrosis. While regeneration is initially
effective, repeat cycles of myofibre necrosis and subsequent regeneration (associated
with inflammation and accumulating fibrosis) eventually leads to the replacement of
myofibres by fatty and/or fibrotic connective tissue. The specific mechanism(s) leading
to myofibre necrosis are still unclear, yet there are strong associations with excessive
inflammation, increased intracellular calcium levels, elevated oxidative stress and
metabolic abnormality (Davidson and Truby, 2009; Evans et al, 2009a; Even et al, 1994;
Kuznetsov et al, 1998; Radley et al, 2008; Tidball and Wehling-Henricks, 2007;
Whitehead et al, 2008).
(For more detailed information on Skeletal Muscle Necrosis, see section 1.4 of this Introduction).
13
_____________________________________________________Chapter 1 - Introduction
The absence of dystrophin protein in the DGC also causes the incorrect anchoring of
neuronal nitric oxide synthase (nNOS) at the myofibre sarcolemma. Nitric oxide is
produced after increased muscle activity and plays an important role in regulating
vasodilation and oxygenation. Disruptions in nNOS expression and signalling negatively
impacts dystropathology [Reviewed in (Le Rumeur et al, 2010)]. Deregulation of nNOS
may also contribute to high levels of reactive oxygen species (ROS) in dystrophic
muscle (Grisotto et al, 2000; Thomas et al, 1998; Wehling et al, 2001).
(For more detailed information on The Role of ROS in Skeletal Muscle Necrosis, see section 1.4 of this
Introduction).
1.2.4 Animal models for Duchenne Muscular Dystrophy:
Several animal research models for DMD have been identified and provide a wealth of
information on the pathophysiology of dystrophin deficiency. Each animal model
differs from both DMD patients and other animal models [Reviewed in (Allamand and
Campbell, 2000; Banks and Chamberlain, 2008; Collins and Morgan, 2003; Grounds et
al, 2008b; Partridge, 1997; Vainzof et al, 2008; Willmann et al, 2009)]. While
dystrophin deficient worms (Caenorhabditis elegans) and zebra fish (Danio rerio) have
been identified and described [Reviewed in (Collins and Morgan, 2003)], only the
mammalian models are discussed here. The most widely used animal model is the mdx
mouse (C57BL/10ScSnmdx/mdx) which was discovered during a screening for mutant
glycolysis enzymes and occurred as a spontaneous mutation on the C57BL/10ScSn
background (Bulfield et al, 1984). The mdx mouse (mdx/mdx homozygous females and
mdx/Y heterozygous males) has total absence of the dystrophin protein and elevated
serum CK levels. Mdx mice show some progressive disease symptoms, although they
are not severe until later life [Reviewed in (Grounds et al, 2008b)].
(For more detailed information of the mdx mouse please see section 1.3 of this Introduction).
At least 2 dystrophin deficient cat models of muscular dystrophy (HFMD –
Hypertrophic Feline Muscular Dystrophy) have been identified. Both show cycles of
skeletal muscle necrosis and regeneration, marked myofibre hypertrophy and severe
cardiac symptoms. Lingual and/or diaphragmatic muscle hypertrophy becomes so
14
_____________________________________________________Chapter 1 - Introduction
severe that dystrophic cats succumb to de-hydration and/or starvation; the HFMD cats
are not widely used in pre-clinical DMD research [Reviewed in (Carpenter et al, 1989;
Collins and Morgan, 2003; Hoffman, 2001; Partridge, 1997)].
A sporadic mutation in golden retriever dogs (GRMD – Golden Retriever Muscular
Dystrophy) discovered in 1983 was found to cause muscular dystrophy and follow Xlinked recessive inheritance; the causative mutation was found to be a splicing
mutation in exon 8 of the dystrophin gene. GRMD dogs most closely resemble DMD
patients with a rapid course of muscle weakness, wasting and fibrosis. Severe onset of
the disease is around 3 months of age with death occurring anywhere from the
neonatal period to adulthood (Hoffman, 2001; Kornegay et al, 1988). GRMD colonies
are challenging to work with due to their extremely severe phenotype, large size (2530kg), high maintenance costs and high variability in muscle condition both between
and within individual dogs (Ambrosio et al, 2009; Zucconi et al, 2010).
Recently, smaller dog animal models such as the Beagle, German Short Haired Pointer
and Cavalier King Charles Spaniel have been established (Schatzberg et al, 1999;
Walmsley et al, 2010). A dystrophic beagle colony, established and phenotyped in
Japan (CXMDJ – Canine X-linked Muscular Dystrophy in Japan) has an obvious
dystrophic phenotype, with a high neonatal death rate, elevated serum CK levels,
diaphragmatic and limb muscle degeneration, and severe dysphagia and macroglossia
in ‘old’ animals (Shimatsu et al, 2003; Shimatsu et al, 2005), although their phenotype
is less severe and progresses more slowly in comparison to the Golden Retrievers.
CXMDJ dogs have recently been the animal model of choice for pre-clinical cell and
gene therapy studies due to their severe phenotype yet smaller size and thus greatly
reduced pre-clinical research costs.
1.3. The Mdx mouse - overview:
The classic biochemical and genetic mouse model of DMD is the mdx mouse
(C57BL/10ScSnmdx/mdx), a mutant with an inherited X-linked recessive trait, discovered
in 1981 in a C57BL/10ScSn colony at the University of Leicester, UK (Bulfield et al,
1984; Hoffman et al, 1987). Dystrophic mdx mice have been widely used for pre15
_____________________________________________________Chapter 1 - Introduction
clinical DMD research and intensively described in numerous papers (Coulton et al,
1988a; Coulton et al, 1988b; Grounds et al, 2008b; Willmann et al, 2009). The absence
of dystrophin is due to a point mutation on the X chromosome, resulting in a
premature stop codon in the mRNA of exon 23 of the dystrophin gene (Sicinski et al,
1989). The mdx mouse has no detectable dystrophin protein in skeletal muscles;
however, sporadic revertant myofibres can express dystrophin which can complicate
the interpretation of gene or cell replacement studies (Hoffman et al, 1990; Yokota et
al, 2006).
The dystropathology of mdx mice begins early in embryonic development with the
absence of dystrophin resulting in disrupted myogenesis, changes in skeletal and
cardiac muscle patterning (Merrick et al, 2009) and disorganisation of Z-discs (Torres
and Duchen, 1987). Some muscle necrosis is observed from 5 days of age in muscles of
the head and shoulder and occasionally in the limb muscles from 14 days (Torres and
Duchen, 1987). It must also be noted that a low level of apoptosis precedes any
detectable necrotic changes in mdx muscle, and that this low level of apoptotic events
continues in dystrophic muscle through the life cycle of the mdx mouse (Lim, 2004;
Tidball et al, 1995). Dystrophic mdx mice experience severe dystropathology in both
skeletal (Figure 1.3.1) and cardiac muscle (Grounds et al, 2008b; McNally and
MacLeod, 2005; Zhang et al, 2008). Cardiac muscle falls outside the scope of this Thesis
and only skeletal muscle is discussed here.
Mdx mice undergo an onset of severe skeletal myofibre necrosis and subsequent
regeneration in limb and paraspinal muscles postnatally around 3 weeks of age (Figure
1.3.1) (Coulton et al, 1988b; Grounds et al, 2008b). The high level of muscle necrosis
between 21 and 28 days provides an excellent model to study therapeutic
interventions designed to prevent or reduce muscle necrosis, as a reduction in
dystropatholgy is easily observed (Grounds and Torrisi, 2004; Radley et al, 2008;
Radley and Grounds, 2006; Shavlakadze et al, 2004; Stupka et al, 2006). Skeletal
muscle necrosis peaks around 4 weeks (Figure 1.3.2) and then decreases significantly
and stabilises by 8-12 weeks of age to a relatively low level of damage where
approximately 6% of each skeletal muscle is actively necrotic (McGeachie et al, 1993)
(Radley-Crabb et al, submitted). Beyond 12 weeks of age, mdx mice show few clinical
16
_____________________________________________________Chapter 1 - Introduction
dystrophic symptoms, they exhibit a low level of muscle necrosis and creatine kinase
levels reduce dramatically, there is however a high level of cumulative
damage/regeneration and a fibrosed diaphragm; the mdx diaphragm is the most
severely affected muscle and its pathology most closely resembles that of DMD
patients (Dupont-Versteegden and McCarter, 1992; Lynch et al, 1997; Stedman et al,
1991). Beyond 20 months of age, mdx mice exhibit a more severe dystropathology;
that is, the extensive replacement of skeletal muscle with fatty and/or fibrous
connective tissue, reduced cardiac and respiratory function, reduced motor strength
and a slightly shortened lifespan (Chamberlain et al, 2007; Lefaucheur et al, 1995;
Willmann et al, 2009). The striking differences in disease severity between mdx mice
and DMD patients, may largely reflect vast differences in the growth phase, final body
size and life span of the two species [Reviewed in (Grounds, 2008; Grounds et al,
2008b; Willmann et al, 2009)].
A
A
B
B
Figure 1.3.1 Myofibre necrosis in Haematoxylin and Eosin stained dystrophic mdx muscle. (A) Prenecrotic muscle in 20 day old mdx quadriceps characterised by peripheral nuclei and a normal
appearance. (B) Necrotic muscle in 23 day old mdx quadricpes (outlined in yellow), characterised by
inflammatory cells and fragmented sarcoplasm. Scale bar represents 100µm.
Some muscles of the head region, including extraocular muscles (Fisher et al, 2005),
masseter (Muller et al, 2001) and the laryngeal muscles (Marques et al, 2007; Smythe,
2009), show a very mild dystropathology and are relatively spared from myonecrosis.
The reasons for the mild dystropathology are not clear, although it is noted that these
muscles may have an improved ability to regulate calcium homeostasis (Khurana et al,
1995; Marques et al, 2007) and selected mechanical properties that offer resistance to
damage (Wiesen et al, 2007).
17
_____________________________________________________Chapter 1 - Introduction
Muscle necrosis
Peak
Onset
1
2
3
4
Age in weeks
6
7
8
9
10
Figure 1.3.2 The relative level of myofibre necrosis across the first 10 weeks of life in a mdx mouse.
There is an acute bout of myofibre necrosis around 3-4 weeks of age, beyond this myofibre necrosis
remains at a low level.
The stimulus for the acute onset of myofibre necrosis around 21 days (Figure 1.3.2) is
unknown. Studies suggest that it may be the result of an increase in motor activity
(movement) at the time of weaning (Mokhtarian et al, 1995), onset is also associated
with huge disruptions in intracellular signalling and excessive inflammation [Reviewed
in (Evans et al, 2009a)]. Hind limb immobilization in young mdx mice (d14-28) results in
reduced myofibre necrosis (and subsequent regeneration), suggesting that myofibre
contraction and movement play an important role in the onset of myofibre necrosis
(Mokhtarian et al, 1999).
The dystrophic mdx mouse strain (C57Bl/10ScSnmdx/mdx) is an inbred strain, therefore
unlike humans, both female (homozygous for the absence of dystrophin) and male
mice (heterozygous) are used in pre-clinical research. Both sexes exhibit a similar
pattern of myofibre necrosis and regeneration across their lifespan and have
consistently elevated serum CK levels. However, specific differences have been
documented between sexes – for example male mdx mice have a 2x fold higher blood
serum CK level compared to age matched females at 12 weeks of age, yet they have a
4x fold lower serum CK level at 48 weeks of age (Salimena et al, 2004). It is
recommended that when conducting pre-clinical research, experimental groups
consistent of only one sex (Grounds et al, 2008b). A high level of variation in the extent
of dystropathology (determined histologically) between individual mice is also seen in
age, sex and muscle matched mdx mice (Radley and Grounds, 2006; Spurney et al,
2009)(Radley-Crabb et al, submitted).
18
_____________________________________________________Chapter 1 - Introduction
1.3.1 Advantages / Limitations of the mdx mouse:
The suitability of the mdx mouse model for pre-clinical DMD research has been the
subject of numerous reviews, conferences and recent discussions (Grounds et al,
2008b; Nagaraju and Willmann, 2009). The advantages and limitations of working with
the mdx mouse strain are summarised in Table 1.3.1.
Advantages
Limitations
Small mammal – low husbandry costs in
Mild phenotype in comparison to DMD patients
comparison to GRMD dogs
and GRMD dogs
Large litters (4-8 pups)
High neonatal mortality rate
Short gestation period (19 – 21 days)
High variation in dystropathology (within and
between both individual mice and whole litters)
Small mammal – low costs to pharmacologically
High number of mdx mice are required in pre-
treat mice in comparison to GRMD dogs
clinical studies due to high level of variation in
dystropathology
Peak in dystropathology at 3-4 weeks of age
Mild phenotype in adult mdx mice - adult mdx
mice require exercise in pre-clinical studies
Willingness to exercise
Marked muscle hypertrophy as adults
(uncharacteristic of DMD patients and GRMD dogs)
Table 1.3.1 Summary of the main advantages and limitations of using the mdx mouse in pre-clinical
DMD research.
1.3.2 The mdx mouse and exercise:
The low level of dystropathology in adult mdx mice can be made significantly worse by
exercise that increases myofibre necrosis and reduces muscle strength (Brussee et al,
1997; Granchelli et al, 1996; Okano et al, 2005; Vilquin et al, 1998), thus enabling
potential therapeutic interventions to be evaluated in adult mice (Archer et al, 2006;
De Luca et al, 2005; Granchelli et al, 2000; Payne et al, 2006; Radley et al, 2008; Radley
and Grounds, 2006). Exercise is a less invasive and a more clinically relevant option in
comparison to previous methods (irradiation) designed to increase the extent of mdx
dystropathology (Wakeford et al, 1991).
19
_____________________________________________________Chapter 1 - Introduction
Our laboratory has used voluntary wheel exercise over 48 hours to increase myofibre
necrosis in adult dystrophic mdx mice and to histologically assess the beneficial effects
of various anti-inflammatory drugs on the muscles of adult mdx mice (Hodgetts et al,
2006; Radley et al, 2008; Radley and Grounds, 2006). Muscle necrosis is roughly
doubled (increases from ~6 to 12%) in quadriceps muscle after 48 hours of voluntary
exercise, although other muscles such as the tibialis anterior (TA) are barely affected
(Archer et al, 2006; Radley et al, 2008). Histological analysis of muscle necrosis after 48
hours of voluntary exercise, allows induced necrosis to be readily measured without
the ensuing complication of regeneration and new muscle formation, as new
myotubes appear only after pproximately 3 days after injury (McGeachie et al, 1993).
Voluntary wheel exercise is inexpensive and causes minimal stress to the animal since
the mice (which are nocturnal) run voluntarily during the night (Hara et al, 2002). Mdx
mice tend to run less total distance (km) and at a lesser speed (km/hr) than nondystrophic control mice (Brussee et al, 1997; Grounds et al, 2008b; Hayes and
Williams, 1996). The running patterns of mdx mice during voluntary exercise have
previously been used as a non-invasive method to determine genotype, as mdx mice
run a very distinct intermittent (stop-start) running pattern in comparison to a
continuous running pattern seen in non-dystrophic mice (Hara et al, 2002; Radley and
Grounds, 2006). However, long-term voluntary exercise has shown beneficial effects
on muscle strength and fatigue resistance in the mdx mouse (Call et al, 2008; DupontVersteegden et al, 1994b; Hayes and Williams, 1996; Landisch et al, 2008) which may
complicate the long-term analysis of pre-clinical drug interventions.
A widely used alternative to voluntary exercise is forced treadmill running. This occurs
at a controlled speed and for a pre-determined length of time, thus potentially
eliminating some of the variables experienced with voluntary exercise. Treadmill
running is usually carried-out for convenience during the day, although normally mice
are active at night and may benefit from being run at night as daytime exercise may be
stressful for them [Reviewed in (Grounds et al, 2008b)]. A protocol of 30 minutes
running on a horizontal treadmill at a speed of 12m/min, twice a week for at least 4
weeks, significantly increases the dystropathology of adult mdx mice (Burdi et al, 2009;
20
_____________________________________________________Chapter 1 - Introduction
De Luca et al, 2005; De Luca et al, 2008; De Luca et al, 2003; Granchelli et al, 2000).
The in vivo weakness produced by the above protocol is observed exclusively in mdx
mice, with no effect in normal control C57Bl/10 mice (De Luca et al, 2003). There are
concerns regarding such forced treadmill running because mdx mice can have
problems coping and thus be reluctant to run, although a short warm-up period at a
slower speed and training of the mdx mice to accustom them to this treadmill exercise
appears to help with their running performance (Payne et al, 2006)(Radley-Crabb et al,
submitted) in addition, the motivation for exercise may be influenced by various
environmental and behavioural factors.
The advantages and limitations of both
voluntary and treadmill exercise are summarised in Table 1.3.2.
Voluntary wheel exercise
Controlled treadmill exercise
Advantages:
Advantages:
Mdx mice can run when they choose (usually at
Reduce variation between mice - pre-determined
night)
speed, distance & protocol
Minimal supervision required by researcher
Sacrifice at a known time after exercise
Stop-start running pattern favoured by mdx mice
Up-hill / down-hill running if required
Non-invasive and non-stressful for the mice.
Cage mdx mice in groups when not running
Multiple inexpensive cage set-ups
Limitations:
Limitations:
Variation in speed, distance & duration between
Time consuming for researcher
mice
Mdx mice forced to run during inactive phase
Heavily influenced by behaviour
Can be stressful for the mouse
Wheels can ‘block-up’ overnight
Initially expensive
Must cage mdx mice individually
Table 1.3.2 Summary of the main advantages and limitations of two different exercise models for the
mdx mouse: voluntary wheel exercise and controlled treadmill exercise.
There are numerous mdx treadmill protocols documented in the literature with many
differences in speed, duration, regime and surface incline/decline. Horizontal running
is most common (Burdi et al, 2009; De Luca et al, 2005; De Luca et al, 2008; De Luca et
al, 2003; Granchelli et al, 2000; Grounds et al, 2008b), however some researchers use
up-hill (Ahmad et al, 2009; Okano et al, 2005), or down-hill (eccentric) treadmill
running (Bizario et al, 2009; Brussee et al, 1997; Vilquin et al, 1998). Eccentric muscle
contractions (muscle contraction while muscle is extended) are the most damaging for
21
_____________________________________________________Chapter 1 - Introduction
dystrophic (and normal) muscle. Experimental lengthening contractions induced via
nervous stimulation while the muscle is extended in vivo, avoids behavioural
complications and produces reproducible contraction of the entire musculo-tendinous
unit, allowing for more experimental control (Hamer et al, 2002; Piers et al, 2010;
Ridgley et al, 2008; Weller et al, 1990), although these specialised exercise regimes
using a dynanometer require expensive and specialised laboratory equipment (See
Chapter 8 of this Thesis).
1.3.3 Development of Standard Operating Procedures:
Recently, the important issue of world-wide standard operating procedures for preclinical studies in the mdx mouse has been heavily discussed (Grounds et al, 2008b;
Nagaraju and Willmann, 2009). Researchers have begun establishing a set of
recommended standard operating procedures and endpoints for consistent and
comparable assessment of pre-clinical interventions in the mdx mouse (Grounds et al,
2008b; Guerron et al, 2010; Spurney et al, 2010; Spurney et al, 2009)(Radley-Crabb et
al, submitted). This was catalysed by ‘The first Brazilian International Workshop on
preclinical tests for drug therapies for Muscular Dystrophy’ a 3 day workshop help in
Brazil (Ribeirao Preto, Sao Paulo 2006) (Bizario and Costa, 2007) and an associated
large review manuscript (Grounds et al, 2008b). There are also many standard
operating procedures listed on the TREAT-NMD website http://www.treatnmd.eu/research/preclinical/SOPs. The standard operating procedures listed on the
TREAT-NMD website have been employed (chapters 5 & 6) and contributed to (chapter
2) in this thesis.
1.4. Myofibre Necrosis in dystrophic muscle:
Myofibre necrosis is the fundamental destructive process in several forms of
inflammatory myopathies and muscular dystrophy– especially DMD. In contrast to
healthy skeletal muscle, fragile dystrophic muscles undergo repeated cycles of
sarcolemmal damage, myofibre necrosis and subsequent regeneration. The exact
mechanism by which the absence of dystrophin leads to myofibre necrosis is unknown
however inflammation, oxidative stress, metabolic abnormality, mislocalisation of
22
_____________________________________________________Chapter 1 - Introduction
nNOS and increased intracellular calcium are all heavily implicated in the process
[Reviewed in (Allen et al, 2010; Evans et al, 2009a; Evans et al, 2009b; Grounds et al,
2008a; Leighton, 2003; Radley et al, 2007; Tidball and Wehling-Henricks, 2004b; Tidball
and Wehling-Henricks, 2005; Tidball and Wehling-Henricks, 2007; Wallace and
McNally, 2009)].
The absence of dystrophin does not affect the activation or proliferation of satellite
cells as dystrophin is not normally expressed in satellite cells or myoblasts. Satellite
cells are hyperactivated in dystrophic muscle due to the need for myogenesis in
response to necrosis. Initially satellite cells (myogenesis) can repair the damaged
myofibres but eventually this process fails and dystrophic muscle is replaced with fatty
and/or fibrous connective tissue [Reviewed in (Anderson and Wozniak, 2004; Karpati
and Molnar, 2008)]. There are conflicting data surrounding satellite cell number and
differentiation status between in vitro and in vivo studies and disease states; however,
a recent study examining satellite cell numbers in muscle biopsies from DMD patients
aged 2-7 years, reports an increased number of quiescent satellite cells (Pax-7 positive
nuclei, located beneath the basement membrane on the surface of the myofibre)
(Kottlors and Kirschner, 2010). It has also been proposed that growing dystrophic
skeletal muscle is more susceptible to myofibre necrosis than adult muscle and that
the cellular and molecular events contributing to acute myofibre necrosis (e.g. 3 week
old young mice or exercised adult mdx mice) compared with chronic necrosis (e.g. low
persistent necrosis in adult mdx mice) are quite different (Grounds, 2008).
1.4.1 Inflammation and dystrophic muscle:
A persistent inflammatory state is observed in dystrophic skeletal muscle, including
(but not limited to) an increased presence of inflammatory (and immune) cells (e.g.
neutrophils, macrophages, T-lymphocytes and mast cells), elevated levels of various
pro-inflammatory cytokines (e.g. TNF and TGFβ) and increased oxidative stress (Figure
1.4.1) [Reviewed (Evans et al, 2009a; Evans et al, 2009b; Grounds et al, 2008a; Spencer
and Tidball, 2001; Tidball and Wehling-Henricks, 2005; Tidball and Wehling-Henricks,
2007)]. It is hypothesised that activation of the inflammatory cascade, in response to
sarcolemmal damage, exacerbates myofibre damage, leading to myofibre necrosis
23
_____________________________________________________Chapter 1 - Introduction
rather than repair of small sarcolemmal tears (Evans et al, 2009b; Grounds et al,
2008a; Grounds and Torrisi, 2004; Radley et al, 2008; Radley and Grounds, 2006;
Spencer and Tidball, 2001).
Figure 1.4.1. Sequence of the inflammatory response in dystrophic skeletal muscle. Resident mast cells
act early in the inflammatory cascade and rapidly degranulate in response to damage to release TNF and
many other pro-inflammatory mediators. Figure taken from Radley et al (2006) (Radley and Grounds,
2006).
After muscle contractions which damage (tear) the dystrophic sarcolemma, there is an
aggressive invasion of inflammatory (and immune) cells into dystrophic muscle, which
can exacerbate myofibre damage, through the release of pro-inflammatory cytokines,
increased oxidative stress and proteolysis (Figure 1.4.1) (Radley et al, 2008; Spencer et
al, 2001; Tidball, 2005). Resident and rapidly infiltrating neutrophils can also damage
dystrophic muscle via the production of hypochlorous acid (HOCl) and via the secretion
of superoxide (SO•) which is then converted to the highly damaging hydroxyl radical
(OH•) (Tidball, 2005).
24
_____________________________________________________Chapter 1 - Introduction
Strong evidence that inflammatory cells can contribute to necrosis of healthy muscle
cells comes from studies investigating the role of neutrophils, macrophages and
oxidative damage in vitro (McLoughlin et al, 2003; Nguyen and Tidball, 2003b; Pizza et
al, 2001; Toumi et al, 2006) and in vivo (Cheung and Tidball, 2003; Nguyen and Tidball,
2003a; Pizza et al, 2002). It has similarly been proposed that an excessive inflammatory
response can directly damage myofibres in myopathic conditions such as dystrophies
or myositis (Porter et al, 2002; Tidball and Wehling-Henricks, 2005) and recent data
increasingly implicate inflammation and specifically tumour necrosis factor (TNF) in
myofibre necrosis [Reviewed in (Evans et al, 2009b; Grounds et al, 2008a)].
TNF, previously referred to as TNFα
(Clark, 2007), is a major pro-inflammatory
cytokine that is expressed by a wide range of inflammatory cells and by myoblasts,
myotubes and damaged skeletal muscle (Collins and Grounds, 2001; Kuru et al, 2003).
TNF is also produced by adipose tissue (Fruhbeck, 2004; Weisberg et al, 2003) that is
often pronounced within DMD patients as the disease progresses. In response to
myofibre damage, TNF is rapidly released from resident mast cells and also by
neutrophils that accumulate quickly at sites of tissue damage (Radley and Grounds,
2006; Tidball, 2005). TNF is a potent chemokine that attracts further inflammatory cells
to the injured site. The chemotactic role of TNF was demonstrated in normal mouse
muscle, where administration of TNF resulted in the accumulation of neutrophils and
macrophages in the absence of any sarcolemma damage (Peterson et al, 2006).
Elevation of TNF protein locally in dystrophic muscles is supported by
immunohistological studies that show increased staining for TNF associated with
necrotic areas of dystrophic muscles of the mdx mouse (Collins, 1999) and biopsies
from DMD patients (Kuru et al, 2003) and also with Western blotting analysis using
anti-TNF antibodies in mdx muscle extracts (Messina et al, 2006). Few studies have
accurately quantitated TNF in dystrophic muscles or blood; due mainly to problems
with sensitivity thresholds of such immunodetection in muscle and blood. Numerous
examples from the literature which demonstrate an increase in TNF in dystrophic
muscle are summarised in Table 1.4.
25
_____________________________________________________Chapter 1 - Introduction
Authors
Saito et al.
Chahbouni
et al
Porreca et
al.
Pan et al.
Bizario et al.
Tews et al.
Kuru et al.
Hnia et al.
Grounds et
al.
Messina et
al.
Messina et
al.
Measurements of TNF in mdx mice and DMD patients.
1) TNF Protein
Reference
Technique
TNF level
Clin Chem 2000; Immuno-PCR assay –
Serum TNF 27.8ng/L in
46: 1703-4.
serum (Novel
DMD patients vs
technique).
0.027ng/L in healthy
controls.
+
J. Pineal Res
Linco-Plex plate - blood Plasma TNF 4.43ng/L ( /2010; 48: 282plasma.
0.61) in DMD patients vs
+
289.
2.9ng/L ( /- 0.29) in
healthy controls.
Thromb
ELISA – serum
Serum TNF 54ng/L in
Haemost 1999;
(Amersham Int UK).
DMD/BMD patients vs
81: 534-61.
25ng/L in healthy controls.
+
Mol Cells 2008;
ELISA – serum
Serum TNF 120ng/L ( /- 40)
25: 531-37.
(JingMed Co PR China). in 28 day old mdx mice vs
+
10ng/L ( /- 10) in C57Bl/10
controls.
J Neuroimmuno Cytometric bead array
Serum TNF 25ng/L (+/- 5)
2009; 212: 93and flow cytometry
in chronic downhill
101.
(BD biosciences Corp
exercised 10 wk old mdx
CA).
mice.
J Neuropathol
Immunohistochemistry TNF detected in 5/8 DMD
Exp Neurol
– muscle biopsy
patient limb muscle
1996; 55: 342(Genzyme USA).
biopsies vs undetected in
47.
healthy controls.
Acta
Immunohistochemistry TNF positive muscle fibres
+
neuropathol
– muscle biopsy
15.8% /- 5.1% in DMD
2003; 105: 217- (Genzyme USA).
patient biopsys vs 0% TNF
24.
positive fibres in healthy
controls.
Am J pathol
Immunocytochemistry TNF detected in all mdx
2008; 172:
and western blot –
diaphragm sections and in
1509-79.
mdx diaphragm
all mdx protein samples
(Abcam UK).
used in western blots.
Clin exp pharma Immunohistochemistry TNF detected in mdx
physiol 2008;
– mdx and C57Bl/10
muscle section and
35: 846-51.
limb muscle sample.
undetected in C57Bl/10
control section.
Am J pathol
Western blot – mdx
TNF in mdx muscle approx
2006; 168: 918- and C57Bl/10
6x higher than in control
26.
quadriceps muscle
muscle (arbitrary values).
(Chemicon USA).
Exp neurol
Western blot – mdx
TNF in mdx muscle approx
2006; 196: 234- and C57Bl/10
4x higher than in control
41.
quadriceps muscle
muscle (arbitrary values).
(Chemicon USA).
RadleyCrabb et al.
2009
unpublished
Authors
Lundberg et
al.
Reference
J neuroimmunol
1995; 63: 9-16.
Huang et al.
FASEB 2009;
March epub.
Elisa – serum
(Invitrogen Aus).
TNF in serum of unex/ex
C57BL/10 and mdx mice –
undetectable (<15.6ng/L).
2) TNF Gene expression
Technique
TNF level
Reverse transcription
TNF mRNA detected in 3/5
Polymerase chain
DMD patients vs 2/5
reaction – muscle
healthy controls.
biopsys.
Reverse transcription
TNF mRNA 2.5x higher in
Polymerase Chain
mdx diaphragm muscle in
Change
1000x increase.
2x increase
2x increase.
12x increase.
No comparison
to C57Bl/10
controls.
Increased
detection.
Increased
detection.
No comparison
to C57Bl/10
controls.
Increased
detection.
6x increase.
4x increase.
-
Change
Slight increase in
detection.
2.5x increase.
26
_____________________________________________________Chapter 1 - Introduction
Bizario et al.
J Neuroimmuno
2009; 212: 93101.
RadleyCrabb et al
2010
unpublished
RadleyCrabb et al
2010 submitted
Kuru et al.
Reaction – diaphragm
muscle samples.
Reverse transcription
Polymerase Chain
Reaction –
gastrocnemius muscle
samples.
Reverse transcription
Polymerase chain
reaction –
gastrocnemius muscle
samples.
Reverse transcription
Polymerase chain
reaction –quadriceps
muscle samples.
In-situ hybridisation –
frozen muscle biopsys
(Genzyme USA).
comparison to C57Bl/10
control muscle.
TNF mRNA detected in
chronic downhill exercised
10 wk old mdx mice.
No comparison
to C57Bl/10
controls.
TNF mRNA 4x higher in
mdx gastrocnemius
muscle in comparison to
C57Bl/10 control muscle.
4x increase.
TNF mRNA 10 higher in
mdx quadriceps muscle in
comparison to C57Bl/10
control muscle.
TNF mRNA detected in
muscle fibres of DMD
patient biopsys.
5x increase.
Acta
No comparison
neuropathol
to controls.
2003; 105: 21724.
Table 1.4. A comprehensive list of measurements of TNF in mdx mice and DMD patients. Both TNF
protein and gene expression are listed.
Elevated TNF may exacerbate muscle damage through several pathways (Clark et al,
2004). One of the contributors to TNF induced muscle necrosis could be the
inflammatory transcription factor nuclear factor-kappa beta (NFκB), since NFκB is
activated in limb muscles (Acharyya et al, 2007) and the diaphragm (Acharyya et al,
2007; Kumar and Boriek, 2003) of mdx mice and also in muscle samples from DMD
patients (Monici et al, 2003). Blockade of NFκB by pyrrolidine dithiocarbamate reduces
skeletal muscle degeneration in mdx mice (Messina et al, 2006) and it has recently
been shown that heterozygous deletion of the p65 subunit of NFκB is sufficient to
decrease muscle necrosis in mdx mice (Acharyya et al, 2007). Dysregulation of
signalling through activation of the NFκB pathway appears to be particularly important
in many diseases (associated with inflammation) and chronically activated NFκB is a
feature of DMD and mdx muscles (Peterson and Guttridge, 2008).
1.4.2 Anti-inflammatory therapies in dystrophic muscle:
A wide range of immunosuppressant, anti-inflammatory and specific anti-cytokine
drugs have been tested in both mdx and DMD patients, with benefits for various
indicators of muscle damage [Reviewed (Bogdanovich, 2004; Evans et al, 2009b;
Manzur and Muntoni, 2009; Radley et al, 2007; Spurney et al, 2009)]. Treatments that
27
_____________________________________________________Chapter 1 - Introduction
include (but are not limited to); blockade of TNF (Grounds and Torrisi, 2004; Hodgetts
et al, 2006; Pierno et al, 2007; Radley et al, 2008), cromolyn prevention of mast cell
degranulation (Granchelli et al, 1996; Radley and Grounds, 2006), cyclosporine
treatment (De Luca et al, 2005), and depletion or reduction of CD4+ or CD8+ cells (T
cells) (Spencer et al, 2001), neutrophils (Hodgetts et al, 2006) and macrophages
(Wehling et al, 2001), reduce the severity of dystropathology in mdx mice. Other antiinflammatory therapies, less specific and less commonly used, such as Imatinib and
Melatonin have also demonstrated beneficial effects in young mdx mice (Bizario et al,
2009; Chahbouni et al, 2010; Huang et al, 2009).
Two drugs have been used to block TNF activity in the mdx mouse; Remicade® (an
antibody to human TNF, also known as Infliximab) (Grounds and Torrisi, 2004); and
Enbrel® (soluble receptor to TNF, also known as Etanercept) (Hodgetts et al, 2006;
Pierno et al, 2007) are in wide clinical use to treat inflammatory disorders such as
arthritis and Crohn’s disease. The high specificity of these anti-cytokine drugs,
combined with their clinical success in other diseases and relatively few side effects
suggests that they may be attractive alternatives to the existing use of corticosteroids
to treat DMD. In the mdx mouse, long-term studies have further demonstrated that
the mouse-specific antibody to TNF (cV1q) has equal efficacy to Remicade® and
Enbrel® (Radley et al, 2008) (See chapter 3 of this thesis). It is noted that cV1q
blockade of TNF has no effect on the low levels of chronic myofibre damage in unexercised dystrophic muscle, in striking contrast to the marked protective effect on
exercise-induced acute myofibre necrosis, raising the possibility of different roles for
TNF (and other molecules) in these two different situations of myopathology (Radley
et al, 2008). These results strongly support a key role for TNF in both inflammation and
necrosis in dystrophic muscle.
While systemic depletion of inflammatory cells or cytokines is not clinically viable,
studies in gene ‘knock-out’ mice provide a wealth of knowledge regarding
inflammatory signalling in dystrophic muscle and the potential compensatory upregulation of other cytokines (Rafael et al, 1997; Spencer et al, 2000). It was
hypothesised that generation of dystrophin and TNF double knock-out mice (TNF /mdx), would result in improved dystropathology. However, serum CK level and whole
28
_____________________________________________________Chapter 1 - Introduction
body strength was unchanged in these mice, the extent of dystropathology was
inconsistent across muscle groups and body weight of the mice was significantly
reduced. These results suggest that the effects of complete TNF knock-out are
complex, unpredictable, and may result in the up-regulation of numerous other
inflammatory cytokines to compensate for the absence of TNF (Spencer et al, 2000).
Phenotypic examination of these dystrophin and TNF gene ‘knock-out’ mice (TNF /mdx) was carried out on sedentary 8 week old mice, and different effects might have
been seen in exercised mice.
1.4.3 Reactive oxygen species and oxidative stress:
The term reactive oxygen species (ROS) is used to describe oxygen radicals, including
superoxide (O2•-) and hydroxyl (OH•) and the non radical derivatives of oxygen,
including hydrogen peroxide (H2O2) and hypochlorous acid (HOCl). ROS are generated
by a wide range of biological functions in the human body. The most reactive of the
ROS, the hydroxyl radical (OH•) is formed from hydrogen peroxide (H2O2) in the
presence of metal ions (iron) via Fenton chemistry (Figure 1.4.2). Oxidative stress
occurs when ROS generation is not counteracted by antioxidants; substances that
delay prevent or remove oxidative damage to a target molecule Three of the main
anti-oxidant systems are 1) superoxide dismutase enzymes (SODs) which remove
superoxide O2•- by accelerating its conversion into hydrogen peroxide H2O2, 2) catalase
enzymes which convert hydrogen peroxide H2O2 into water H2O and oxygen O2 and 3)
the glutathione system which removes hydrogen peroxide H2O2 by using it to oxidize
reduced glutathione GSH to oxidized glutathione GSSG. An excess production of ROS
and/or an insufficient anti-oxidant defence within a cell can lead to a situation of
oxidative stress [Reviewed in (Arthur et al, 2008; Halliwell and Gutteridge, 2007)]. ROS
have a diverse range of actions in cells, ROS can cause irreversible oxidation (a gain of
oxygen or loss of electrons) of cellular components, cross linking or fragmentation of
protein, damage to DNA and permanent damage to membrane phospholipids (lipid
peroxidation) (Figure 1.4.2) (Arthur et al, 2008; Berlett and Stadtman, 1997). These
processes can lead to irreversible cell damage and death by both apoptosis and
necrosis (Rando, 2002; Rando et al, 1998). ROS can also affect cell signalling pathways
through the reversible modification of transcription factors and signal transduction
29
_____________________________________________________Chapter 1 - Introduction
proteins, mediated by the reduction/oxidation (redox) state of protein thiols
[Reviewed in (Arthur et al, 2008; Lui et al, 2010)].
Figure 1.4.2. Formation and properties of some ROS molecules . As shown, ROS are generated by the
sequential addition of electrons. Figure taken from Arthur et al (2008) (Arthur et al, 2008).
1.4.4 Dystrophic muscle and oxidative stress:
After contraction (that damages vulnerable dystrophic myofibres) there is an
aggressive invasion of inflammatory cells (Hodgetts et al, 2006; Radley et al, 2008;
Radley and Grounds, 2006; Spencer et al, 2001; Spencer and Tidball, 2001), which can
exacerbate myofibre damage, through cellular processes such as phagocytosis and
proteolysis (Tidball, 2005; Tidball and Wehling-Henricks, 2005). Neutrophils can
damage dystrophic muscle via the production of hypochlorous acid (HOCl) and the
secretion of superoxide (SO•) which is then converted to hydroxyl radical (OH•) and
this increase in ROS leads to further damage of the muscle tissue (Tidball, 2005; Tidball
and Wehling-Henricks, 2005).
While it has been established that oxidative stress is increased in dystrophic muscle
(shown by lipid peroxidation, glutathione status and Dihydroethidium staining - a
fluorescent probe for superoxide) (Austin et al, 1992; Burdi et al, 2009; Disatnik et al,
2000; Dudley et al, 2006; Whitehead et al, 2008) and that dystrophic myotubes and
myofibres are much more susceptible to oxidative damage (Ragusa et al, 1997; Rando
et al, 1998), the exact role of oxidative stress in necrosis of dystrophic myofibres is
uncertain. Some research suggests that ROS generation in dystrophic muscle is a
primary occurrence, due to deregulation of nitric oxide (NO) (Grisotto et al, 2000;
Thomas et al, 1998; Wehling et al, 2001), but others have suggested it is secondary to
30
_____________________________________________________Chapter 1 - Introduction
excessive calcium influx and inflammation by contraction induced damage (Whitehead
et al, 2008; Yeung et al, 2005).
In normal healthy muscle, neuronal nitric oxide synthase (nNOS) is directly bound to
syntrophins in the dystroglycan complex (DGC) but, due to the absence of dystrophin,
in dystrophic muscle nNOS is displaced from the sarcolemma into the cytoplasm.
nNOS is an important member of the DGC complex as it synthesises nitric oxide (NO), a
molecule that reacts with free radicals and regulates vascular tone. During muscle
contraction, the production of NO allows blood vessels to vasodilate which prevents
ischemia and subsequent damaging reperfusion. In the absence of dystophin, NO
production is significantly reduced and may lead to free radical mediated damage
[Reviewed in (Brenman et al, 1995; Wallace and McNally, 2009)]. Lack of NO is
proposed as a mechanism for damage in dystrophic muscle, and over-expression of
nNOS in transgenic mdx mice decreases myofibre damage, CK levels and macrophage
invasion (Wehling et al, 2001).
ROS production in dystrophic muscle could also be a secondary consequence of
contraction induced injury, facilitated by excessive intracellular calcium and
inflammatory cell cascade. Contraction of dystrophic muscle leads to excessive
intracellular calcium via either increased sarcolemma tears or altered calcium channel
function (Turner et al, 1988; Wallace and McNally, 2009; Yeung et al, 2005). This
calcium imbalance can lead to excessive activation of proteases such as calpain
(Whitehead et al, 2008; Whitehead et al, 2006a) which promote proteolysis (protein
degradation). An overload of mitochondrial calcium stimulates the generation of ROS
(Brookes et al, 2004).
Oxidative stress may also contribute to the pathology of dystrophies by disrupting cell
signalling. Cellular protein thiols (sulfhydryl groups) respond to an increase in cellular
ROS and oxidation leads to a reversible formation of disulphide bonds. These
modifications can directly affect the activity of signal transduction proteins or
transcription factors as well as cause indirect changes in signal transduction through
modifications of ion transport proteins (Arthur et al, 2008; Kourie, 1998; Suzuki and
Ford, 1999). Nuclear factor kappa B (NFκB) is an important redox regulated inducible
31
_____________________________________________________Chapter 1 - Introduction
transcription factor activated by oxidative stress (Macaione et al, 2007). It regulates
the expression of many genes, including those involved in the inflammatory and stress
response. In particular, activation of the NFκB pathway increases pro-inflammatory
cytokines such as TNF, and levels of both NFκB and TNF (Table 1.4) are elevated in mdx
mice (Acharyya et al, 2007; Kumar and Boriek, 2003; Peterson and Guttridge, 2008).
1.4.5 Antioxidant therapies in dystrophic muscle:
An alternative (or perhaps complementary) approach to treating DMD is to decrease
oxidative stress. N-acetylcysteine (NAC) is an anti-oxidant which decreases oxidative
stress by scavenging ROS (hypochlorus acid, hydroxyl radicals and hydrogen peroxide)
(Aruoma et al, 1989). NAC also has many effects on cell signalling and is a precursor to
the cellular antioxidant glutathione which is preferentially oxidized by ROS thus
protecting vital macromolecules such as DNA, proteins and lipids (Zafarullah et al,
2003). When administered to mdx mice, NAC reduced cellular ROS (shown by DHE
staining), decreased nuclear protein expression of NFκB, and reduced muscle damage
(histological quantitation of centrally nucleated myofibres) (Whitehead et al, 2008).
The in vivo effects of NAC in exercised mdx mice were not investigated by Whitehead
et al (2006); thus further analysis of myofibre necrosis and more specific ROS
measurements were carried out in Chapter 5 of this Thesis.
Numerous in vivo studies have shown green tea (specifically the major polyphenol (-)epigallocatechin gallate –EGCG) to have anti-oxidant properties in vivo and beneficial
effects on the extent of dystropathology in mdx mice (Buetler et al, 2002; Call et al,
2008; Dorchies et al, 2006; Evans et al, 2010; Nakae et al, 2008), possibly via downregulation of NFκB (Evans et al, 2010). Call et al (2008) reported additional benefits of
EGCG treatment when administered in combination with voluntary exercise in young
mdx mice (21-42 days of age), possibly due to the beneficial effects of long-term
voluntary exercise (Call et al, 2008).
NAC and EGCG function as antioxidants by scavenging radicals and reducing oxidative
stress, but it is feasible that a more powerful way to prevent oxidative stress is to
prevent ROS from forming. As mentioned previously, formation of the highly reactive
32
_____________________________________________________Chapter 1 - Introduction
hydroxyl radical requires an iron catalyst. Free iron, or Fe (II) is capable of catalysing
the production of the hydroxyl radical. In this context, it is relevant to note that when
skeletal muscle of rats is overloaded with iron, there is increased oxidative stress,
decreased muscle strength and increased muscle atrophy (Reardon and Allen, 2009). In
contrast, dystrophic mdx mice fed a low iron diet show a decrease in macrophageinvaded necrotic myofibres (Bornman et al, 1998).
In addition to excess (dysregulated) inflammation and oxidative stress, it is also
proposed that the absence of dystrophin in skeletal muscle leads to many metabolic
abnormalities, which may also be involved in the process of myofibre necrosis.
1.5 Skeletal muscle metabolism:
Metabolism refers to the sum of all chemical processes which occur in a living
organism in order to maintain life. Metabolic processes enable living organisms to
complete vital functions such as respiration, reproduction, growth, maintenance, and
repair. Metabolism is divided into two types of processes – catabolic and anabolic.
Catabolism is the set of metabolic processes that break down large molecules into
components that can be used in anabolic processes e.g. digesting dietary protein into
amino acids. Anabolism, on the other hand, is the set of constructive metabolic
processes where the energy released by catabolism is used to synthesize complex
molecules. Some of the main metabolic processes carried out by skeletal muscle are
the synthesis and degradation of proteins, fatty acid catabolism (βoxidation of fatty
acids produces Acetyl-CoA which is used in the citric acid cycle), mobilization of
glycogen stores for energy production and glycogen synthesis from glucose for energy
storage. Hydrolysis of adenosine triphosphate (ATP) is the main source of energy for
many enzymatic reactions that occur in skeletal myofibres, ATP is supplied by
mitochondria located in the myofibre cytoplasm and since contraction is an energy
demanding process myofibres are rich in mitochondria. Depending on the level of
skeletal muscle activity, ATP is replenished from phosphocreatine stores, through the
process of glycolysis or via oxidative metabolism [Reviewed in (Bettelheim et al, 2001;
MacIntosh et al, 2006)].
33
_____________________________________________________Chapter 1 - Introduction
1.5.1 Protein turnover in skeletal muscle:
Skeletal muscle protein mass is tightly regulated by (subtle) changes in the relative
rates of protein synthesis and degradation. The whole-body protein synthesis rate
represents the compilation of protein synthesis rates of all tissues and organs in the
body and skeletal muscle represents the largest protein mass in the body. Growth,
aging, nutrition and disease can all affect the protein synthesis rates of tissues and
organs. Protein turnover is the net balance between protein synthesis and protein
degradation. More synthesis than breakdown indicates an anabolic state that builds
lean tissues, whereas more breakdown than synthesis indicates a catabolic state that
burns lean tissues. Proteins are constantly synthesized and degraded in cells.
Depending on the tertiary structure of the protein (stability), turnover rate can vary
from minutes to weeks; rapid protein turnover allows cells and tissues to respond
quickly to constantly changing conditions. Protein synthesis involves the process of
translation of mRNA into an amino acid sequence to form a peptide, which occurs on
ribosomes in the cytoplasm. Protein breakdown occurs generally in the lysosomes or
proteasomes. The most commonly employed method for measuring actual protein
synthesis is the use of isotopic tracers to measure the rate of incorporation of a
labeled amino acid into tissue protein over-time [Reviewed in (Bettelheim et al, 2001;
Davis et al, 1999a)].
Such issues of protein turnover are also influenced by nutrition, especially specific
amino acids derived from protein ingestion, by available energy provided by different
foods and influenced by the balance of energy demands within the body. It has been
proposed (Leighton, 2003) that dystrophic boys have an altered metabolic rate and
therefore this was investigated to explore the potential basis for meaningful dietary
interventions or nutritional supplements for DMD boys.
1.5.2 Metabolic alteration in dystrophic muscle:
Boys with DMD can move between the spectra of over-nutrition to under-nutrition
within their shortened lifespan. Young DMD patients are often over-weight or obese
34
_____________________________________________________Chapter 1 - Introduction
(due to a combination of inactivity, steroid therapies and proposed metabolism
defects), yet patients are also often underweight and experience malnutrition in later
life [Reviewed in (Davidson and Truby, 2009; Leighton, 2003)]. Despite these
observations, there is limited literature on the nutritional and energy requirements for
DMD patients. DMD is considered to be a hyper-metabolic condition [Reviewed in
(Leighton, 2003)] with the basal metabolic rate (kcal/kg bw/24hr), dietary protein
requirements (g protein/kg bw/24hr) and urinary excretion of 3-methylhistidine (an
indicator of muscle protein breakdown) all higher in DMD patients (aged 11- 29 years)
in comparison to healthy controls (Davidson and Truby, 2009; Leighton, 2003; Okada et
al, 1992). Resting energy expenditure when adjusted to free fat mass (lean muscle
tissue) is also higher in DMD patients (6-13 years) (Hankard et al, 1996; Zanardi et al,
2003).
Studies in mdx mice also indicate that dystrophin defects lead to altered skeletal
muscle metabolism and an impaired energy status. Repeated cycles of myofibre
necrosis and regeneration, increased demand on the sarcoplasmic reticulum to
regulate intracellular calcium, defective mitochondrial function, increases in both
protein synthesis and protein degradation rates, and disruption in nNOS signalling may
all contribute to an altered metabolic state (Dupont-Versteegden et al, 1994a; Griggs
and Rennie, 1983; Han et al, 2006; Kuznetsov et al, 1998; Landisch et al, 2008;
MacLennan and Edwards, 1990; Passaquin et al, 2002; Whitehead et al, 2006b; Zanardi
et al, 2003). In support of an altered metabolism in dystrophic muscle, 48 hours fasting
significantly increased myofibre necrosis in muscles from the hind limb and lumbar
region of 6 month old mdx mice (yet no change in muscle morphology of control
C57BL/10 mice), suggesting a strong dependence on caloric intake to maintain
dystrophic muscle structure (Helliwell et al, 1996).
Changes in protein metabolism have been reported in various mdx muscles both in
vivo and ex vivo (Helliwell et al, 1996; MacLennan and Edwards, 1990; MacLennan et
al, 1991a; MacLennan et al, 1991b; Turner et al, 1988). The rates of both protein
synthesis and protein degradation are elevated in the gastrocnemius muscle of mdx
35
_____________________________________________________Chapter 1 - Introduction
mice (various ages, sex unspecified) and it is suggested that this process is mediated by
an increase in ribosome concentration in mdx muscles (MacLennan and Edwards,
1990). Interestingly, in mdx liver where there is no dystropathology, the rate of protein
synthesis is similar to control C57BL/10 mice, indicating that altered protein
metabolism in mdx mice is restricted to skeletal muscles (MacLennan and Edwards,
1990). The extent to which this reflects the increased protein demand of growing
myotubes and myofibres compared with mature myofibres is unclear in dystrophic
muscle.
Adult mdx mice maintain skeletal muscle mass and experience significant skeletal
muscle hypertrophy (Anderson et al, 1987; Coulton et al, 1988a; Mokhtarian et al,
1996; Peter and Crosbie, 2006; Shavlakadze et al, 2010; Spurney et al, 2009), indicating
that protein synthesis must outweigh protein degradation in adult mdx mice. Despite
adult mdx mice experiencing significant muscle hypertrophy, dystrophic muscles are
still ‘weaker’ than those from control C57Bl/10 mice due to the absence of dystrophin
protein (Ridgley et al 2009). Skeletal muscle hypertrophy in adult mdx mice is also a
feature of young DMD boys (~ 5 years of age) where hypertrophic calf muscles are a
diagnostic feature (Bushby et al, 2010a; Hoffman, 2001). This is in striking contrast to
older (6-18 years), wheel chair bound, DMD patients that lose lean muscle mass at an
approximate rate of 4% per year (Griffiths and Edwards, 1988). Skeletal muscle
hypertrophy in adult mdx mice may be linked to an increase in AKT signalling (Peter
and Crosbie, 2006), since AKT is a key downstream molecule of IGF-1 signalling that
results in increased protein synthesis and hypertrophy [Reviewed in (Shavlakadze and
Grounds, 2006)]. There are also reports of increased circulating growth hormone in 810 month old female mdx (Anderson et al, 1994) and increased circulating IGF-1 levels
in 8-10 week old (sex unspecified) mdx mice (De Luca et al, 1999).
It was hypothesised by Dupont-Vesteegden et al (1994) that continuous cycles of
myofibre necrosis and subsequent regeneration, increased levels of intracellular
calcium and elevated protein synthesis in dystrophic mdx muscles could lead to an
overall increase in whole body metabolic rate; however they reported no change in the
metabolic rate (kcal/kg bodyweight0.75/24hr) of one year old adult mdx mice (mixed
sex). In contrast, there was a decrease in the whole body metabolic rate (kcal/kg
36
_____________________________________________________Chapter 1 - Introduction
bodyweight0.75/24hr) of 4-6 week old (mixed sex) mdx mice, and it was proposed that
this was a consequence of reduced physical activity and reduced food consumption (g
food /g weight) (Dupont-Versteegden et al, 1994a). Mokhtarian et al (1996) also
reported no difference in total energy expenditure, basal energy expenditure and
spontaneous activity between control C57Bl/10 and mdx mice (sex unspecified) at 6-12
months of age (Mokhtarian et al, 1996). Clearly, mouse age, extent of dystropathology
and the presence of regenerating (growing) muscle, may have a major impact on
metabolic demand and energy usage.
1.5.3 Nutritional therapies for dystrophic muscle:
Dystrophin deficiency does not adversely affect muscle growth or function, in fact
adult mdx mice experience muscle hypertrophy (Anderson et al, 1987; Coulton et al,
1988a; Mokhtarian et al, 1996; Peter and Crosbie, 2006; Shavlakadze et al, 2010;
Spurney et al, 2009), however the absence of dystrophin does eventually lead to
muscle wasting in both ‘old’ mdx mice and DMD patients (Bushby et al, 2010a; Emery,
2002; Sinnreich, 2010). Dietary supplementations have shown beneficial effects in
both mdx mice and DMD patients [Reviewed in (Bogdanovich, 2004; Davidson and
Truby, 2009; Leighton, 2003; Pearlman and Fielding, 2006; Radley et al, 2007)], in
addition to this some recently completed clinical trials have examined the role of
creatine (organic acid) and glutamine (amino acid) supplementation in DMD patients
with some beneficial effects (Escolar et al, 2005; Mok et al, 2006; Tarnopolsky et al,
2004).
Creatine is a major intermediate of cellular energy transfer; and when fed to newborn
mdx mice (10% w/w in chow) strongly reduced the onset of muscle necrosis in the fasttwitch EDL muscle and also improved mitochondrial respiration capacity (Passaquin et
al, 2002). In addition, creatine, taurine (organic acid) and glutamine treatments have
shown beneficial effects in treadmill exercised adult mdx mice (De Luca et al, 2003;
Granchelli et al, 2000). A combined nutritional therapy (creatine monohydrate –
organic acid, conjugated linoleic acid – essential fatty acid, α-lipoic acid –
mitochondrial coenzyme and antioxidant, and β-hydroxy-β-methylbutyrate –leucine
metabolite) administered, in addition to prednisolone, for 8 weeks increased muscle
37
_____________________________________________________Chapter 1 - Introduction
strength and reduced the extent of dystropathology in 12 week old treadmill exercised
mdx mice (Payne et al, 2006). A combination of taurine (1g/kg bw/day - orally) and
prednisolone (1mg/kg bw/day - i.p. injection) treatment of treadmill exercised mdx
mice (4-8 weeks of age) also markedly improved forelimb grip strength, compared to
either taurine or prenisolone treatment alone (Cozzoli et al, 2010). In addition, a high
protein diet (50%) improved muscle morphology in dystrophic laminin deficient
(129ReJ dy/dy) mice and caused a shift to a more ‘normal’ protein metabolism
(Zdanowicz et al, 1995).
A preliminary study to test the effects of a high fat diet on the dystrophic mdx heart
(Personal communication - Andrew Hoey 2005, Queensland Australia, data
unpublished), reported a striking difference in bodyweight of dystrophic mdx and
control mice fed a high fat diet (~15%) for 9 weeks (from 6–15 weeks of age). As
expected C57Bl/10 mice showed significantly increased body weight and % body fat,
however; the bodyweight of mdx mice was unaffected by the change in diet. The
dystropathology of skeletal muscles was not examined in this preliminary study, but
these interesting results led us to propose that a high fat diet may reduce the extent of
dystropathology, and may be metabolically beneficial to mdx mice due to their
impaired energy status - see chapter 6 of this Thesis.
The information presented in this review of background literature summarises 3 main
points 1) the suitability of the mdx mouse as an animal model for pre-clinical DMD
research, 2) the dystropathology of mdx mice, in particular myofibre necrosis and 3)
the identification of 3 potential therapeutic interventions that could be modulated in
attempt to reduce dystropathology in mdx mice. The central hypothesis of the thesis is
that inflammation, oxidative stress and metabolic abnormality all play a role in the
extent of dystropathology and that modulation of these parameters via therapeutic
intervention can ameliorate dystropathology. The experimental research presented in
this thesis is an investigation of this central hypothesis.
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_____________________________________________________Chapter 1 - Introduction
1.6 Aims and Hypotheses of the Thesis
Overall Aim:
To identify potential therapeutic interventions for Duchenne Muscular Dystrophy using
the mdx animal model.
Specific Aims:
1. Examine the potential therapeutic benefits of an anti-inflammatory drug (specifically
cV1q a mouse specific TNF antibody) in both young and adult dystrophic mdx mice.
(Chapter 3).
2. Develop a short (30 minute) and repeatable treadmill exercise protocol to efficiently
test pre-clinical therapeutic drugs in adult mdx mice and identify key markers of
muscle damage to rapidly monitor efficacy of pre-clinical drug treatments. (Chapter 4).
3. Examine the potential therapeutic benefits of an anti-oxidant drug (specifically Nacetylcysteine) in adult mdx mice and determine (if any) the mechanism(s) of
protective action. (Chapter 5).
4. Examine the potential therapeutic benefits of a high fat and high protein diet in both
sedentary and exercised dystrophic mdx mice, with respect to i) myofibre integrity; (ii)
muscle growth and maintenance of muscle mass; (iii) the balance between muscle and
fat formation. (Chapter 6).
5. Determine the metabolic differences (food intake, energy expenditure, activity level,
body composition and protein synthesis rate) between young and adult mdx and
C57Bl/10 mice. (Chapter 7).
Overall Hypothesis:
The central hypothesis of the thesis is that inflammation, oxidative stress and
metabolic abnormality all play a role in the extent of myofibre necrosis in dystrophic
muscle and that modulation of these parameters via therapeutic intervention can
ameliorate dystropathology.
39
_____________________________________________________Chapter 1 - Introduction
Specific Hypotheses:
1. TNF plays a key role in necrosis of dystrophic muscle and in vivo administration of
cV1q a TNF antibody will ameliorate dystropathology in both young and adult mdx
mice.
2. A single 30 minute treadmill exercise protocol will increase markers of muscle
damage in adult mdx mice and thus provide a quick and efficient way to screen
potential pre-clinical therapeutic drugs in adult mdx mice.
3. Oxidative stress plays a key role in myofibre necrosis of dystrophic muscle in vivo
and
administration
of
N-acetylcysteine
(an
antioxidant)
will
ameliorate
dystropathology in both sedentary and treadmill exercised adult mdx mice.
4. Dystrophic muscle is subject to metabolic abnormalities and consumption of a high
fat or high protein diet will ameliorate dystropathology in both sedentary and
treadmill exercised adult mdx mice.
5. Due to the continuous cycles of muscle necrosis and regeneration in mdx mice there
will be significant metabolic differences between mdx and C57Bl/10 mice. In addition,
these differences will be amplified in young mdx mice where dystropathology is most
severe.
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____________________________________________Chapter 2 – Materials and Methods
Materials and Methods.
Overview: This methods chapter contains both an unpublished part and 1 published
methods protocol (Publication #4). Details of the published protocol along with the
candidate’s individual contribution to this protocol are listed below. A re-print of the
published protocol is included in section 2.5 of the Methods chapter.
1) Grounds M.D, Radley-Crabb H.G, Nagaraju K, Aartsma-Rus A, van Putten M, Rüegg
M.A, de La Porte S. Hematoxylin and Eosin staining for histology of dystrophic skeletal
muscle M.1.2_007. TREAT NMD Neuromuscular Network. Pre-clinical testing for DMD:
Endpoints in the mdx mouse. http://www.treat-nmd.eu/research/preclinical/SOPs/
Candidate completed approximately 40% of the work required for the method.
Each paper (each Results chapter) within this thesis contains all specific information for
the experimental methods specific to that paper. The following general text (Chapter
2) describes in detail the routine experimental methods employed throughout this
Thesis.
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2.1 In vivo animal protocols:
2.1.1 Animal usage and animal ethics: All experiments were carried out in nondystrophic (normal) C57Bl/10ScSn and dystrophic mdx (C57Bl/10mdx/mdx) mice.
C57Bl/10ScSn mice are the parental strain for mdx. For experiments conducted at the
University of Western Australia, all mice were obtained from the Animal Resource
Centre, Murdoch, Western Australia. Mice were maintained at the UWA on a 12-h
light/dark cycle, under standard conditions, with free access to food and drinking
water. All animal experiments were conducted in strict accordance with the guidelines
of the National Health and Medical Research Council Code of practice for the care and
use of animals for scientific purposes (2004) and the Animal Welfare act of Western
Australia (2002) and were approved by the Animal Ethics committee at the University
of Western Australia.
(For details of animal experiments conducted in the USA see section 2.4 of this Materials and Methods
chapter).
2.1.2 Voluntary exercise: The low level of muscle damage in dystrophic adult mdx mice
can be elevated by voluntary exercise (Grounds et al, 2008b). One individual mouse is
placed into a cage with a voluntary running wheel (Figure 2.1) and allowed free access
to the wheel. Large perspex cages are used and the wheel is mounted to the roof of
the cage; this allows for the height of the wheel (300mm) and prevents bedding on the
cage floor from blocking up the exercise wheel.
Figure 2.1. Voluntary exercise set-up. A metal exercise wheel (arrow) is attached to the cage
lid with cable ties and the monitor from a bicycle speedometer (*) is taped to the inside of the
mouse’s cage. The bicycle monitor records distance and speed run by the mouse.
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Exercise data were collected via a small magnet attached to the mouse wheel, and a
sensor from a bicycle pedometer attached to the back of the cage. The pedometer
records single wheel revolutions, allowing total distance (km) run by an individual
mouse to be determined, as per (Hodgetts et al, 2006; Radley et al, 2008; Radley and
Grounds, 2006). The mice run the most during the night since they are normally
nocturnal (Hayes and Williams, 1996; Radley and Grounds, 2006). Mice are monitored
daily throughout the experiment for food consumption, distance run and wheel
function (See Results chapters 3 & 6).
2.1.3 Treadmill exercise: Based on previous research (De Luca et al, 2003; Granchelli et
al, 2000) and as per the TREAT-NMD recommended standard protocol “Use of
treadmill and wheel exercise for impact on mdx mice phenotype M.2.1_001”
http://www.treat-nmd.eu/research/preclinical/SOPs/ a treadmill exercise regime
consisting of 30 minutes treadmill running at a speed of 12m/minute was used. The
rodent treadmill used was an Exer 3/6 from Columbus Instruments (USA) (Figure 2.2).
Treadmill Setup: Individual running lanes were separated by clear Perspex dividers so
that the mice could see each other while exercising. The treadmill was horizontal (0°
incline) and mdx mice were run in groups of 3 or 4 as it is time consuming and
inefficient to run mdx mice individually (Figure 2.2). For consistency, the treadmill
running was routinely conducted between 8am – 11am.
Figure 2.2. Treadmill exercise set-up. Lanes are separated by clear perspex dividers (arrow) and each
mouse is given an individual lane to run in. Soft foam blocks (*) are placed at the end of each lane to
prevent the mouse falling off the back of the treadmill.
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____________________________________________Chapter 2 – Materials and Methods
Exercise Protocol: Groups of 3 or 4 mdx mice were all (1) settled for 2 mins with the
treadmill belt stationary, (2) then acclimatized with gentle walking for 2 mins at a
speed of 4m/min, followed immediately by (3) a warm–up of 8 minutes at 8m/min and
then (4) the main exercise session for 30 minutes at 12m/min. If during the 30 minutes
exercise session a mouse fatigued and could no longer run, the procedure was as
follows: turn the treadmill belt off and give all mice a 2 minute rest, turn the belt on at
4m/min for 2 minutes, increase the speed to 12m/minute and run for the remainder of
the 30 minutes. Repeat this process if fatigue occurs again (up to 5 times for an
individual mdx mouse) (Radley-Crabb et al, submitted) (See Results Chapters 4 & 5).
2.1.4 Forelimb Grip strength: Grip strength was measured using a Chatillon Digital
Force Gauge (DFE-002) and a triangle metal bar, as per the TREAT-NMD recommended
standard protocol “Use of grip strength meter to assess limb strength of mdx mice –
M.2.2_001” http://www.treat-nmd.eu/research/preclinical/SOPs/. In brief, the mouse
was placed on the front of the triangle bar (attached to a force transducer) and pulled
gently until released. Each mouse underwent 5 consecutive grip-strength trials; the
grip strength value for each mouse was recorded as the average of the 3 best efforts.
Average grip strength was then normalized for body weight [Force (kg)/BW (g)].
Change in normalised grip strength was determined by subtracting normalised grip
strength (8 weeks) from normalised grip strength (12 weeks) (See Results Chapter 4).
2.1.5 Dietary interventions: The three customised semi-pure diets; Control (7% fat,
19% protein - AIN93g), High Fat (16% fat, 19% protein - sf 06-040) and High Protein (7%
fat, 50% protein - sf 00-252) were manufactured by and purchased from Specialty
Feeds Glen Forest Western Australia (www.specialtyfeeds.com.au). The three diets
were designed and manufactured specifically for laboratory mice by Mr Warren Potts.
Pilot studies were conducted with all custom diets for palatability, consumption and
toxicity. All custom diets were administered as standard 12mm cubes. All mice were
fed a cereal based standard rat and mouse chow (5% fat, 19% protein) prior to the
commencement of any dietary interventions (this is the standard cereal diet fed to all
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____________________________________________Chapter 2 – Materials and Methods
mice at UWA). Mice were monitored twice weekly for food consumption (weight g)
and bodyweight (See Results Chapter 6).
2.1.6 Intra-peritoneal injections of cV1q antibody (Anti-inflammatory drug
administration): Intra-peritoneal (IP) injections of cV1q and the isotype-matched,
negative control antibody (cVaM) both provided by Centocor were given at a
concentration of 20µg/g bodyweight /week. This procedure is simple and requires no
anaesthesia; mice were picked up (scruffed) and held by a researcher while being
injected. A 27.5 gauge insulin syringe (Sigma Aldrich Z192082) was used for the
injection. Both cV1q and cVaM were diluted in PBS (See Results Chapter 3 & 8).
2.1.7 Drinking water administration of N-acetylcysteine (Anti-oxidant drug
administration): N-acetylcysteine (Sigma Aldrich A7250) was added to standard
acidified laboratory mouse drinking water (pH 2.5 – 3) at a concentration of either 1%
or 4%. Mice were then given free access to the treated drinking water for 6 weeks (1%)
or 1 week (4%). Drinking water was changed once a week as per strandard animal
husbandry practices. Mice were monitored every second day for drinking water
consumption (ml) and bodyweight. It is estimated that laboratory mice drink
approximately 3-5mls per day, this equates to 30-50µg N-acetylcysteine consumption
per day, (e.g 1.6µg NAC/g bw/day [for a 30g mdx mouse]) (See Results Chapter 5).
2.1.8 Evans Blue Dye injection: A single Evans Blue Dye (Sigma Aldrich 46160) injection
was given intraperitoneally (IP) 24 hours prior to animal sacrifice and sampling (Hamer
et al, 2002). A 1% solution of EBD (diluted in filtered PBS) was injected at a dose of
1mg/g bodyweight (1% of mouse’ bodyweight). A 27.5 gauge insulin syringe (Sigma
Aldrich Z192082) was used for the injection.
2.1.9 Whole muscle graft surgery: Whole muscle autografts were used as an in vivo
bio-assay to assess the anti-inflammatory properties of cV1q in non-dystrophic
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____________________________________________Chapter 2 – Materials and Methods
C57Bl/10 mice. Six weeks old female C57BL/10 mice were anaesthetised using 2% (v/v)
isoflurane (Bomac Australia). The Extensor Digitorum Longus (EDL) muscle with both
tendons was removed from the anatomical bed and transplanted onto the surface of
the Tibialis Anterior (TA) muscle, the EDL tendons were sutured to the TA, the skin
closed and wound left to heal, as described in (Grounds et al, 2005b; Roberts and
McGeachie, 1992; Shavlakadze et al, 2009; White et al, 2000) (Figure 2.3). It is well
documented that this procedure severs the nerve and blood supply to the EDL muscle
and results in necrosis and subsequent inflammation and regeneration of the graft;
grafts were examined histologically 5 or 7 days after surgery (See Results Chapter 3).
Figure 2.3 Schematic diagram of whole muscle graft surgery. The EDL is removed from the muscle
bed and placed on top of the TA muscle. The EDL is sutured into place and collected after 5-7 days.
2.1.10 Animal sacrifice and tissue collection: An individual mouse was placed into a
small perspex box connected to an anaesthetic machine for initial anaesthesia with 2%
v/v isoflurane (Bomac Australia) (see section 2.3.1). The anaesthetised mouse was
then transferred into a small nose cone and remained under anaesthesia while 0.5ml
of blood was collected from the mouse via direct cardiac puncture using a 27.5 gauge
tuberculin syringe (Sigma Aldrich Z192082) - it is necessary to do this while the heart is
still beating (see section 2.3.4). The anaesthetised mouse was then sacrificed by
cervical dislocation. The tissues were collected in a strict sequence: firstly, tissues were
rapidly collected for molecular or biochemical analysis by snap freezing immediately in
liquid nitrogen. Secondly, tissues were collected for either frozen or paraffin histology.
Lastly, tissues were collected for weight and morphometric measurements (e.g tibial
length).
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2.2 Histological protocols:
2.2.1 Sample preparation: For paraffin histology, muscles were removed from the
mouse and immediately placed into 4% paraformaldehyde (Sigma Aldrich 158127) for
fixation. Where possible, placing an entire leg into paraformaldehyde allows the
muscles to hold their shape and length, which minimises curling of the muscles. The
muscles were left in paraformaldehyde for up to 48hours (depending on the size of the
muscle/leg) and once fixed moved into 70% ethanol. As an alternative to
paraformaldehyde, various commercially available formalin solutions (Confix –
Australian BioStain) can be used for basic muscle histology although these may not be
suitable for antibody staining. After fixation, muscles were cut transversely, ‘caged’
and processed overnight in an automatic processor e.g. Shandon, (ethanol through to
paraffin wax) and embedded into paraffin blocks. Skeletal muscle tissue sections were
generally cut at a width of approximately 5µm for paraffin blocks, collected onto
uncoated glass slides and stored in the dark at room temperature until stained.
For frozen histology, the fresh muscles were bisected transversely and the 2 pieces
mounted side-by-side on cork squares using tragacanth gum (Sigma Aldrich G1128).
The muscles were routinely frozen in a slurry of isopentane cooled in liquid nitrogen.
Isopentane reduces surface tension and avoids trapping air around the muscle (which
can slow the freezing process) thus producing a better frozen sample and excellent
histology (with little or no freeze artefact). Frozen sections were cut at 8µm (on a
cryostat) directly onto uncoated or silinated glass slides: ideally these were stained
immediately but could be stored at -20ºC until stained.
2.2.2 Haematoxylin and Eosin stain: Skeletal muscle pathology can be accurately
assessed on frozen or paraffin sections stained with Haematoxylin and Eosin (H&E)
(Publication #4 - section 2.5) (Grounds et al, 2008b). Haematoxylin stains eosinophilic
structures (e.g. muscle sarcoplasm) pink and Eosin stains basophilic structures (e.g.
nuclei) dark purple, with high RNA producing paler purple staining in the cytoplasm
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____________________________________________Chapter 2 – Materials and Methods
(e.g. in young myotubes). The basic H&E staining protocol was: a) Dewax paraffin
sections – incubate slides at 60ºC for 20minutes. b) Dewax sections – Xylene wash for
3 minutes (repeat x3). c) Rehydrate sections – 100% ethanol for 3 minutes (repeat x2).
d) Rehydrate sections – 70% ethanol for 3 minutes. e) Rinse – double distilled water for
1-3 minutes. f) Stain - Haematoxylin for 30 seconds (Note – Can start staining frozen
sections at ‘f’). g) Stain - Remove excess stain in tap water (until nuclei turn blue). h)
Stain - 70% ethanol for 3 minutes. i) Stain - Eosin for 15 seconds. j) Dehydrate – 100%
ethanol for 3 minutes (repeat x3). k) Clear – Xylene wash for 3 minutes (repeat x3). l)
Mount - DPX mountant and coverslip.
2.2.3 Sirius red stain: Sirius red stain is used to assess fibrosis as it stains collagen red.
Sirius red stain is also useful to measure myofibre cross sectional area as it stains
myofibres yellow and collagenous myofibre membranes red. The Sirius red staining
protocol was: a) Bring sections to distilled water (heat for 20mins at 60ºC, dewax in 3x
changes of xylene, dehydrate in 2x changes of 100% ethanol, 1x change of 70% ethanol
and rinse in distilled water). b) Stain nuclei with Haematoxylin for 1 minute. c) Rinse
well in tap water. d) Stain in Solution A – Pico Sirius red for 70 minutes. This gives an
equilibrium staining which does not increase with a longer time. e) Wash in two
changes of Solution B – Acidified water. (Separate the 500ml of acidified water into 2
glass staining containers, discard after use). f) Remove most of the acidified water
from the slides by shaking and blotting (do not rinse). g) Dehydrate in 3x changes of
100% ethanol. h) Clear in 3x changes of xylene and mount with DPX.
Solution A: Pico-sirius red: Sirius Red stain F3B (CI 35782) – 0.5g. Saturated aqueous
solution of picric acid – 500ml (add a little solid picric acid to the solution to ensure
saturation). Do not filter.
Solution B – Acidified water: Add 2.5ml of acetic acid to 500ml of distilled water. Make
up fresh for each run and discard after use.
2.2.4 Image collection and analysis: Non-overlapping tiled images of transverse
muscle sections provided a picture of the entire muscle cross section. Images were
acquired using a Leica DM RBE microscope, a personal computer, a Hitachi HVC2OM
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digital camera, Image Pro Plus 4.5.1 software and Vexta stage movement software.
Histological analysis of muscle morphology was carried out on whole muscle cross
sections. Muscle morphology was drawn manually by the researcher using Image Pro
Plus 4.5.1 software and all section analysis was done ‘blind’. Muscle sections were
routinely analysed for the following morphological features: normal /undamaged
myofibres (myofibres with peripheral nuclei), myofibre necrosis (myofibres with
fragmented sarcoplasm and/or areas of inflammatory cells), regeneration (myofibres
with central nuclei), fibrosis and myofibre cross-section area (Grounds et al, 2008b). All
Histological analysis was completed as per the TREAT-NMD recommended standard
protocol “Hematoxylin and Eosin staining for histology of dystrophic skeletal muscle M.1.2_007” http://www.treat-nmd.eu/research/preclinical/SOPs/ (Publication #4 section 2.5).
2.3 Molecular and Biochemical protocols:
2.3.1 Sample preparation: When using tissue samples for molecular or biochemical
analysis is it important to remove the tissue from the mouse and snap freeze it
immediately in liquid nitrogen to prevent degradation of various parameters. Snap
frozen tissues were stored at -80ºC until analysed. Anaesthetic regimes should be
standardised as changes may affect gene expression, protein synthesis and oxidative
stress (Heys et al, 1989; Juhl, 2010). The standard terminal anaesthesia regime used
throughout this thesis is: 400ml NO2 (BOC Australia), 1.5L O2 (BOC Australia) and 2%
v/v isoflurane (Bomac Australia) (see section 2.1.10).
2.3.2 RNA extraction and reverse transcription: RNA was extracted from either the
snap frozen gastrocnemius or quadriceps muscle using Tri-reagent (Sigma T9424) and
quantitated using a Nano Drop Spectrophotometer (ND 1000) and ND 1000 software
version 3.5.2. The RNA was DNAse treated using Promega RQ1 RNAse free DNAse
(M610A), RQ1 RNAse free 10x buffer (M198A) and RQ1 DNAse stop solution (M199A).
RNA was reverse transcribed into cDNA using Promega M-MLV Reverse Transcriptase
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(M3682), random primers (C1181) and 10mM dNTPs (U1515) and the cDNA was
purified using a MoBiol Clean up kit (12500-250).
2.3.3 Reverse Transcription Polymerase Chain Reaction (RT-PCR): RT-PCRs were run
on a Corbett 3000 (Corbett Research) using QIAGEN quantifast SYBR green PCR mix
(204054) and QIAGEN Quantitect Primer Assays for TNF (QT00104006), IL-1β
(QT01048355), IL-6 (QT00098875), PPAR alpha (QT00137984), PPAR beta/delta
(QT00166292), PPAR gamma (QT00100296), PGC-1alpha (QT00095578)
and
standardised to a house keeping gene L-19 (QT01779218). mRNA expression was
calculated and standardised using Roto-gene 6.1 and Microsoft Excel software (See
Results Chapters 4 & 6).
2.3.4 Creatine Kinase assay: While mice were under terminal anaesthesia, whole
blood (approx 0.5ml) was collected via cardiac puncture using a 27.5 gauge tuberculin
syringe (Sigma Z192082), into a 1.5ml tube (see section 2.1.10). Extensive
experimentation revealed that storage of blood samples overnight at 4ºC to enable
clotting leads to a false increase in serum CK levels and therefore blood samples were
immediately spun down in a refrigerated centrifuge for 5 minutes (12000g), serum was
removed and aliquoted (Radley-Crabb et al, submitted). Blood serum CK activity was
determined in duplicate using the CK-NAC kit (Randox Laboratories) and analysed
kinetically using a BioTek Powerwave XS Spectrophotometer using the KC4 (v 3.4)
program. A minimum of 10µl serum is required to complete this assay. Prior to
establishing this assay in our laboratory some CK assays were performed commercially
at Murdoch Veterinary Hospital, WA (Radley et al, 2008).
2.3.5 TNF Elisa: A commercially available TNF Elisa (Invitrogen KMC3012 pre-coated 96
well plate) was used to measure TNF protein level in blood serum, according to
manufacturer’s instructions. In brief the method consisted of: 1) Add standards, water
blank, chromagen blank and TNF controls to first 12 wells. 2) Add 50µl of standard
diluent buffer and 50µl of each serum sample to the remaining wells. 3) Add 50µl of
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biotinylated anti-TNFa solution into each well (except chromagen blank) and tap plate
to mix. 4) Cover plate and incubate at room temperature for 90 minutes. 5) Prepare
working solution of Streptavidin-HRP (at approx 70 mins incubation). Warm
streptavidin-HRP concentrate (100x concentrate in 50% glycerol) to room temperature
and gently mix. Dilute 10ul of streptavidin-HRP concentrate in 1ml of streptavidin-HRP
diluent (for each 8 well strip). 6) After 90 minutes empty wells and wash x4. 7) Add
100ul of Streptavidin-HRP working solution to each well (except chromagen blank). 8)
Cover plate and incubate at room temperature for 30 minutes. 9) Empty well and wash
x4. 10) Add 100ul of stabilized chromagen to each well (liquid in the wells will begin to
turn blue). 11) Incubate at room temperature (and in the dark) for 30 minutes.
Incubation time is determined by the microtiter plate reader used. Some plate readers
have a maximum optical density (O.D) of 2.0. O.D values should be monitored and the
reaction should be stopped before the positive wells exceed the instruments limit (2.0
O.D lies between std 2 and std 3). The O.D values can only be read at 450nm once the
stop solution has been added. If using an instrument that has a maximum O.D of 2.0 an
incubation time of 20 -25mins is suggested. 12) Add 100ul of stop solution to each
well. 13) Blank the plate against the chromagen blank (well 10 – contains only
chromagen and stop solution). 14) Read the absorbance of each well at 450nm
(generate a standard curve for standards/wells 1-8). 15) Correct for dilutions of
controls and samples in step 4,5,6 ( i.e. added 50ul sample and 50ul of dilution buffer
so need to multiply mouse TNF values by 2).
2.3.6 Novel ROS analysis (ratio of oxidised to reduced protein thiols): The following
method was conducted by Miss Jessica Terrill (co-author on Publications #6&7).
Oxidative stress was measured as a ratio of oxidised (di-sulphide) to reduced
(sulfhydryl) protein thiols using a novel technique (USA PATENT #20080305495)
developed by Dr Peter Arthur (Armstrong et al, submitted)(Lui et al, 2010). Muscle
samples were collected very rapidly and immediately snap frozen in liquid nitrogen
(section 2.1.10). Frozen quadriceps muscles were crushed under liquid nitrogen,
before protein extraction with 20% trichloroacetic acid/acetone.
Protein was
solubilised in 0.5% sodium dodecyl sulphate 0.5 M tris (SDS buffer), pH 7.3 and thiols
were labelled with the fluorescent dye BODIPY FL-N-(2-aminoethyl) maleimide (FLM,
85
____________________________________________Chapter 2 – Materials and Methods
Invitrogen).
Following removal of the unbound dye using ethanol, protein was
resolubilised in SDS buffer, pH 7 and oxidised thiols were reduced with tris(2carboxyethyl) phosphine (TCEP, Sigma) before the subsequent unlabelled reduced
thiols were labelled with a second fluorescent dye texas red maleimide (Invitrogen).
The sample was washed in pure ethanol and resuspended in SDS buffer. Samples were
read using a fluorescent plate reader (Fluostar Optima) with wavelengths set at
excitation 485, emission 520 for FLM and excitation 595, emission 610 for texas red. A
standard curve for each dye was created using ovalubumin and all results were
expressed per mg of protein, quantified using Detergent Compatible protein assay
(BioRad) (See Results Chapters 4 & 5).
2.4 Metabolism protocols: In May 2008, the candidate participated in a 5 week
internship in the laboratory of Dr Marta Fiorotto, Baylor College of Medicine, Houston
USA. The following protocols were all conducted during the internship and have been
written-up as a collaborative manuscript (Publication #9 – Results Chapter 7).
2.4.1 Animal usage and animal ethics: All animal experiments and analysis were
carried out at The Children’s Nutrition Research Centre (CNRC) at Baylor College of
Medicine, Houtston USA. Experiments were conducted in strict accordance with the
U.S. National Research Council’s Guide for the Care and Use of Laboratory Animals and
all procedures were approved by the Baylor Medical College Animal Care and Use
Committee. The young 3 - 4 week old male mice (mdx and C57Bl/10ScSn) were bred at
the CNRC from breeders originally from Jackson Laboratories.
2.4.2 Whole Body Composition Scans. Total body lean muscle and fat mass were
measured by dual X-ray absorptometry (DXA) using a PIXImus (General Electric, USA).
Whilst under isoflurane anaesthesia mice were aligned on a PIXImus measuring tray
and then inserted into the machine for measurements. All mice were measured in
duplicate and then returned to their cage to recover from the anaesthetic, a minimum
of 2 days elapsed before protein turnover measurements were performed. Correction
factors derived specifically for the CNRC PIXImus were applied to the fat and lean mass
86
____________________________________________Chapter 2 – Materials and Methods
data to adjust for the inherent errors in fat and lean mass obtained using all PIXImus
instruments (http://www.bcm.edu/cnrc/intranet/MMRU/PIXImus%20corrections.xls).
Figure 2.4. Whole body
composition scans in a
PIXImus machine. Mice
were anaesthetised
(arrow), laid out on a
measuring tray and
composition scans were
performed in duplicate.
2.4.2 Calorimetry. Mice were placed individually into calorimetry chambers for 72 hrs
and energy expenditure was measured using a Columbus Instruments (Columbus OH)
CLAMS Oxymax System. A known flow of air was passed through the chambers and the
O2 and CO2 gas fractions were monitored at both inlet and outlet ports. The gas
fraction and flow measurements were used to calculate VO2 and VCO2, from which the
Respitatory Exchange Ratio and heat production (kcal/hr) are calculated. At the same
time spontaneous activity level (both horizontal and vertical movements) was
monitored from the interruption of infra-red beams projected across the cage (See
Results Chapter 7).
2.4.3 Isotope infusion and tissue collection to measure rate of protein synthesis.
Food was removed from all mice 7 hours prior to isotope infusion. A flooding dose of L4-[3H]-phenylalanine (American Radiolabeled Chemicals, Inc., St. Louis, MO) at
20mL/kg BW, 1.5mmol phenylalanine/kg and 250µCi/mouse was given through the tail
vein fifteen minutes after injection mice were sacrificed (by decapitation) and trunk
blood was immediately collected and acidified to 0.2M Perchloric acid (PCA). The
supernatant was frozen and reserved for estimation of the specific radioactivity of
blood free phenylalanine pool. The hindlimbs were immediately detached, wrapped in
foil and chilled on ice. A range of muscles including; gastrocnemius and plantaris
complex (referred to hereafter as the gastrocnemius), quadriceps, soleus, tibialis
87
____________________________________________Chapter 2 – Materials and Methods
anterior, diaphragm and heart, were then dissected on ice and frozen in liquid
nitrogen. The weights of all muscles were recorded and muscles were stored at -80ºC.
Laboratory method: These are based on Fiorotto et al (2000) and Welle et al (1993)
with some modifications (Fiorotto et al, 2000; Welle et al, 1993).
Total Protein, and Myofibrillar Proteins isolation: Frozen muscles from each mouse
were powdered, weighed and 50mg was homogenized in a low salt homogenising
buffer (50 mM potassium phosphate, 0.25 M sucrose, 1% Triton X-100 pH 7). For each
muscle, an aliquot of the homogenate was retained for measurement of total protein;
a second aliquot was acidified to 0.2M PCA and centrifuged; the supernatant
containing the muscle free amino acid precursor pool was collected and neutralized as
described below. The PCA-insoluble precipitate was washed, and after assaying for
total RNA, were processed for the measurement of L-[4-3H] phenylalanine
incorporated into all muscle proteins. The remainder of the low salt/sucrose buffer
homogenate was left to incubate with agitation at 4°C for 60 minutes; the proteins
that were insoluble after low speed centrifugation (1,500 g at 4ºC for 10min) contained
the myofibrillar proteins (Fiorotto et al, 2000; Welle et al, 1993). After several washes
of the the pellet in the low salt/sucrose buffer followed by ice-cold water, the purified
myofibrils were acidified to 0.2M PCA. The acid insoluble myofibrillar and total protein
precipitates were separated from the supernatant by centrifugation at 10,000 g at 2°C
for 30 min, and the pellets were washed three times in 0.2M PCA. The supernatant
from the total protein fraction and the pellets from all protein fractions were
processed for determination of L-[4-3H] phenylalanine specific radioactivity.
Quantification of L-[4-3H] phenylalanine specific radioactivity in the protein bound
and tissue free amino acid precursor pools: The acid-insoluble pellets were first
hydrolysed for 24 hrs in 6M HCL(ultrapure) at 110oC under N2 gas. After evaporation of
the HCl in a Savant Speedvac concentrator, the remaining precipitate was subject to
several washes in water and finally resuspended in mQ water for HPLC analysis. The
PCA-treated blood and muscle free pool supernatants samples were neutralized on ice
with 4M KOH, evaporated to dryness and resuspended in 1M acetic acid. The amino
acids were purified over a Dowex anion exchange column (AG-50w-x8, 100 - 200 mesh,
Biorad). The amino acids were eluted in 3M NH4OH evaporated to dryness in the
Savant Speedvac concentrator and finally dissolve in MQ water. All samples were
filtered using a 0.45µm syringe filter, supernatant dried down and each sample was re88
____________________________________________Chapter 2 – Materials and Methods
dissolved in 100ul of MQ water. Phenylalanine in the total and myofibrillar protein
hydrolysates, muscle homogenate and blood supernatants were isolated by anion
exchange HPLC (AminoPac1 Analytical column; Dionex, Sunnyvale, CA). Amino acids
were post-column derivatized with o-phthalaldehyde reagent and detected with an online fluorimeter. The fraction of the eluant that included the phenylalanine peak was
collected, and the associated radioactivity was measured in a liquid scintillation
counter (Packard Tricarb, Perkin Elmer, Waltham, MA). The phenylalanine
concentration was determined by comparing the peak areas of the samples with that
of a known standard (Pierce Labs.)
Calculations of in-vivo fractional protein synthesis rates:
The fractional rate of
protein synthesis (FSR), i.e., the percentage of the protein mass synthesized in a day
was calculated as follows: FSR = (SB/ SA )  (1440/t) 100
Where SB is the specific radioactivity of the protein-bound labelled amino acid, SA is the
specific radioactivity of the precursor pool and t is the the labelling time in minutes
(Davis et al, 1999a). It has been demonstrated that the specific radioactivity of the
tissue free phenylalanine after a flooding dose of phenylalanine is in equilibrium with
the aminoacyl-tRNA specific radioactivity; hence, the tissue free phenylalanine reflects
the specific radioactivity of the tissue precursor pool (Davis et al, 1999b). Blood and
tissue free phenylalanine specific radioactivity values were compared to verify
equilibration.
Protein quantification: Total protein concentration of all muscle samples was
determined using a standard BCA protein (Smith et al, 1985) assay after first
solubilising the aliquot of total muscle homogenate in 0.1 M NaOH for 1 hr at 37 oC.
Total RNA measurement: Because the RNA content of a tissue is dominated by
ribosomal RNA, total RNA was measured quantitatively on all muscle samples and
provides an estimate of ribosomal abundance. Total RNA was quantified in the total
protein PCA-insoluble precipitate using a modified Schmidt-Thannhauser procedure as
described by Munro and Fleck (1966) (Munro and Fleck, 1966).
2.5 TREAT-NMD SOP published protocol:
Re-print included in pages 90-100.
89
_______________________________________________________________Chapter 3.
Chapter 3 (Results 1).
Reduced muscle necrosis and long-term benefits in dystrophic mice after cV1q
(blockade of TNF) treatment.
Tumour necrosis factor (TNF) is a potent inflammatory cytokine that exacerbates
damage of dystrophic muscle damage in vivo (see Introduction 1.4). This chapter
examined the effects of cV1q, a monoclonal murine specific antibody that neutralises
mouse TNF, in dystrophic mdx mice in vivo. This work was published in 2008
(Publication #5) in the Journal of Neuromuscular Disorders 18: 227-238 and has
received 12 citations (as of September 2010). The candidate completed approximately
85% of the work required for this manuscript.
There is strong evidence to suggest that inflammatory cells and cytokines play a role in
skeletal muscle damage [reviewed in (Evans et al, 2009a; Grounds et al, 2008a; Tidball,
2005)]. Antibody blockade of TNF with the human/mouse chimeric antibody infliximab
(Remicade®) in young mdx mice results in a striking protective effect on dystrophic
myofibres and suppresses the early acute phase of myofibre necrosis (Grounds and
Torrisi, 2004). A similar protective effect in young dystrophic muscle was
demonstrated with etanercept (Enbrel®) a soluble TNF receptor (Hodgetts et al, 2006;
Pierno et al, 2007).
These results strongly support a key role for TNF in both
inflammation and necrosis in dystrophic muscle.
There is controversy regarding the cross-species binding of infliximab based on in vitro
tests, yet at least 4 papers report anti-inflammatory effects in vivo in mice (Grounds
and Torrisi, 2004), rats (Oruc et al, 2004; Woodruff et al, 2003) and pigs (Olmarker et
al, 2003). As a result this controversy Centecor (USA) generally supplied us with the
mouse specific anti-TNF antibody cV1q in 2007. The present paper tested the
effectiveness of the cV1q (mouse-specific anti-TNF) antibody in comparison to
infliximab in both dystrophic and non-dystrophic mice, in both short and long-term (up
to 90 days) studies combined with voluntary exercise. Treatment with cV1q
significantly delayed inflammation and thus muscle regeneration in whole muscle
101
_______________________________________________________________Chapter 3.
autografts in normal non-dystrophic C57Bl/10 mice and confirmed the bio-activity of
cV1q. TNF antibody treatment (cV1q treatment) of dystrophic mdx mice reduced the
extent of dystropathology in both young mdx litters and voluntarily exercised adult
mdx mice. This paper highlights the central role of TNF in myofibre necrosis in
dystrophic muscle and suggests the use of anti-inflammatory drugs as a potential
clinical therapy for DMD patients.
Additional experimental work testing the protective effects of the cV1q antibody on in
vivo functional parameters in adult mdx and normal mice using a dynamometer was
conducted in collaboration with Mr Adam Piers, an Honours student, in 2009 and
presented in Chapter 8 of this thesis (Publication #10).
102
_______________________________________________________________Chapter 4.
Chapter 4 (Results 2).
The early consequences of two different treadmill exercise protocols on dystrophic
mdx mice.
This chapter describes in detail two standardised protocols for the treadmill exercise of
mdx mice and profiles changes in molecular and cellular events after a single 30
minute treadmill session or after 4 weeks of treadmill exercise. This work was resubmitted in November 2010 to the Journal of Neuromuscular Disorders (Publication
#6). The candidate completed approximately 75% of the work required for this
manuscript.
Bi-weekly treadmill exercise for 4 weeks is widely used in pre-clinical experiments to
increase the extent of dystropathology in mdx mice, yet the cellular consequences of a
single 30 minute treadmill exercise session have not been described. The present study
standardised a protocol and conducted time-course analysis of molecular and cellular
changes after a single 30 minute treadmill exercise session (Protocol A). These data
were compared to data from age matched mdx mice subjected to 4 weeks of treadmill
exercise (Protocol B) a protocol currently widely used for pre-clinical drug screening in
mdx mice (Burdi et al, 2009; De Luca et al, 2003; Granchelli et al, 2000; Grounds et al,
2008b).
Both treadmill protocols increased multiple markers of muscle damage. It was
concluded that a single 30 minute treadmill exercise session is a sufficient and
conveniently fast screening test to evaluate the benefits of pre-clinical drugs in vivo.
Myofibre necrosis, blood serum CK and oxidative stress (specifically the ratio of
oxidised to reduced protein thiols) are reliable markers of muscle damage after
exercise; however sampling time (after exercise) for these parameters is critical.
A more precise understanding of the changes in dystrophic muscle after exercise aims
to identify the immediate molecular and cellular changes resulting from a single short
treadmill exercise session; this novel information has the potential to identify new
115
_______________________________________________________________Chapter 4.
reliable biomarkers and new therapeutic drug targets for Duchenne Muscular
Dystrophy.
The detailed information gathered in this study (treadmill protocol, sampling time
points and markers of muscle damage) was used to design an in vivo experiment which
examined the potential benefits of N-acetylcysteine (NAC) treatment in dystrophic
mdx mice (Chapter 5, Publication #6).
116
_______________________________________________________________Chapter 5.
Chapter 5 (Results 3).
The benefits of N-acetylcysteine treatment in dystrophic mdx mice.
In addition to inflammation (discussed in chapters 4 & 8) and altered metabolism
(discussed in chapters 6 & 7), oxidative stress is another proposed mechanism by
which the absence of functional dystrophin protein leads to myofibre necrosis.
Whether it is by direct macromolecule damage or via reversible modifications to
protein function, oxidative stress has the potential to be a major contributor to the
pathology associated with Duchenne Muscular Dystrophy [Reviewed in (Arthur et al,
2008; Tidball and Wehling-Henricks, 2007; Whitehead et al, 2006b)].
N-acetylcysteine (NAC) is an anti-oxidant which decreases oxidative stress by
scavenging ROS; NAC is highly attractive as a clinical treatment since it is readily
available, inexpensive and has few adverse side effects. This body of work initially
began as a direct comparison between the potential therapeutic benefits of the antiinflammatory drug cV1q (TNF antibody – see chapter 4) and the antioxidant drug NAC.
It was hypothesised that conducting two parallel in vivo experiments testing the
effects of both cV1q and NAC, by using identical experimental methods and analysis
measures, would enable identification of the more ‘promising’ therapeutic
intervention. Unfortunately, due to numerous problems with the cV1q experiments
(e.g. concerns regarding efficacy of different antibody batches and conflicting results),
the cV1q work was abandoned and only the NAC work was continued.
The final chapter (manuscript #7) examines the benefits of NAC treatment (at 2
different doses and treatment regimes) in both sedentary and treadmill exercised
adult mdx mice in vivo and also attempts to determine the mechanism by which NAC
works in dystrophic muscle. This chapter puts into practice the methods developed in
chapter 4, including the exact 30 minute treadmill exercise protocol, mouse sampling
time-points (after exercise) and biomarkers to measure muscle damage after treadmill
exercise.
In sedentary mdx mice NAC treatment had no effect on myofibre necrosis or
membrane damage (measured by serum CK level). NAC treatment had no effect on
158
_______________________________________________________________Chapter 5.
malondialdehyde (MDA) level or protein carbonylation (two indicators of irreversible
oxidative damage); however 4% NAC treatment did significantly reduce oxidized
glutathione and oxidized protein thiols in sedentary mdx mice. Also, in treadmill
exercised mdx mice, NAC treatment prevented exercise induced myofibre necrosis and
membrane damage. These novel in vivo data show that NAC protects dystrophic
muscle from exercise induced myofibre necrosis, the primary cause of severe muscle
pathology seen in DMD. The exact mechanism by which NAC acts is unclear at this
stage; however it appears to be multi-factorial.
This work was submitted to The International Journal of Biochemistry and Cell
Biology in January 2011 a copy of the manuscript (manuscript #7) in included. The
candidate completed approximately 40% of the work required for this manuscript.
159
_____________________________________________________________________Chapter 6.
Chapter 6 (Results 4).
The different impacts of a high fat diet on dystrophic mdx and control C57Bl/10 mice.
The absence of functional dystrophin protein in dystrophic mdx mice leads to fragile
myofibre membranes and cycles of myofibre necrosis and regeneration. Studies in
both DMD patients and mdx mice indicate that the absence of dystrophin may also
(directly or indirectly) lead to alterations in skeletal muscle metabolism and an
impaired energy status (Davidson and Truby, 2009; Dupont-Versteegden et al, 1994a;
Landisch et al, 2008; MacLennan and Edwards, 1990; Passaquin et al, 2002; Zanardi et
al, 2003). In addition to anti-inflammatory and antioxidant drug interventions such as
cV1q and NAC, aimed to reduce the severity of dystropathology (see Chapters 3, 5 &
8), dietary interventions (e.g specific amino acid supplementation) have shown
beneficial effects in both mdx mice and DMD patients [Reviewed in (Leighton, 2003;
Pearlman and Fielding, 2006; Radley et al, 2007)].
This study (Chapter 6) directly compares the response of control C57Bl/10 and
dystrophic mdx mice to a high protein (50%) and a high fat (16%) diet. Surprisingly,
consumption of a high protein diet had minimal effects on the body composition or
dystropathology of C57Bl/10 and mdx mice. In contrast, striking differences between
the strains were seen in response to the high fat (16%) diet; this response also varied
between mdx mice aged <24 weeks and up to weeks. C57Bl/10 mice demonstrated
many negative side effects after consuming a high fat diet with significant weight gain,
increased body fat and elevation of pro-inflammatory cytokines. In contrast, mdx mice
(< 24 weeks) remained lean with minimal fat deposition and were resistant to changes
in body composition after consuming a high fat diet. These results support the
proposal that dystrophic mdx mice have an altered ‘energy status’ compared to
normal C57Bl/10 mice since the mdx mice were more capable of efficiently processing
and utilising increased dietary fat. However, older mdx mice (24-40 weeks old)
exhibited some effects of a high fat diet but to a lesser extent than age matched
C57Bl/10 mice. A high fat diet significantly increased the running ability (distance - km)
of voluntarily exercised mdx mice and significantly reduced myofibre necrosis in 24
192
_____________________________________________________________________Chapter 6.
week old sedentary mdx mice, suggesting that increased dietary fat (and increased
energy intake) can assist energy deficient mdx mice metabolically, providing them with
additional energy to run further while maintaining muscle integrity. This research
clearly identifies an ‘altered’ response to a high fat diet in dystrophic mdx mice
compared to C57 controls. Energy deficient mdx mice appear to be utilising excess
dietary fat to reduce the severity of dystropathology and increase voluntary exercise
ability. Our new data highlight the potential benefits of a high fat diet on dystrophic
muscle.
It must be noted that the custom diets used in this study, high fat and high protein,
were initially designed to be isocaloric based on digestible energy content (HP – 18.2
MJ/Kg vs. HF – 18.1 MJ/Kg). Major differences in body composition were seen in
response to the high fat and high protein diets and it became apparent that
metabolisable energy content was a more appropriate way to assess energy intake of
the mice. There are large differences in the metabolisable energy content of the high
fat and high protein diet (HP – 14.09 MJ/Kg vs. HF – 16.96 MJ/Kg) which explains the
major differences in energy intake and thus body composition in response to the two
diets.
This manuscript has been prepared for submission in Neurobiology of Disease and has
been formatted accordingly. The candidate completed approximately 90% of the work
required for the manuscript.
In addition to the work presented in this chapter (Chapter 6), dietary interventions
involving an Omega-3 enriched diet and dietary interventions in mdx litters and female
mdx mice are briefly discussed in Appendix 1. These were major studies conducted
over approximately 8 months that involved approximately 100 mice; however the
work was fatally flawed due to errors in diet manufacture and experimental design.
Because of these fatal flaws and the time and resources that would have been
required to repeat this studies, the project and full analyses were terminated. This
information is provided as an Appendix (Appendix 1 in this thesis) in case reference to
the design of the study and the available data are of use for future work.
192
_____________________________________________________________________Chapter 7.
Chapter 7 (Results 5).
Comparison of metabolism and protein synthesis rates in young and adult dystrophic
mdx and control C57Bl/10 mice.
In May 2008, the candidate participated in a 5 week internship in the laboratory of Dr
Marta Fiorotto, at Baylor College of Medicine, Houston USA. The following work was
conducted during the internship and has been written-up as a collaborative
manuscript (Publication #9 – Chapter 7 that follows). The candidate completed
approximately 50% of the work required for the manuscript. The manuscript has been
prepared for submission to the Journal of Clinical Investigation and has been
formatted accordingly.
Dystrophic skeletal muscle is subject to many metabolic abnormalities that can greatly
impact on the maintenance of muscle mass and function. Repeated cycles of myofibre
necrosis and regeneration, increased demand on the sarcoplasmic reticulum to
regulate intracellular calcium, defective mitochondrial function, increases in both
protein synthesis and protein degradation rates, and disruption in nNOS signalling may
all contribute to an altered metabolic state (Dupont-Versteegden et al, 1994a; Griggs
and Rennie, 1983; Han et al, 2006; Kuznetsov et al, 1998; Landisch et al, 2008;
MacLennan and Edwards, 1990; Passaquin et al, 2002; Whitehead et al, 2006b; Zanardi
et al, 2003).
Changes in protein metabolism have been reported in various mdx muscles both in
vivo and ex vivo (Helliwell et al, 1996; MacLennan and Edwards, 1990; MacLennan et
al, 1991a; MacLennan et al, 1991b; Turner et al, 1988) and, in contrast to older DMD
patients, adult mdx mice maintain skeletal muscle mass and experience significant
skeletal muscle hypertrophy (Mokhtarian et al, 1996; Peter and Crosbie, 2006;
Shavlakadze et al, 2010; Spurney et al, 2009), indicating that protein synthesis must
outweigh protein degradation in adult mdx mice. A previous study that examined the
metabolic state of dystrophic mdx mice reported no change in the metabolic rate
[kcal/(kg bodyweight0.75. d)] of one year old adult mdx mice, yet a decrease in the
metabolic rate [kcal/(kg bodyweight0.75.d)] of 4-6 week old (mixed sex) mdx mice
238
_____________________________________________________________________Chapter 7.
(Dupont-Versteegden et al, 1994a). Mokhtarian et al (1996) also reported no
difference in total energy expenditure, basal energy expenditure and spontaneous
activity between control C57Bl/10 and mdx mice (sex unspecified) at 6-12 months of
age (Mokhtarian et al, 1996). Clearly, the mouse age, extent of dystropathology and
the presence of regenerating (growing) muscle, may have a major impact on metabolic
demand and energy usage.
The purpose of this longitudinal study (over approximately 10 days) was to measure
food intake, energy expenditure, activity level, body composition and protein synthesis
rate (of both whole muscles and of the myofibrillar fraction) of dystrophic mdx and
control C57BL/10 mice at 2 different ages; 1) Young mice aged 3-4 weeks (shortly after
the acute onset of muscle necrosis in mdx mice when muscle damage regeneration is
at a peak); and 2) Adult mice aged 13-14 weeks (when adult mdx mice exhibit a lower
level of muscle damage and dystropathology).
Young mdx mice had a ‘stunted’ growth; they were significantly lighter with reduced
fat free mass (lean muscle mass). Young mdx mice also had a significantly increased
protein synthesis rate (in both whole muscle samples and isolated myofibrillar
fraction) and increased metabolic rate (Heat kcals/24hr/kg ffm). The acute onset of
myofibre necrosis and regeneration at 3-4 weeks of age appears to have had a
significant effect on many parameters related to muscle mass and function, and
metabolic rate in young mdx mice. There was significant ‘catch-up’ growth in mdx mice
between 4 and 14 weeks of age, with 14 week old adult mdx mice being heavier, with
significantly increased muscle mass and bone length than C57Bl/10 mice. The adult
mdx mice were very lean, with significantly increased fat free mass (lean muscle mass)
and increased protein synthesis rates (in both whole muscle samples and isolated
myofibrillar fraction) compared with non-dystrophic control mice.
These findings emphasise major metabolic differences, mainly due to protein synthesis
rates, between control and dystrophic mdx mice and have many implications for
understanding the basic pathology of DMD.
239
_____________________________________________________________________Chapter 8.
Chapter 8 (Results 6).
Additional work completed during candidature; not central to the Thesis.
In addition to the 5 main results chapters included in this thesis, the candidate made a
significant contribution to 2 further published manuscripts.
Re-prints of both
manuscripts are included in this chapter (Publications #10 & 11).
1) Piers AT, Lavin T, Radley-Crabb HG, Bakker AT, Grounds MD, Pinniger G. Blockade of
TNF (using cV1q antibody) in vivo reduces contractile dysfunction of skeletal muscle in
response to eccentric exercise in dystrophic mdx and normal mice. Neuromuscular
Disorders – Nov 3rd 2010 [Epub ahead of print].
Candidate completed approximately 20% of the work required for the manuscript. The
candidate advised and trained on all aspects of in vivo work including animal
treatment, cV1q dosages, animal sacrifice and tissue sampling, histological analysis and
interpretation of histological results. This work is a direct continuation from Chapter 3
(Publication #5) in this thesis and was conducted in order to show the in vivo
functional benefits of cV1q treatment in mdx mice in addition to the histological
benefits already shown (Publication #5).
This study shows that blockade of the pro-inflammatory cytokine tumour necrosis
factor (TNF) with the mouse specific antibody cV1q, protects dystrophic skeletal
myofibres against initial contractile dysfunction and subsequent myofibre necrosis in
adult mdx mice subject to eccentric exercise. Damaging eccentric exercise was
conducted in vivo using a custom built mouse dynamometer (Hamer et al, 2002).
278
_____________________________________________________________________Chapter 8.
2) Klyen B.R, Armstrong J.J, Adie S.G, Radley H.G, Grounds M.D, Sampson D.D. 2008.
Three-dimensional optical coherence tomography of whole muscle autografts as a
precursor to morphological assessment of muscular dystrophy in mice. Journal of
Biomedical Optics. 13(1) 011003.
The candidate completed approximately 20% of the work required for the manuscript.
The candidate advised and trained on all aspects of in vivo work including animal
ethics, whole muscle autograft surgery, animal sacrifice and tissue sampling,
histological analysis and interpretation of histological results. The collaborative work
was conducted as part of an ARC grant investigating new ways of imaging whole
muscles and development of new technologies as potential alternatives to animal
sacrifice and subsequent histology.
This manuscript describes in detail the ability of Three-dimensional optical coherence
tomography (3D-OCT) to evaluate the structure and pathology of whole muscle
autografts ex vivo. 3D-OCT images, co-registered with histology, can readily distinguish
necrotic and inflammatory tissue from intact healthy muscle fibres. These preliminary
finding suggest that with further development 3D-OCT could be used as a tool for
evaluation of small-animal (mouse) morphology in vivo.
279
___________________________________________________________Chapter 9 - Discussion
Overall discussion and general conclusion.
Myofibre necrosis is the fundamental destructive process in Duchenne Muscular
Dystrophy (DMD). The exact mechanism by which the absence of dystrophin leads to
myofibre necrosis is unknown, however inflammation, oxidative stress, metabolic
abnormality, mislocalisation of nNOS and increased intracellular calcium are all heavily
implicated in the process (summarised in Figure 9.1) [Reviewed in (Evans et al, 2009a;
Evans et al, 2009b; Grounds et al, 2008a; Leighton, 2003; Radley et al, 2007; Tidball
and Wehling-Henricks, 2004b; Tidball and Wehling-Henricks, 2005; Tidball and
Wehling-Henricks, 2007; Wallace and McNally, 2009)]. It seems likely that myofibre
necrosis is due to a combination of many of these dysregulated mechanisms and not
the result of one sole mechanism. The mdx mouse is a very useful animal model to
examine the potential mechanisms by which the absence of dystrophin leads to
myofibre necrosis and to test potential therapeutic interventions to reduce the extent
of dystropathology in vivo. While replacement of the defective dystrophin by cell or
gene therapy, or molecular interventions is the definitive treatment option for DMD,
there are many problems to overcome before clinical application (Bushby et al, 2009;
Cossu and Sampaolesi, 2007; Grounds and Davies, 2007; Guglieri and Bushby, 2010;
Manzur and Muntoni, 2009; Nagaraju and Willmann, 2009; Odom et al, 2010; Odom et
al, 2007; Partridge, 2010; Tremblay and Skuk, 2008; Wells, 2008).
The present body of work examined the potential benefits of 3 therapeutic
interventions on dystrophic mdx and control mice in vivo: 1) an anti-inflammatory
drug, cV1q a mouse specific TNF antibody, 2) an antioxidant drug, N-acetylcysteine a
scavenger of ROS, and 3) dietary interventions, specifically a high fat (16%) and high
protein (50%) diet, plus metabolic analyses of the 2 strains.
All 3 therapeutic
interventions showed some benefits in mdx mice, confirming the involvement of
inflammation, oxidative stress and metabolic abnormality in myofibre necrosis in
dystrophic muscle. A detailed discussion is given for each of these interventions in the
appropriate manuscript/chapter and here an overall discussion is presented with a
focus on potential clinical relevance.
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Figure 9.1 Mechanisms for membrane degeneration in dystrophic muscle. The dystrophin glycoprotein
complex (DGC) is a complex that links the extracellular matrix component to the actin cytoskeleton in
muscle cells (Left). Absence of dystrophin causes DMD characterised by sarcolemmal disruptions (right).
In the absence of proper membrane-matrix attachment, tears may simply develop as a result of the
mechanical stress. An increase in free radicals (green triangles) may be caused by the displacement of
NOS from the plasma membrane. The high levels of free radicals in dystrophic muscle are thought to
contribute to muscle degeneration via the oxidation of muscle membranes and recruitment of
macrophages. Calcium-sensitive pathways also contribute to muscle degeneration. Calcium (pink
spheres) may enter dystrophic muscle through membrane lesions or through calcium channels. Calcium
dysregulation may also lead to abnormal mitochondrial function as well as the activation of the calciumdependent protease calpain to degrade muscle membrane proteins. Adapted from Wallace and
McNally (2009) (Wallace and McNally, 2009).
Anti-inflammatory therapy: The potential therapeutic benefits of cV1q, a monoclonal
murine specific antibody that neutralises mouse TNF, in dystrophic mdx mice were
examined (Chapter 3). Both short and long-term studies demonstrate that cV1q
antibody treatment reduces the extent of dystropathology in both young and
voluntarily exercised adult mdx mice (Radley et al, 2008) and supports the original
hypothesis that TNF contributes to necrosis of dystrophic muscle. The protective
benefits of cV1q were similar to those seen with the human/mouse chimeric TNF
antibody infliximab (Remicade®) in young mdx mice (Grounds and Torrisi, 2004) and
with etanercept (Enbrel®) a soluble TNF receptor (Hodgetts et al, 2006; Pierno et al,
2007). All these results strongly support a key role for TNF in both inflammation and
necrosis in dystrophic muscle. Interestingly, long-term treatment with cV1q had no
beneficial effects on sedentary adult mdx mice. This indicates that the already low
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___________________________________________________________Chapter 9 - Discussion
level of background dystropathology in adult mdx mice is not further reduced by cV1q
blockade of TNF, whereas cV1q clearly protected against exercise induced acute
myofibre damage (more representative of the severity of the human condition). The
lack of effect in unexercised mice is in striking contrast to the benefits of cV1q in
exercised mdx mice: whether this reflects differences in the cellular responses in these
two situations is unknown (Radley et al, 2008). The study confirms that a drug
intervention may not show benefits in unexercised mdx mice, thus exercise appears an
essential part of the ‘standard operating procedure’ to effectively test the benefits of
anti-inflammatory drugs in mdx mice in vivo [Reviewed in (Grounds et al, 2008b)].
Numerous studies report the beneficial effects of anti-inflammatory therapies in
dystrophic mdx mice [Reviewed (Bogdanovich, 2004; Evans et al, 2009b; Manzur and
Muntoni, 2009; Radley et al, 2007; Spurney et al, 2009)], but this is the first study to
report the in vivo effects of a mouse specific antibody that targets TNF in dystrophic
mdx mice. TNF blockade may be a relatively expensive clinical treatment option for
DMD and the potential long-term side effects of TNF blockade in DMD patients is
unknown, although infliximab (Remicade®) is widely used clinically to treat other
human conditions in adults and children such as rheumatoid arthritis and Crohn’s
disease with much success and relatively mild side-effects (Kageyama et al, 2006). The
optimal regime and dosage, as well as the most appropriate humanised anti-TNF drug
needs to be selected for possible trials in DMD boys. The development of new TNF
drugs, such as TNF-TeAb (mini antibodies), further complicate the decision (Liu et al,
2007). We propose that TNF targeted therapies are more specific and may have fewer
side effects than corticosteroid treatment and it is surprising that clinical trials of TNF
based therapies have not yet been conducted.
Anti-oxidant therapy: The second drug treatment examined in this Thesis was the
antioxidant, N-acetylcysteine (NAC) (Chapter 5). The beneficial effects of NAC were
examined in both sedentary and treadmill exercised mdx mice, using the exact
experimental methods and analysis methods optimised in Chapter 4. NAC was chosen
as the antioxidant therapy, as it contains a thiol which can directly scavenge some
types of ROS directly (Zafarullah et al, 2003), it is also a precursor of L-cysteine which is
required in the synthesis of the major intracellular antioxidant glutathione (Dilger and
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___________________________________________________________Chapter 9 - Discussion
Baker, 2007), NAC can be administered orally, it is readily available and it is widely
used clinically. NAC has previously shown to improve muscle strength ex vivo and
reduce some aspects of dystropathology in sedentary mdx mice (Whitehead et al,
2008), and sedentary Stra13 knock-out mice (which exhibit a dystrophy-like pathology)
(Vercherat et al, 2009). To our knowledge this is the first study to extensively
demonstrate the in vivo benefits of NAC treatment on muscle histopathology, serum
CK level and oxidative state of both sedentary and exercised adult mdx mice.
NAC was administered via drinking water (1% or 4%) for a maximum duration of 6
weeks. NAC is reported to have an unpleasant bitter taste (Crouch et al, 2007;
Pendyala and Creaven, 1995) and administration of 4% NAC to mice via drinking water
resulted in a slight reduction in water intake and a slight reduction in body weight. For
this reason, it is important to consider an alternative method of delivery when
conducting long-term experiments with high concentrations of NAC. Miss Jessica
Terrill, a PhD student in our laboratory, is currently investigating appropriate
administration methods (e.g dietary supplement or gavage) and the best concentration
of NAC for therapeutic in vivo studies in mdx mice. The ‘bitter’ taste of NAC is less of a
factor when administering to human patients as it can be rationally tolerated or
administered in an enclosed capsule; however a bitter taste may influence the
compliance rate of treatment.
This work in Chapter 5 initially began as a comparison between the potential in vivo
benefits of NAC treatment vs. cV1q treatment in exercised adult mdx mice. It was
hypothesised that conducting two parallel in vivo experiments testing the effects of
both cV1q and NAC, (using identical experimental methods and analysis measures)
would enable identification of the more ‘promising’ therapeutic intervention.
Unfortunately, due to numerous problems with the cV1q experiments (e.g. concerns
regarding efficacy of different antibody batches and conflicting results), the cV1q work
was abandoned and only the NAC work was continued; therefore a direct comparison
between the in vivo effects of NAC vs. cV1q could not be made. However, when the
relative improvements seen in the two separate studies (Chapter 3 vs. Chapter 5) are
compared, both interventions appear to ameliorate dystropathology to a similar level.
For example, both 1% and 4% NAC treatment, reduced myofibre necrosis
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___________________________________________________________Chapter 9 - Discussion
approximately 3 fold (15% vs 6%) after 30 minutes of treadmill exercise and 4% NAC
treatment reduced blood serum CK approximately 3 fold (12 000U/L vs 32 000U/L)
after treadmill exercise. In comparison, cV1q treatment reduced myofibre damage (in
young 24 day old mice) approximately 2 fold (15% vs. 30%), reduced myofibre necrosis
after 48hrs voluntary exercise approximately 2 fold (7% vs 13%) and reduced serum CK
in long-term voluntarily exercised mice approximately 4 fold (4000U/L vs. 18000U/L)
(Radley et al, 2008).
Similar to the results seen with cV1q administration (Radley et al, 2008), differences in
the extent of dystropathology (and thus the effectiveness of NAC treatment) varied
between sedentary and exercised mdx mice, that is both 1% and 4% treatment showed
limited beneficial effects in sedentary adult mdx mice (Chapter 5). This again
demonstrates the importance of exercising adult mdx mice and confirms that exercise
should be part of the ‘standard operating procedure’ to effectively test the effects of
potential pre-clinical drugs in adult mdx mice in vivo [Reviewed in (Grounds et al,
2008b; Nagaraju and Willmann, 2009)]. NAC treatment showed many in vivo benefits
in mdx mice and our novel in vivo data supports the proposed clinical trials of NAC in
genetic myopathies. http://www.treat-nmd.eu/about/TACT/previous-reviews---june2010/
Dietary interventions: Previous studies in both DMD patients and mdx mice indicate
that the absence of dystrophin may also (directly or indirectly) lead to alterations in
skeletal muscle metabolism and an impaired energy status (Davidson and Truby, 2009;
Dupont-Versteegden et al, 1994a; Landisch et al, 2008; MacLennan and Edwards,
1990; Passaquin et al, 2002; Zanardi et al, 2003). In addition to these published data,
the striking differences in response to a high fat diet between mdx and C57Bl/10 mice
(identified in chapter 6) strongly indicating that dystrophic mdx mice have an ‘altered’
metabolic state.
Chapter 6 directly compared the response of control C57Bl/10 and dystrophic mdx
mice to a high protein (50%) and a high fat (16%) diet and identified striking
differences between strains in response to the high fat (16%). C57Bl/10 mice
demonstrated many negative side effects after consuming a high fat diet with
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___________________________________________________________Chapter 9 - Discussion
significant weight gain, increased body fat and elevation of pro-inflammatory
cytokines. In contrast, mdx mice (< 24 weeks) remained lean with minimal fat
deposition and were resistant to changes in body composition after consuming a high
fat diet. These results support the proposal that dystrophic mdx mice have an altered
‘energy status’ compared to normal C57Bl/10 mice since the mdx mice were more
capable of efficiently processing and utilising increased dietary fat. However, older
mdx mice (24-40 weeks old), where energy intake (kj/day and kj/g bw /day) was
significantly increased compared to mdx mice on the control diet, exhibited some
negative effects of a high fat diet (increase in both bodyweight and standardised fat
pad weight) but to a lesser extent than age matched C57Bl/10 mice. The negative
effects of a high fat diet seen in older mdx mice (24-40 weeks) also corresponds to a
very low level of myonecrosis and a cease in myofibre hypertrophy (indicative of
diminished new muscle formation and thus reduced additional energy demands due to
the status of the endogenous dystropathology). A high fat diet (and increased energy
intake) significantly increased the running ability (distance - km) of voluntarily
exercised mdx mice and significantly reduced myofibre necrosis in 24 week old
sedentary mdx mice, suggesting that increased dietary fat (and increased energy
intake) can assist energy deficient mdx mice metabolically, providing them with
additional energy to run further while maintaining muscle integrity. This supports the
notion of a high fat (or high energy) diet to reduce the severity of DMD, although this
must be balanced against the potential for obesity.
It must be noted that the custom diets used in this study (Chapter 6), high fat and high
protein, were initially designed to be isocaloric based on digestible energy content (HP
– 18.2 MJ/Kg vs. HF – 18.1 MJ/Kg). Major differences in body composition were seen in
response to the high fat and high protein diets and it became apparent that
metabolisable energy content was a more appropriate way to assess energy intake of
the mice. There are large differences in the metabolisable energy content of the high
fat and high protein diet (HP – 14.09 MJ/Kg vs. HF – 16.96 MJ/Kg) which explains the
major differences in energy intake and thus body composition in response to the two
diets. Therefore it is important to determine if the lack of benefits on dystropatholgy
with a high protein diet is due to diet composition (i.e. an increase in dietary protein
vs. Fat) or because consumption of the high protein diet did not result in an equivalent
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___________________________________________________________Chapter 9 - Discussion
increase in energy intake (kj/day). This question could be addressed by administering a
higher protein diet with a higher level of metabolisable energy and repeating the
analysis methods employed in Chapter 6: this additional experiment is now planned.
This is clearly of much interest since protein supplements are indeed used in an
attempt to maintain muscle mass in DMD boys, yet there is little conclusive evidence
for such nutritional supplements. It would seem that a high protein diet (with
increased energy intake) may be more desirable than a high fat diet as a source of
energy due to the potential adverse effects of a high fat diet.
Altered metabolism in mdx mice: The striking differences in response to a high fat diet
between mdx and C57Bl/10 mice (chapter 6) raised the question ‘Why don’t mdx mice
get fat after consuming a high fat diet?’ This led us to evaluate the differences in
metabolism, body composition and protein turnover in muscle between dystrophic
mdx and control C57Bl/10 mice, this research was conducted in collaboration with Dr
Marta Fiorotto at Baylor Medical college, Houston USA (Chapter 7).
Young mdx mice (~4 weeks) have a ‘stunted’ growth, increased protein synthesis rates
and increased metabolic rate and it appears that activity levels are reduced and energy
intake is increased in order to help to compensate for this. It is highly likely that the
acute onset of myofibre necrosis and initiation of new muscle formation during
regeneration at 3-4 weeks of age has a significant effect on metabolic rate in young
mdx mice and the onset of muscle hypertrophy, since growing new myofibres remain
highly responsive to growth factors, whereas mature myofibres are less responsive to
growth factors such as IGF-1 (Shavlakadze et al, 2010). There is significant ‘catch-up’
growth in mdx mice between 4 and 14 weeks of age, and 14 week old adult mdx mice
are heavier than C57Bl/10 mice, with significantly increase muscle mass and bone
length. Adult mdx mice are very lean, with significantly increased fat free mass (muscle
mass) and increased protein synthesis rates. Similar to the results seen in Chapter 6,
adult mdx mice appear to be focused on maintenance (and hypertrophy) of muscle
mass and show very little fat deposition. In addition to increased protein synthesis
rates and continuous cycles of myofibre necrosis and regeneration, dystrophic mdx
mice exhibit significant myofibre hypertrophy up to 24 weeks of age (chapter 6),
protein synthesis is a very energy expensive procedure and may explain why in mdx
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___________________________________________________________Chapter 9 - Discussion
mice (with the same increase in energy intake compared to C57Bl/10 mice) do not
exhibit the same negative side-effects of a high fat diet.
Significant differences in body composition (e.g. muscle mass and fat mass) between
young and adult mdx mice and between mdx and C57Bl/10 mice, may lead to changes
in the metabolism and clearing of various therapeutic drugs and thus, where possible,
dosage and half-life of different therapeutic interventions should be considered and
optimised for mdx mice of various ages (Hanley et al, 2010).
At this stage, it is still unclear if the metabolic differences between strains are an
innate feature of dystrophic muscle or a result of the continuous cycles of myofibre
necrosis and regeneration, and the high energy demands of growing myofibres in
dystrophic muscle. This question could be addressed by conducting similar metabolic
studies on young ‘pre-necrotic’ mdx mice (<20 days old). However, it must be noted
that conducting metabolic studies on young rapidly growing un-weaned mdx mice may
be complicated by dystrophic mothers being energy deficient themselves and this
impacting on milk availability for mdx pups, thus studies would need to be conducted
on cross-fostered (non-dystrophic mother) mdx litters. Alternatively, metabolic
analysis could be conducted in vitro on mature myotubes in primary muscle cultures
(Dystrophic vs. Control) since mdx myotubes do not manifest necrosis under routine
tissue culture conditions.
In collaboration with Dr Matt McDonagh (Department of Primary Industries, Victoria),
we are currently conducting a metabolomics screening study, using Nuclear Magnetic
Resonance (NMR) spectroscopy (Griffin and Des Rosiers, 2009; Jones et al, 2005), of
skeletal muscle metabolites in both pre-necrotic (15 day old litters) and adult (12 week
old adult) mdx and C57Bl/10 mice. Information gathered from this collaboration will
help to further understand possible subtle differences in metabolism between both
pre-necrotic and adult dystrophic and control mice.
Combined therapies: It seems likely that excess inflammation, elevated oxidative
stress and metabolic abnormality all contribute to myofibre necrosis via slightly
different mechanisms and that targeting one mechanism alone may only partially
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___________________________________________________________Chapter 9 - Discussion
ameloriate dystropathology, and may not prevent it completely. Although not
examined in this thesis, it is highly likely that combined therapeutic interventions (e.g.
an anti-TNF or antioxidant drug treatment + a high fat/ high protein diet) may have
cumulative beneficial effects in dystrophic muscle and should be pursued in pre-clinical
mdx studies. Inflammation and oxidative stress are tightly linked [Reviewed in (Arthur
et al, 2008; Tidball, 2005; Tidball and Wehling-Henricks, 2007)] and we have shown
that the antioxidant NAC can act to reduce both oxidative stress and inflammation in
dystrophic muscle (Chapter 5), therefore combined therapeutic intervention of both
an anti-inflammatory and an antioxidant drug might not result in any additional
benefit. However, combined therapy of a high fat diet (increased energy intake) with
an anti-inflammatory or antioxidant drug may have a cumulative benefit as the two
appear to be working through different mechanisms. Increasing numbers of recent
studies in mdx mice are trialling such combined therapies and demonstrate that
dietary interventions in combination with prednisolone have additive benefits to
reduce the severity of dystropathology (Cozzoli et al, 2010; Payne et al, 2006). A
deeper understanding of the precise mechanisms underlying the different aspects of
dystropathology, specifically further clarification of the enigma of metabolic
differences and benefits of different nutritional supplements, will greatly enhance the
optimal design of such combined therapies.
Development of a 30 minute treadmill protocol: Chapter 4 in this Thesis, focused on
the development of a short and repeatable in vivo treadmill exercise protocol. Biweekly treadmill exercise is widely used in pre-clinical experiments to increase the
extent of dystropathology in mdx mice (Burdi et al, 2009; De Luca et al, 2003;
Granchelli et al, 2000; Grounds et al, 2008b), yet the cellular consequences of a single
30 minute treadmill exercise session had not previously been described. The present
study (Chapter 4) standardised a protocol and conducted time-course analysis of
molecular and cellular changes after a single 30 minute treadmill exercise session. It
was concluded that a single 30 minute treadmill exercise session is a sufficient and
conveniently fast screening test to evaluate some of the benefits of pre-clinical drugs
in vivo. This has major applications to speed up pre-clinical drug screening in mdx mice.
This study also showed that myofibre necrosis, blood serum CK and oxidative stress
(specifically the ratio of oxidised to reduced protein thiols) are reliable markers of
307
___________________________________________________________Chapter 9 - Discussion
muscle damage after such treadmill exercise; however sampling time (after exercise)
for these parameters is critical. This study further emphasises the need for standard
operating procedures in order to compare experimental data between different
laboratories. The methods developed in the study were employed when assessing the
beneficial effects of NAC in vivo (Chapter 5).
While Chapter 4 identified 30 minutes of treadmill exercise to be an efficient screening
method to assess potential therapeutic drugs in vivo, there are also many benefits of
using voluntary exercise. For example, distance run (km) is a valuable indicator (albeit
indirect) of exercise capacity and thus muscle function after therapeutic intervention
(Call et al, 2008 11905; Radley et al, 2008), voluntary exercise also allows the mice to
run at night during their normal ‘active’ hours and is thus less stressful (Grounds et al,
2008b; Radley and Grounds, 2006). The whole process does not require intensive
researcher involvement and is easily conducted over many months. Further discussion
of the advantages and limitations of both exercise methods is covered in section 1.3.2
of this Thesis.
The TNF antibody, cV1q (Chapter 3) (Radley et al, 2008) and the high fat and high
protein dietary interventions (Chapter 6) were assessed in voluntarily exercised adult
mdx mice. One reason for using voluntary exercise in these 2 studies is that they were
conducted prior to the availability of a rodent treadmill in our laboratory group, plus
the voluntary running is especially convenient for long term studies. It is striking that a
single 30 minute bout of treadmill exercise resulted in a similar increase in myofibre
necrosis to that seen with 48hrs voluntary exercise, endorsing a similar muscle
susceptibility to both types of exercise.
Specific exercise regime and gender
Myofibre necrosis in the quadriceps (% CSA)
48 hours voluntary exercise (F)
12.93% ( /- 3.08)
8 weeks voluntary exercise (F)
10.19% ( /- 1.96)
30 minutes treadmill exercise (24hrs post) (M)
15.06% ( /- 6.01)
4 weeks treadmill exercise (M)
11.83% ( /- 2.22)
+
+
+
+
Table 9.1 Myofibre necrosis after exercise in mdx mice. Myofibre necrosis in the quadriceps muscle is
similar across the 4 exercise protocols used in this Thesis.
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___________________________________________________________Chapter 9 - Discussion
Indeed, it is very interesting to note that the amount of myofibre necrosis (% crosssectional area) in the quadriceps muscle of exercised adult (6-12 weeks) mdx mice is
similar after all 4 types of exercise used in this Thesis (Table 9.1)(Radley et al, 2008)
and did not differ between male and female mice. This suggests that there is a certain
number of myofibres at any one time that are ‘vulnerable’ to exercise induced damage
and that increasing the duration of exercise (e.g increasing voluntary exercise from
48hrs up to 8 weeks) or increasing the number of exercise sessions a mouse is required
to complete (e.g. a single 30 minute session vs. 8x 30 minute sessions) does not
increase the percentage of myofibre necrosis. These data also demonstrates that
dystrophic myofibres which undergo necrosis after contraction induced damage are
fully capable of regenerating between exercise bouts (Chapter 4).
It is possible that if the ‘severity’ of the exercise protocol is increased (e.g. eccentric
downhill running) that even more myofibre necrosis would be seen. While myofibre
necrosis in the quadriceps muscle after eccentric exercise does not seem to be
reported in the literature, higher levels of myofibre damage after eccentric downhill
running for other muscles is described, for example: 3 downhill running sessions (15º
decline) for 10 minutes at a speed of 10m/minute, caused 22.92% ( +/- 1.08) damage in
the triceps and 23.83% (+/- 1.18) damage in the gastrocnemius, as measured by Evans
Blue Dye positive myofibres (Brussee et al, 1997). It is noted that Evans Blue Dye
identifies ‘myofibre leakiness’ which does not always correspond to myofibre necrosis
(Archer et al, 2006; Grounds et al, 2008b; Hamer et al, 2002; Straub et al, 1997) but
these data do support higher levels of myofibre damage after more severe types of
exercise.
Variation in mdx mouse: A high level of variation in the extent of dystropathology in
mdx mice (within and between both individual mice and whole litters) is identified
throughout this thesis (Chapter 3 & 4). This is an extremely important point to consider
when conducting pre-clinical studies in mdx mice and treatment groups should always
consist of age, sex and weight (where possible) matched mdx mice (Grounds et al,
2008b)(Willmann et al, in preparation). Analysis (histological or molecular) should also
be conducted on the same muscle (or muscle group), since the extent of
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___________________________________________________________Chapter 9 - Discussion
dystropathology can also vary between muscles (Chapter 4) (Grounds et al, 2008b;
Radley et al, 2008).
A clear example of variation is documented in Chapter 4. Numerous experiments
(control groups from 9 separate experiments) conducted in our laboratory revealed
high levels of variation in the extent of dystropathology in various muscles of
unexercised adult mdx mice. The variation in myofibre necrosis (% CSA) in the
quadriceps muscle from 12 week old sedentary male mdx mice ranges from 1.04 –
23.1% for individual muscles (Chapter 4). When results from the 9 experiments were
pooled together (n=60 quadriceps) the average amount of myofibre necrosis in the
quadriceps muscle of a 12 week old unexercised male mdx mouse is 6.12% (Chapter 4).
Similar variation was seen in the triceps and gastrocnemius muscles. These ‘pooled’
histological data were used in the assessment of myofibre necrosis after treadmill
exercise (Chapter 4) and to assess the beneficial effects of NAC treatment (Chapter 5).
Standard Operating Procedures: Recently, the important issue of world-wide standard
operating procedures for pre-clinical studies in the mdx mouse has been heavily
discussed (Grounds et al, 2008b; Nagaraju and Willmann, 2009) and researchers have
begun establishing a set of recommended standard operating procedures and
endpoints for consistent and comparable assessment of pre-clinical interventions in
the mdx mouse (Grounds et al, 2008b; Guerron et al, 2010; Spurney et al, 2010;
Spurney et al, 2009)(Radley-Crabb et al, submitted; Willmann et al, in preparation).
There are also many standard operating procedures listed on the TREAT-NMD website
http://www.treat-nmd.eu/research/preclinical/SOPs
and
where
possible
these
guidelines have been followed throughout this Thesis (Chapter 4 & 5). Included in this
thesis (Methods – Chapter 2) is a standard operating procedure for ‘Haematoxylin and
eosin staining for histological analysis of dystrophic muscle’. This method is published
on the TREAT-NMD website and available worldwide for all researchers to follow and
discuss.
Conclusion: The mdx mouse is a very useful animal model to examine the potential
mechanisms by which the absence of dystrophin leads to myofibre necrosis and to test
potential therapeutic interventions to reduce the extent of dystropathology in vivo.
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___________________________________________________________Chapter 9 - Discussion
The exact mechanism by which the absence of dystrophin leads to myofibre necrosis is
unknown, however the body of work presented in this thesis confirms the involvement
of inflammation, oxidative stress and metabolic abnormality and that therapeutic
interventions targeting these 3 processes can ameliorate dystropathology. It seems
likely that myofibre necrosis is due to a combination of many dysregulated processes
and combined pre-clinical therapeutic interventions should be pursued. The high level
of variation in extent of dystropathology in mdx mice emphasises the need for
establishing Standard Operating Procedures to enable comparison of data between
laboratory groups world-wide when designing and conducting pre-clinical in vivo
therapeutic trials in mdx mice.
311
_________________________________________________________Chapter 10 - References
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The International Journal of Biochemistry & Cell Biology 39 (2007) 469–477
Medicine in focus
Duchenne muscular dystrophy: Focus on pharmaceutical
and nutritional interventions
H.G. Radley a , A. De Luca b , G.S. Lynch c , M.D. Grounds a,∗
a
c
School of Anatomy and Human Biology, University of Western Australia, Crawley, Australia
b Unit of Pharmacology, Department of Pharmacobiology, University of Bari, Bari, Italy
Basic and Clinical Myology Laboratory, Department of Physiology, University of Melbourne, Parkville, Australia
Received 4 July 2006; received in revised form 8 September 2006; accepted 29 September 2006
Available online 10 October 2006
Abstract
Duchenne muscular dystrophy is a lethal X-linked muscle disease resulting from a defect in the muscle membrane protein dystrophin. The absence of dystrophin leads to muscle membrane fragility, muscle death (necrosis) and eventual replacement of skeletal
muscle by fat and fibrous connective tissue. Extensive muscle wasting and respiratory failure results in premature death often by the
early 20s. This short review evaluates drug and nutritional interventions designed to reduce the severity of muscular dystrophy, while
awaiting the outcome of research into therapies to correct the fundamental gene defect. Combinations of dietary supplementation
with amino-acids such as creatine, specific anti-inflammatory drugs and perhaps drugs that target ion channels might have immediate
realistic clinical benefits although rigorous research is required to determine optimal combinations of such interventions.
© 2006 Elsevier Ltd. All rights reserved.
Keywords: Duchenne muscular dystrophy; Mdx mouse; Pharmaceuticals; Nutritional supplements; Therapy
1. Duchenne muscular dystrophy and the mdx
mouse model of DMD
There are many forms of muscular dystrophy
but only Duchenne muscular dystrophy (DMD) is
discussed here. DMD is an X-linked lethal muscle
wasting disorder, affecting approximately 1/3500 male
births. The disease is caused by a mutation in the
gene that encodes for the sub-sarcolemmal protein
dystrophin (Biggar, 2006). Dystrophin links the muscle
cytoskeleton through a membrane complex to the extracellular matrix. Dystrophic myofibres are susceptible to
∗ Corresponding author. Tel.: +61 8 6488 3486;
fax: +61 8 6488 3486.
E-mail address: [email protected] (M.D. Grounds).
1357-2725/$ – see front matter © 2006 Elsevier Ltd. All rights reserved.
doi:10.1016/j.biocel.2006.09.009
damage during mechanical contraction, damage leads
to myofibre necrosis and ultimately the replacement of
myofibres by fibrous and fatty connective tissue (due
to failed regeneration). While the genetic defect was
identified in 1987, the specific mechanism of myofibre
damage is still unclear (Whitehead, Yeung, & Allen,
2006) and there is still no effective treatment for DMD.
Therapeutic approaches for DMD fall into three main
strategies: (i) replacement of dystrophin by genetic, cell
transplantation or molecular interventions; (ii) enhancement of muscle regeneration or reduction of fibrosis
to combat failed regeneration; (iii) reduced muscle
necrosis. This latter approach is the main focus of this
review which outlines pharmacological interventions
and nutritional supplementation as potential therapies to
reduce myofibre necrosis in DMD. Most experimental
studies use mdx mice and therefore data from this
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H.G. Radley et al. / The International Journal of Biochemistry & Cell Biology 39 (2007) 469–477
model, along with clinical studies, form the basis of
this review. DMD primarily affects skeletal and cardiac
muscle and in addition other tissues (Biggar, 2006), but
only the effects on skeletal muscle will be addressed.
The mdx mouse is the most widely used animal model
for DMD. The absence of dystrophin results in a distinct
disease progression with an acute onset of skeletal muscle necrosis around 3 weeks of age in young mdx mice
(Fig. 1), necrosis then decreases significantly after 4–6
weeks to a relatively low level in adult mice (McGeachie,
Grounds, Partridge, & Morgan, 1993): the pathology
is far more benign than in DMD. The acute onset of
myofibre necrosis provides an excellent model to study
therapeutic interventions to prevent or reduce necrosis.
In contrast, reduced necrosis can be difficult to detect in
adult mice where there is little active myofibre breakdown but high cumulative muscle pathology. For this
reason, exercise is often used to induce muscle damage
Fig. 1. Skeletal muscle necrosis in the mdx mouse. (A) Pre-necrotic
(normal) skeletal muscle in the quadriceps muscle of a 20-day-old
mdx mouse, characterized by healthy myofibres with peripheral nuclei
(PN). (B) Necrotic skeletal muscle in the quadriceps muscle of a 23day-old mdx mouse, characterized by infiltrating inflammatory cells
(IC) and degenerating myofibres (DM). Transverse muscle sections
stained with haematoxylin and eosin. Scale bar represents 50 ␮m.
enabling potential therapeutic interventions to be evaluated in adult mdx mice (Granchelli, Pollina, & Hudecki,
2000; Payne et al., 2006). The simplest form of exercise
is voluntary wheel running, where muscle necrosis in
the quadriceps is doubled (from ∼6 to 12%) after 48 h
(Hodgetts, Radley, Davies, & Grounds, 2006; Radley &
Grounds, 2006). Forced exercise greatly increases muscle damage with the most severe injury resulting from
forced downhill running (eccentric exercise), although
such severe muscle damage caused by eccentric exercise is a poor model for pre-clinical drug screening.
The symptoms of dystropathology are cumulative, with
fibrosis becoming increasingly pronounced in older (>15
months) mdx mice. Symptoms are most severe in the
mdx diaphragm that more closely resembles the severe
pathology of DMD (Stedman et al., 1991).
Numerous parameters are measured to assess the in
vivo effects of various interventions. In mdx mice, measurements on whole animals are combined with extensive tissue analysis. Physiological parameters such as
Rotarod tests (to test motor co-ordination and fatigue
resistance) and grip-strength tests (to measure the maximum amount of force an animal applies by grasping)
assess changes in muscle endurance, muscle strength
and overall functional capacity. Further physiological
tests are conducted in vivo and on isolated muscles
in situ or in vitro. Blood sampling and serum creatine
kinase (CK) levels provide a qualitative indicator of muscle damage. Histological assessment of tissue sections
quantifies cumulative muscle necrosis and regeneration,
along with leaky myofibres and immunohistochemical
staining identifies changes in location and levels of specific proteins. Other measurements include alterations
in channel (Ca2+ and Cl− ) function that contribute (or
sensitise) to disrupted calcium homeostasis and to muscle necrosis (De Luca et al., 2003). A positive result
with mdx mice can eventually lead to clinical studies
in DMD patients. In humans, the main parameters measured are muscle strength, functional tests and CK levels.
The Cooperative International Neuromuscular Research
Group (CINRG) performs clinical trials on young DMD
patients with various compounds, some of which showed
positive results in an early screening program on dystrophic mice (Granchelli et al., 2000): while some of
these trials have been published, ongoing results are
available on http://www.cinrgresearch.org.
2. Steroids and anti-inflammatory drugs
Until a cure for DMD is found, treatment will involve
the administration of corticosteroids combined with
interventions to alleviate cardiac and respiratory prob-
H.G. Radley et al. / The International Journal of Biochemistry & Cell Biology 39 (2007) 469–477
lems. Corticosteroids have a catabolic effect on muscle
(non-exercised muscle) and act to preserve existing
muscle fibres and reduce inflammation, although their
exact mechanism of action in dystrophic skeletal muscle
is unknown. The two main corticosteroids used to
treat DMD, Prednisone and Deflazacort, both seem
equally effective in delaying the progression of muscle
wasting. Unfortunately both are associated with adverse
side-effects although these, particularly weight gain, are
less severe with Deflazacort. Side-effects of Deflazacort
include an increase in appetite requiring strict dietary
control, retarded growth resulting in short stature and
asymptomatic cataracts. It is recommended that daily
calcium and Vitamin D supplementation is taken in conjunction with corticosteroids to maintain bone mineral
density and reduce bone fractures (Biggar, 2006). The
administration of steroids is not a cure for DMD but a
therapy to improve quality of life and prolong lifespan.
DMD is characterized by aggressive inflammation
and there is strong evidence that this contributes to
myofibre necrosis both in vitro and in vivo (reviewed
in Refs. (Tidball & Wehling-Henricks, 2005)). Other
immunosuppressive drugs have demonstrated benefits
in mdx mice, leading to increasing recognition for the
damaging role of inflammation in DMD. Early clinical
trials with the immunosuppressive drug cyclosporine in
DMD returned promising results as 8 weeks of treatment
(5 mg/kg/day) resulted in a significant increase in muscle
force generation (Sharma, Mynhier, & Miller, 1993) and
cyclosporine reduced the dystropathology in mdx mice
(De Luca et al., 2005). However, cyclosporine exerts
multiple dose-dependent effects and a correct dosage
must be established especially when administered to
young dystrophic patients during muscle development.
These results are of clinical relevance, as immunosuppression may be required for enhancing efficiency of
future gene/cell therapies.
Another potential anti-inflammatory drug that has
attracted attention is pentoxifylline. Pentoxifylline has
a wide range of anti-inflammatory and anti-coagulant
effects; it reduces TNF␣ production in vitro (Vary et
al., 1999), reduces fibrosis and may also play a role in
normalising blood flow in dystrophic muscle. On the
basis of increased muscle strength in exercised adult
mdx mice (Granchelli et al., 2000), pentoxifylline is now
the subject of two CINRG trials in DMD patients (one
completed recently). However, a recent study in mdx
mice involving pentoxifylline (16 mg/kg/day) administration for 4 weeks failed to reduce fibrosis or improve
the contractile force of the diaphragm muscle (Gosselin
& Williams, 2006), this study does not support the use of
pentoxifylline as an anti-fibrotic drug in DMD patients.
471
However, pentoxifylline counteracts, both in vitro and in
vivo, the abnormal activity of calcium channels responsible for high sarcolemmal calcium permeability of dystrophic myofibres, suggesting a possible amelioration
of dystrophic condition through alternative pathways
(Rolland et al., 2006). Both cyclosporine and pentoxifylline (like corticosteroids) affect many cellular events
and can have severe adverse side-effects, careful animal
studies will be helpful in this regard.
Other anti-inflammatory drugs such as oxatomide
(Granchelli et al., 2000) and cromolyn (Granchelli,
Avosso, Hudecki, & Pollina, 1996; Radley & Grounds,
2006) that block mast cell degranulation are used widely
for clinical treatment of allergies such as asthma and
show benefits in mdx mice, indicating that mast cell
products (including TNF␣) have detrimental effects in
dystrophic muscle. A recent CINRG trial with oxatomide (based on the study by (Granchelli et al., 2000)) in
DMD showed some minor benefits.
3. Specific anti-cytokine drugs
Another promising approach involves targeting specific aspects of the inflammatory response (rather than
the broadly acting anti-inflammatory drugs) in order
to reduce muscle necrosis. While systemic depletion
of specific inflammatory cells may not be clinically
viable, modulation of specific cytokines has been very
successful clinically in several severe inflammatory disorders. tumour necrosis factor-alpha (TNF␣) is a key
pro-inflammatory cytokine that stimulates the inflammatory response and pharmacological blockade of TNF␣
activity with the neutralising antibody infliximab (Remicade) is highly effective clinically at reducing symptoms
of inflammatory diseases such as rheumatoid arthritis
and Crohn’s disease (Feldman & Maini, 2003). Similar
successful clinical blockade of TNF␣ by the drug etanercept (Enbrel) results from the use of soluble receptors
to TNF␣. In mdx mice, infliximab delays and reduces
the necrosis of dystrophic muscle in young mdx mice
(Grounds & Torrisi, 2004). A protective effect of TNF␣
blockade is reinforced by two recent studies using etanercept that clearly reduces muscle necrosis in young mdx
mice (Hodgetts et al., 2006; Pierno et al., 2006) and in
exercised adult mdx mice (Hodgetts et al., 2006) with
additional physiological benefits on muscle strength,
chloride channel function and reduced CK levels being
demonstrated in chronically treated exercised adult mdx
mice (Pierno et al., 2006). Such emerging highly specific anti-inflammatory drugs designed for use in other
clinical conditions, appear an attractive alternative (to
steroids) for DMD, although their potential to reduce
472
H.G. Radley et al. / The International Journal of Biochemistry & Cell Biology 39 (2007) 469–477
the severity of DMD remains to be determined. It may
be possible to limit the use of these drugs to periods of
intensive muscle growth in boys when muscle damage
and deterioration can be especially pronounced. Patients
undergoing long-term anti-cytokine treatment must be
monitored carefully for serious infections, as is the case
for any immunosuppressive drug.
4. Other pharmaceutical interventions
Antioxidants. It is widely recognized that high levels
of reactive oxygen species can damage tissues, including skeletal muscle (Rando, 2002). Antioxidants that
reduce oxidative damage in cells, such as Coenzyme Q10
(CoQ10) and green tea extract [(−)-epigallocatechin gallate] are the subject of recent research in mdx mice and
DMD patients. Green tea extract supplemented diets fed
to mdx mice (from birth), significantly reduced muscle
damage (necrosis and regeneration) in the EDL muscle
of 4-week-old mice and improved muscle function in
8-week-old mice after 5 weeks of treatment (Buetler,
Renard, Offord, Schneider, & Ruegg, 2002; Dorchies
et al., 2006). CoQ10 is essential for several enzymatic
steps in the production of energy and functions as
an antioxidant. CoQ10 was the subject of a CINRG
pilot study in DMD patients to assess the effectiveness
and safety in combination with steroid treatment.
CoQ10 increased strength in some muscle groups and
a larger follow-up study was recommended. This larger
study that is currently being conducted by CINRG,
is a 13-month, prospective, randomized study comparing daily Prednisone treatment (0.75 mg/kg/day),
CoQ10 (>2.5 ␮g/mL) and a combined treatment
(Prednisone and CoQ10) in older non-ambulatory
patients.
4.1. Anabolic effects of β2 -agonist drugs
Anabolic agents result in a net increase in protein content and muscle size and this is usually (but not always)
associated with increased strength. Although commonly
recognized as asthma drugs, high doses of some ␤2 agonists have anabolic effects on muscle and thus the
potential to slow muscle degeneration. A 3-month pilot
trial of the ␤2 -agonist albuterol given to patients with
fascioscapulohumeral disease improved maximum voluntary strength. This was followed by a year long trial
where patients were treated with up to 16 mg of albuterol
twice daily resulting in improved muscle mass and grip
strength. Albuterol administered for 28 weeks (Fowler,
Graves, Wetzel, & Spencer, 2004) to boys with DMD
produced a modest increase in strength with no reported
side-effects. It is noted that studies with ␤2 -agonists in
mdx mice have returned inconsistent results (DupontVersteegden, Katz, & McCarter, 1995; Lynch, Hinkle, &
Faulkner, 2000) and that ␤2 -agonists are associated with
numerous undesirable side-effects including increased
heart rate and tremors, which have limited their therapeutic potential. More recently synthesized ␤-agonists such
as formoterol, have anabolic effects on skeletal muscle with minimal cardiac side-effects (Ryall, Sillence, &
Lynch, 2006) when administered at micro-molar doses
in mice and thus may offer a greater potential for DMD
(Harcourt, Schertzer, Ryall, & Lynch, 2006).
4.2. Disturbed ion channels and drugs to inhibit
proteases
Damaged muscle membranes disturb the passage of
calcium ions into the myofibre, and disrupted calcium
homeostasis activates many enzymes, e.g. proteases, that
cause additional damage and muscle necrosis. Ion channels that directly contribute to the pathological accumulation of calcium in dystrophic muscle are potential targets for drugs to treat DMD. There is evidence
that some drugs, such as pentoxifylline, block exercisesensitive calcium channels (Rolland et al., 2006) and
antibiotics that block stretch activated channels reduce
myofibre necrosis in mdx mice and CK levels in DMD
boys (Whitehead et al., 2006). Calpains are calcium
activated proteases that are increased in dystrophic muscle and may directly account for myofibre degeneration
(Spencer, Croall, & Tidball, 1995). A new compound,
BN 82270 (Ipsen) that has dual action as both a calpain inhibitor and an antioxidant (targeting both calpain
and ROS induced muscle damage) increased muscle
strength, decreased serum CK and reduced fibrosis of
the mdx diaphragm, suggesting a potential therapeutic
effect with this new compound (Burdi et al., 2006). A
promising new compound of Leupeptin/Carnitine (Myodur) has recently been proposed for clinical trials in
DMD patients.
Beyond the diverse approaches mentioned already,
there has been much research into substances to help
reduce the dystropathology via maintenance or improvement of myofibre size, strength and function. However,
strategies such as increasing IGF-1 or other growth factors, inhibition of myostatin, normalising nitric oxide
production and the use of poloxamer (P188), do not
readily translate into the clinical situation at present.
Although these interventions (and many others) have
shown promising effects in mdx mice, adverse or
unknown long-term systemic effects currently limit their
therapeutic potential.
H.G. Radley et al. / The International Journal of Biochemistry & Cell Biology 39 (2007) 469–477
Chinese herbal medicine is becoming increasingly
popular as an alternative approach to reduce the severity of symptoms associated with many diseases. For
example, ginseng has a diverse range of effects in vivo
and a study that showed a reduction in exercise-induced
damage in normal muscle after ginseng supplementation (Hsu, Ho, Lin, Su, & Hsu, 2005) suggests possible benefits for DMD. A review of traditional Chinese
medicines, such as massage, acupuncture and capsules
(that contained herbs and other ingredients) that claimed
to alleviate symptoms in DMD patients, was conducted
in Beijing in 2003 (Urtizberea, Fan, Vroom, Recan, &
Kaplan, 2003). Due to the small number of cases no
definitive conclusions could be drawn from this study,
although an overall mild frequency of contractures was
noted and related possibly to the positive influence of
acupuncture and massage. A follow-up study confirmed
that high levels of glucocorticoids were present in the
capsules and may account for the anecdotal improvement
seen in patients (Courdier-Fruh, Barman, Wettstein, &
Meier, 2003). Chinese herbal medicine was also found
to improve locomotor activity in mdx mice (Chen, 2001).
Traditional Chinese medicine is not formally regulated
which potentially creates a large risk for incorrect dosing
and dangerous content as medicines may unknowingly
contain various heavy metals, herbicides, drugs, pesticides and micro-organisms. Furthermore, adverse drug
interactions with cumulative effects may result when
administered to DMD boys already receiving steroid
treatment.
5. Nutritional interventions
Deficiencies of many substances (Selenium, Vitamin
E or Vitamin D) can cause severe myopathies, suggesting
the importance of nutritional supplementation to counteract these deficiencies: however it seems difficult to
justify the application of such supplements to situations
of adequate diet or DMD. Muscle wasting is associated
with changes in the biochemistry of skeletal muscle
that lead to reduced protein synthesis, increased protein
breakdown (catabolism) and increased oxidative cell
damage. The use of protein powders and specific amino
acid supplements has been proposed for attenuating muscle protein loss and to provide a favourable environment
for increasing protein synthesis and muscle mass and has
received much attention in sports medicine and ageing.
Dietary supplementation is a form of protective therapy
potentially available for immediate use and broadly
falls into three categories; antioxidants or Chinese herbs
(both discussed above) and amino acid supplementation.
473
5.1. Amino acids
Amino acids such as creatine, taurine, glutamine and
l-arginine have all been trialed in the mdx mouse with
some benefits on muscle strength or dystropathology.
Creatine is directly involved in the energy supply of
muscle cells and supplementation into the diet (10%,
w/w, in chow) of both exercised adult mdx mice and
pregnant mdx mothers increased strength and improved
dystropathology (De Luca et al., 2003; Passaquin et al.,
2002). Similar benefits were seen after intraperitoneal
injection of 10mg/kg in exercised adult mice (Granchelli
et al., 2000). A double-blinded randomized creatine
monohydrate (0.10 g/kg/day) trial in boys with DMD for
4 months showed increased hand grip strength and fatfree mass, independent of steroid usage (Tarnopolsky
et al., 2004). A more recent clinical trial conducted by
CINRG tested two amino acids (creatine 5g/day and
glutamine 0.6 g/kg/day) in DMD boys aged 4–10 years
(grouped as 4–7 years and 7–10 years). Although it
did not significantly improve muscle strength, creatine
was well tolerated by all patients and there was a trend
towards less deterioration in other outcomes (Escolar
et al., 2005). Creatine monohydrate supplementation in
DMD is the subject of a recent review (Pearlman &
Fielding, 2006) that concludes that it should be considered as a therapeutic agent for DMD, due to the
potential for increased fat-free mass and increased muscle strength. However, additional long-term studies are
required to elucidate the role of creatine in skeletal muscle growth and to accurately assess the degree to which
creatine exerts protective musculoskeletal effects, without unwanted side-effects such as weight gain and kidney
problems.
Taurine is a free amino acid abundant in skeletal
muscle that exerts a wide spectrum of actions, ranging from osmolyte control, antioxidant action and antiinflammatory effects. In skeletal muscle, taurine modulates ion channel function and calcium homeostasis
(Conte Camerino et al., 2004). Supplementation of taurine (10%, w/w, in chow) is relatively safe and counteracts exercise-induced weakness after chronic exercise
and ameliorates gCL (macroscopic chloride conductance, an index of degeneration-regeneration) in EDL
muscles of mdx mice (De Luca et al., 2003). Clinically,
taurine has been used with varying degrees of success
in a wide variety of conditions (Birdsall, 1998) and is
present in commercial food and beverages claimed to
work as energizers.
Screening of numerous amino acids for potential efficacy in the mdx mouse (Granchelli et al., 2000) found
that intraperitoneal injections of glutamine (10 mg/kg)
474
Table 1
Summary of potential pharmaceutical and nutritional interventions as therapeutic agents in (mdx mice and) DMD patients
Action
Mdx mice
DMD patients (+/− benefit)
Immediate clinical
potential
Cyclosporine
Immunosuppressant,
anti-inflammatory
Pentoxifylline
Anti-inflammatory,
anti-coagulant, anti-fibrotic
Eight-week trial—increased
muscle force generation
(Sharma et al., 1993)
2×CINRG recent clinical
trials (one trial on-going)
May be (dosage to be
carefully decided due to
potential toxicity)
May be
Oxatomide
Anti-inflammatory histamine
(H1) receptor antagonist
(asthma therapy)
Anti-inflammatory mast cell
stabilizer (asthma therapy)
Maintained muscle strength, cellular
parameters & CK levels in exercised adult
mice (De Luca et al., 2005)
Increased muscle strength in exercised mice
(Granchelli et al., 2000; Rolland et al., 2006).
No improvement in mdx diaphragm fibrosis
(Gosselin & Williams, 2006) amelioration of
calcium homeostasis (Rolland et al., 2006).
Increased muscle strength in exercised mice
(Granchelli et al., 2000)
Recently completed CINRG
clinical trial
Yes
Increased strength and reduce muscle
necrosis in young and adult mice (Radley &
Grounds, 2006)
Reduced muscle necrosis in young mice
(Grounds & Torrisi, 2004)
–
Yes
–
Yes (Monitor for possible
infections)
Reduced muscle necrosis in young and
exercised adult mice (Hodgetts et al., 2006;
Pierno et al., 2006). Maintained muscle
strength, cellular parameters & CK levels in
exercised adult mice (Pierno et al., 2006).
–
–
Yes (monitor for possible
infections)
2× CINRG recent clinical
trials (one trial on-going)
–
Yes
Anecdotal improvement in
patients (Urtizberea et al.,
2003) Steroid content
confirmed (Courdier-Fruh et
al., 2003)
Albuterol, 28 week trial
resulted in strength increase
(Fowler et al., 2004)
–
No (exact composition of
tablets unknown)
Cromolyn
Infliximab (Remicade)
Etanercept (Enbrel)
Coenzyme Q10
Anti-inflammatory TNF␣
antibody (arthritis, Crohn’s
disease therapy)
Anti-inflammatory TNF␣
antibody (arthritis, Crohn’s
disease therapy)
Green tea extract
Antioxidant energy
production
Antioxidant
Chinese herbal medicine
Antioxidant
Old ␤-agonists: clenbuterol,
albuterol
Anabolic effects on muscle.
Possible anti-inflammatory
(asthma therapy)
Anabolic effects on muscle.
Possible anti-inflammatory
New ␤-agonists: formoterol
Reduced necrosis and improve muscle
function in mice (Buetler et al., 2002;
Dorchies et al., 2006)
Improved locomotor activity in adult mdx
mice (Chen, 2001)
Inconsistent results in mdx mice
(Dupont-Versteegden et al., 1995; Lynch et
al., 2000)
Improved muscle function in mdx mice
(Harcourt et al., 2006)
Yes (necessary dosage
still to be determined)
May be (potential for
cardiac problems)
May be/yes (minimal
cardiac side-effects)
H.G. Radley et al. / The International Journal of Biochemistry & Cell Biology 39 (2007) 469–477
Substance
Substance
Action
Mdx mice
DMD patients (+/− benefit)
Immediate
potential
Calpain inhibitors: BN 82270,
Myodur
Inhibit calcium dependent
enzymes with additional
pharmacodynamic or
pharmacokinetic properties
Amino acid (directly involved
in muscle metabolism)
BN 82270, increased muscle strength,
decrease CK and muscle fibrosis in mice
(Burdi et al., 2006)
Myodur proposed for clinical
trials in 2006.
May be (pre-clinical
toxicological studies are
required)
Increased strength and improved
dystropathology in mice (De Luca et al.,
2003; Granchelli et al., 2000; Passaquin et
al., 2002)
Four-month trial—increased
grip strength and decrease fat
mass (Tarnopolsky et al.,
2004) 6 month CINRG
clinical trial—reduced
deterioration of strength
(Escolar et al., 2005)
–
Yes (upon monitoring of
possible side-effects)
Creatine
Taurine
Amino acid (control of
calcium handling in vitro)
Amino acid
Maintained strength in exercised adult mice
(De Luca et al., 2003)
Maintained strength in exercised adult mice
(Granchelli et al., 2000)
l-Arginine
Amino acid (substrate for
NOS)
Chinese massage or
acupuncture
Stimulation of skeletal
muscles and joints
Upregulation of utrophin, improved
dystropathology and reduced exercise
induced muscle damage (Archer et al., 2006;
Voisin et al., 2005)
–
Glutamine
CINRG clinical trial (Escolar
et al., 2005) 10-day
trial—reduced whole-body
protein degradation (Mok et
al., 2006)
–
Anecdotal improvements in
patients (Urtizberea et al.,
2003)
clinical
Yes (upon monitoring of
possible side-effects)
Yes (upon monitoring of
possible side-effects)
Yes (upon monitoring of
possible side-effects)
Yes
Note: Even though some substances are shown as having the potential for immediate clinical intervention, the issue of adverse side-effects requires careful evaluation and, importantly, some of
these substances when administered alone show only marginal benefits.
H.G. Radley et al. / The International Journal of Biochemistry & Cell Biology 39 (2007) 469–477
Table 1 (Continued )
475
476
H.G. Radley et al. / The International Journal of Biochemistry & Cell Biology 39 (2007) 469–477
and a glutamine and alanine combination (10 mg/kg
each) significantly improved whole body strength after
6 weeks of treadmill exercise. Clinical studies with glutamine show that oral supplementation (0.5 g/kg/day)
over 10 days also inhibits whole-body protein degradation in DMD patients (Mok et al., 2006). Combinations
of multiple dietary supplements (including creatine),
both with and without prednisolone have recently shown
improved muscle strength and reduced fatigue in exercised mdx mice (Payne et al., 2006). Such studies build
a good case for promising benefits from combined interventions.
An additional possible treatment for DMD would
be to compensate for the loss of dystrophin and nitric
oxide (NO) with pharmacological agents. Administration of l-arginine (the substrate for nitric oxide synthase)
increases NO production and up regulates utrophin
expression in mdx mice. Six weeks of l-arginine treatment (200 mg/kg – intraperitoneal injection) improved
muscle dystropathology and decreased serum CK in mdx
mice (Voisin et al., 2005) and when given in combination
with Deflazacort (Archer, Vargas, & Anderson, 2006) larginine (0.375% in drinking water) spared limb muscles
from exercise induced damage and increased the distance (km) run voluntarily by an individual mouse. Since
l-arginine can have adverse side effects, the drug isosorbide dinitrate that increases NO might be preferable
for NO-based therapy in muscular dystrophy (Marques,
Luz, Minatel, & Neto, 2005).
While amino acids have been proposed for many
clinical conditions, possible benefits to DMD of supplementation with taurine, glutamine, alanine and arginine
(alone or in combinations) remain to be formally evaluated.
6. Conclusion
The protective interventions briefly outlined in this
review are those with some immediate possibility for
clinical application in the near future; either drugs
already in clinical use for other purposes or nutritional
supplements as summarized in Table 1. When developing therapies for DMD, the goal is to maintain or
promote skeletal muscle mass and function but at the
same time reduce any deleterious side-effects, such as
unwanted cardiovascular complications seen with powerful muscle anabolic agents. Early treatments administered before the pathology manifests are expected to
be the most efficacious. Aggressive inflammation is a
secondary characteristic of the disease and specific antiinflammatory drugs seem to be a logical progression in
the search for an alternative to steroids. Dietary supple-
mentation in conjunction with some anti-inflammatory
drug seems particularly promising and requires further
rigorous investigation.
Acknowledgements
Related research in the author’s laboratories was
made possible by funding from: Akton Benni & Co.
(MG), AFM (MG), Italian Telethon (ADL), National
Health & Medical Research Council (GL) Muscular
Dystrophy Association, USA (GL).
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Miscellaneous
Blackwell
Publishing
Asiain skeletal muscle
MD
TNF
Grounds
and IGF-1
et al.
signalling
doi: 10.1111/j.1440-1681.2007.04868.x
Proceedings of the Australian Physiological Society Symposium:
Myopathies and Muscle Regeneration
IMPLICATIONS OF CROSS-TALK BETWEEN TUMOUR NECROSIS
FACTOR AND INSULIN-LIKE GROWTH FACTOR-1 SIGNALLING IN
SKELETAL MUSCLE
Miranda D Grounds,* Hannah G Radley,* Bijanka L Gebski,* Marie A Bogoyevitch † and Thea Shavlakadze*
*School of Anatomy and Human Biology, The University of Western Australia, Crawley, Western Australia and
†
Department of Biochemistry and Molecular Biology, Bio21 Molecular Science and Biotechnology Institute,
University of Melbourne, Parkville, Victoria, Australia
SUMMARY
1. Inflammation, particularly the pro-inflammatory cytokine
tumour necrosis factor (TNF), increases necrosis of skeletal
muscle. Depletion of inflammatory cells, such as neutrophils,
cromolyn blockade of mast cell degranulation or pharmacological
blockade of TNF reduces necrosis of dystrophic myofibres in the
mdx mouse model of the lethal childhood disease Duchenne
muscular dystrophy (DMD).
2. Insulin-like growth factor-1 (IGF-1) is a very important
cytokine for maintenance of skeletal muscle mass and the
transgenic overexpression of IGF-1 within muscle cells reduces
necrosis of dystrophic myofibres in mdx mice. Thus, IGF-1 usually
has the opposite effect to TNF.
3. Activation of TNF signalling via the c-Jun N-terminal
kinase (JNK) can inhibit IGF-1 signalling by phosphorylation
and conformational changes in insulin receptor substrate (IRS)1 downstream of the IGF-1 receptor. Such silencing of IGF-1
signalling in situations where inflammatory cytokines are elevated
has many implications for skeletal muscle in vivo.
4. The basis for these interactions between TNF and IGF-1
is discussed with specific reference to clinical consequences for
myofibre necrosis in DMD and also for the wasting (atrophy) of
skeletal muscles that occurs in very old people and in cachexia
associated with inflammatory disorders.
Key words: inflammation, insulin like growth factor 1,
necrosis, skeletal muscle, tumour necrosis factor.
Correspondence: Miranda D Grounds, School of Anatomy and Human
Biology, The University of Western Australia, Crawley, WA 6009, Australia.
Email: [email protected]
Presented at the AuPS Sympoisum Myopathies and Muscle Regeneration,
September 2006. The papers in these proceedings were peer reviewed under
the supervision of the AuPS Editor. The papers are being published with the
permission of AupS and were initially published on the AuPS website http://
www.aups.org.au
Received 28 June 2007; revision 18 October 2007; accepted 12 November
2007.
DUCHENNE MUSCULAR DYSTROPHY AND
THERAPIES
Duchenne muscular dystrophy (DMD) is an inherited X-linked
lethal childhood muscle disease caused by a defect in the gene for
dystrophin, which affects young boys, causes extreme wasting and
loss of function of skeletal muscles and leads to death usually by
20 years of age. Dystrophin is located beneath the sarcolemma and
is part of a large dystrophin–dystroglycan complex that forms
a critical link for force transmission between the contractile machinery
of the muscle fibre and the extracellular matrix. Where dystrophin
is defective or absent, the myofibre is fragile and the sarcolemma is
readily damaged in response to exercise, leading to myofibre necrosis.1
Although it is widely considered that mechanical tears in the
sarcolemma are the cause of the initial damage, other data indicate
that changes in ion channels may be responsible for the initial influx
of calcium that causes the damage;2 clearly, an accurate understanding
of the basic mechanism will affect the targeting of potential
therapeutic interventions. Although myofibre necrosis normally
results in new muscle formation, in DMD (and, to a lesser extent,
in the mdx mouse model of DMD) it appears that regeneration fails
over time and the dystrophic muscle is progressively replaced by
fatty and fibrous connective tissue.
Although the defective gene, dystrophin, was identified in 1987,
there is still no effective treatment for DMD boys. Although cell or
gene therapy to replace the defective dystrophin is the ideal scenario,
the clinical application of such therapies is yet to become a reality.3,4
Meanwhile, many preclinical studies continue on the mdx mouse
model of DMD.5
The existing treatment for DMD is administration of corticosteroids;
these are broad-based anti-inflammatory drugs that decrease
inflammatory cell populations in dystrophic muscle6 and increase
myofibre mass, although the precise mechanism of action in DMD
is not yet known and is under intense investigation.7,8 One disadvantage
of steroids is that they are associated with severe adverse side-effects,
such as weight gain and osteoporosis,9 and the response is variable
between individual boys.10
TNF and IGF-1 signalling in skeletal muscle
Fig. 1 Immunohistochemical staining for tumour necrosis factor (TNF) in
transverse sectioned dystrophic (mdx) and non-dystrophic (C57Bl/10) mouse
muscle (TNF stains brown). (a) Non-dystrophic muscle shows no specific
staining for TNF. Bar, 100 mm. (b) Dystrophic muscle shows TNF staining
on the plasma membrane of muscle fibres and also in areas of focal necrosis
(asterisk), with faint staining in some apparently intact myofibres (arrows).
Bar, 100 mm.
While waiting for gene correction therapies to hopefully become
a clinical reality, attention has turned to drug and nutritional
interventions that target inflammation, fibrosis, necrosis, muscle
mass and reactive oxygen species, designed to reduce the severity
of the dystropathology (For reviews, see Tidball and WehlingHendricks11 and Radley et al.12). The best combinations of such
drugs,13 alone or with dietary interventions,14 for long-term use in
DMD remains to be determined.12
ROLE OF TUMOUR NECROSIS FACTOR IN
MUSCULAR DYSTROPHY
There is increasing evidence that inflammation contributes to the
necrosis of dystrophic myofibres.15,16 When myofibre breakdown and
necrosis occurs, inflammation and associated cytokines are essential
for removal of necrotic tissue and for the formation of new skeletal
muscle. Inflammatory cells and a range of cytokines influence
myoblast activation, migration, proliferation, differentiation and
fusion. Strong evidence that inflammatory cells can contribute to
necrosis of healthy muscle cells comes from studies investigating
the role of neutrophils, macrophages and oxidative damage in vitro17–20
and in vivo.21–23 It has similarly been proposed that an excessive
inflammatory response can directly damage myofibres in myopathic
847
conditions, such as dystrophies or myositis,15,24 and recent data
increasingly implicate inflammation, and specifically tumour necrosis
factor (TNF), in myofibre necrosis.
Tumour necrosis factor is a major pro-inflammatory cytokine that
is expressed by a wide range of inflammatory cells and by myoblasts,
myotubes and damaged skeletal muscle.25,26 Tumour necrosis factor
is also produced by adipose tissue,27,28 that is often pronounced
within the wasted skeletal muscles in DMD. In response to even
minor myofibre injury, TNF is rapidly released from resident mast
cells and also by neutrophils, which accumulate quickly at sites of
tissue damage,16,29 and TNF is a potent chemokine that attracts
further inflammatory cells to the injured site. The chemotactic role
of TNF was demonstrated in normal mouse muscle, where
administration of TNF resulted in the accumulation of neutrophils
and macrophages in the absence of any tissue damage.30
In support of the proposal that TNF and neutrophils exacerbate
initial sarcolemmal damage and provoke necrosis of dystrophic
myofibres, in vivo blockade of TNF,31–33 cromolyn prevention
of degranulation of mast cells (which normally release high levels
of TNF)29 or depletion of host neutrophils32 protects dystrophic mdx
mouse muscle from necrosis. Two drugs that were used to block TNF
activity in the mdx mouse model of DMD, namely infliximab (an
antibody to TNF) and etanercept (soluble receptor to TNF) are in
wide clinical use already to treat inflammatory disorders such as
arthritis and Crohn’s disease. The high specificity of these anticytokine
drugs, combined with their clinical success in other diseases and
relatively few side-effects, suggests that they may be attractive
alternatives to the existing use of corticosteroids to treat DMD. In
the mdx mouse, long-term studies have further demonstrated that
the mouse-specific cVIq antibody to TNF has equal efficacy to
Remicade and Enbrel.34 It is noted that this cV1q blockade of TNF
has no effect on the low levels of chronic myofibre damage in
unexercised dystrophic muscle, in striking contrast with the marked
protective effect on exercise-induced acute myofibre necrosis,
raising the possibility of different roles for TNF (and other molecules)
in these two situations of myopathology.34 The impact of exercise,
as well as other factors such as age and gender that affect the severity
of the pathology, should be taken into account when interpreting the
expression profile of different molecules and the impact of drug
interventions and other therapies in mdx mice.
That TNF protein is elevated locally in dystrophic muscles is
supported by immunohistological studies showing increased staining
for TNF associated with necrotic areas of dystrophic muscles of the
mdx mouse (Fig. 1)35 and in biopsies from DMD patients,26 as well
as Western blotting analysis using anti-TNF antibodies in mdx muscle
extracts.36 It is noted that the issue of representative tissue from the
small biopsy sample that can be taken from DMD muscles makes
such measurements very difficult in humans. Few studies have
quantified TNF in dystrophic muscles or blood. Although one study
reported significantly higher plasma levels of TNF in dystrophic
(DMD and Becker muscular dystrophy) patients than in age-matched
control patients,37 another reported low levels of TNF levels in blood
from DMD patients.38 It has proven difficult to detect elevation of
TNF mRNA expression in skeletal muscles of adult non-exercised
mdx mice (T Shavlakadze, unpubl. data, 2007) and it seems that TNF
levels (protein and mRNA) have not been reported for dystrophic dogs.
Elevated TNF may exacerbate muscle damage through several
pathways.39 One of the contributors to TNF induced muscle necrosis
could be the inflammatory transcription factor nuclear factor
848
MD Grounds et al.
Fig. 2 Damaging effects of tumour necrosis factor (TNF) in skeletal muscle
cells are mediated, in part, by interference with insulin-like growth factor
(IGF)-1 receptor signalling. c-Jun N-terminal kinase (JNK) 1 has been
identified as one of the signalling molecules that mediates such interference;
JNK1 can associate with the IGF-1 docking protein insulin receptor
substrate-1 (IRS)-1 and inhibit its activity, which would lead to
downregulation of biological processes mediated by IGF-1 (i.e. stimulation
of protein synthesis, inhibition of protein degradation and cell survival).
Independent of the interaction with IGF-1 signalling, TNF may upregulate
protein degradation, leading to muscle atrophy, and promotes myofibre
necrosis. Nuclear factor-kB is one of the central players in both these
processes. PI3-K, phosphatidylinositol 3-kinase; FOXO, forkhead box, subgroup O transcription factor.
We have undertaken intensive studies using transgenic mice that
overexpress IGF-1 only within skeletal muscle.46,49–51 An important
finding was the demonstration that the reduced pathology in mdx
mice that overexpress the Class 1 IGF-1Ea isoform is likely due to
reduced myofibre necrosis46 and this protective effect may relate to
increased protein synthesis and decreased protein degradation. The
signalling pathways of IGF-1 are highly complex, with effects on
not only atrophy/hypertrophy via promotion of protein synthesis and
inhibition of protein degradation,48 but also on apoptosis, myoblast
proliferation and muscle differentiation.
To further complicate the situation, there are at least six isoforms
of IGF-1 and the specific biological function of different isoforms
of IGF-1 are not defined.52,53 The recent development of transgenic
mice that overexpress these different isoforms (N Winn (EMBL,
Italy), unpubl. data, 2007) will hopefully help clarify their relative
importance in skeletal muscle. It is noted that although the Class 1
IGF-1Ea isoform clearly reduced the dystropathology of mdx
mice,46,54 transgenic mdx mice that overexpress the fully processed
70 amino acid human IGF-1 within myofibres (Rska-actin/ hIGF-1
transgene) and have elevated IGF-1 in muscles and blood showed
no improvement in muscle pathology.51 Whether this lack of effect
reflects the different form of IGF-1 overexpressed within the muscle
or is due to increased IGF-1 levels seen only in the blood as well as
skeletal muscles of these mdx/hIGF-1 transgenic mice is unclear,51
but these contrasting findings emphasize the complexity of
interpreting transgenic data.53
CROSS-TALK BETWEEN TNF AND IGF-1
SIGNALLING PATHWAYS VIA JNK
(NF)-kB, because NF-kB is activated in limb muscles40 and the
diaphragm40,41 of mdx mice, as well as in muscle samples from DMD
patients.42 The blockade of NF-kB by pyrrolidine dithiocarbamate
reduces skeletal muscle degeneration in mdx mice36 and it has
recently been shown that heterozygous deletion of the p65 subunit
of NF-kB is sufficient to decrease muscle necrosis in mdx mice.40
Another possible mechanism for the damaging effects of TNF
could be by activation of c-Jun N-terminal kinase (JNK). This is of
special interest because activated JNK can inhibit the expression of
insulin-like growth factor (IGF)-1 mRNA, as well as IGF-1 signalling,
and such cross-talk between TNF and IGF-1 has many implications
for muscular dystrophy and other conditions where inflammatory
cytokines are elevated. A striking increase in phosphorylation of
JNK1 has been reported in the diaphragm muscles of 7-week-old43
and 12-month-old mdx mice,44 whereas there was little increase in
the limb muscles of 12-month-old mdx mice.44 Another study
reported increased phosphorylation of JNK2, but not JNK1, in the
limb muscle of 16-week-old non-exercised and exercised mdx
mice.45 In marked contrast with the adverse effects of TNF on
dystrophic muscle, increased levels of IGF-1 protect dystrophic
muscle from necrosis46 and the roles of IGF-1 are discussed below.
COMPLEX ROLES OF IGF-1 AND IMPORTANCE
IN SKELETAL MUSCLE
Insulin-like growth factor-1 plays a central role in myofibre
hypertrophy and atrophy47,48 and this balance is of critical importance
for muscle wasting in ageing (sarcopenia), in inflammatory disorders
(cachexia), denervation, disuse atrophy and metabolic syndrome.48
One of the main mechanism by which TNF causes myofibre atrophy
and myofibre necrosis maybe signalling mediated by NF-kB36,55
(Fig. 2). However, the deleterious effects of TNF on skeletal muscle
may also be due to interference with IGF-1 signalling.
Tumour necrosis factor may inhibit IGF-1-dependent events by
downregulation of IGF-1 synthesis56 and inhibition of signalling
pathways downstream of the IGF-1 receptor.57–59 Activation of JNK
appears to play a role in both these processes and this molecule is
the focus of the following discussion. Evidence for an inhibitory
action of TNF on IGF-1 synthesis is based on in vivo and tissue
culture experiments.56 Mice injected with a non-lethal dose of
lipopolysaccharide (known to stimulate inflammation) show
increased TNF protein in blood and increased expression of TNF
mRNA in skeletal muscle, with decreased blood levels of IGF-1 and
decreased IGF-1 mRNA expression in skeletal muscle.56 More direct
proof that TNF regulates IGF-1 expression is provided by a tissue
culture experiment where direct addition of TNF to C2C12
myoblasts and differentiated myotubes decreased IGF-1 mRNA.56
Activation of JNK has been suggested to mediate TNF-induced
inhibition of IGF-1 synthesis in C2C12 myoblasts, because in these
cultured skeletal muscle cells the JNK inhibitor SP600125 blocked
the TNF-induced decrease in IGF-1 mRNA expression.56
Interference by TNF in IGF-1 signalling may occur via inhibition
of the IGF-1 receptor docking proteins insulin receptor substrate
(IRS)-1 and IRS-2,57–59 leading to downregulation of signalling
molecules further downstream of the IGF-1 receptor that are
involved in regulation of the protein balance and cell survival
(Fig. 2). Tissue culture studies suggest that in C2C12 myoblasts
treated with TNF, JNK associates with IRS-1 and phosphorylates
849
TNF and IGF-1 signalling in skeletal muscle
Ser307.60 Phosphorylation of the Ser307 residue leads to dissociation
of IRS-1 from the IGF-1 receptor and inhibition of the tyrosine
phosphorylation of IRS-1, which is required for the downstream
signal transmission from the activated IGF-1 receptor.61 Use of the
JNK inhibitors I-JNK and SP600125 has confirmed the effects of
JNK as a negative regulator of IGF-1 signalling in C2C12
myoblasts,60 but these effects are yet to be tested in vivo. However,
in cultured 3T3-L1 adipocytes, extracellular signal-regulated kinase
(ERK) 1/2 rather than JNK seems to mediate IRS-1 phosphorylation
at the Ser307 residue in response to TNF,62 because inhibition of
ERK1/2 but not JNK1 was sufficient to abolish the Ser307 phosphorylation. In addition, the phosphorylation of IRS-1 on Ser307
takes place not only in response to TNF, but also following treatment
with insulin and IGF-1, which represents a negative feedback loop
responsible for insulin and IGF-1 resistance;62 this inhibition of
IRS-1 by insulin and IGF-1 appears to be distinct from the signalling
pathway activated by TNF.
The activation of a JNK1-mediated signal transduction cascade
has been suggested to contribute to progression of the dystrophic
mdx phenotype, independent of IRS-1 inhibition. Adenoviral
expression of the JNK1 inhibitor JNK-interacting protein (JIP) 1
in skeletal myofibres of mdx mice that also lack MyoD protected
them from degeneration and increased their cross-sectional area.44
That study suggested that the mechanism of JNK1 action in
dystrophic muscle is due, at least in part, to serine phosphorylation
and nuclear exclusion of the calcineurin-sensitive nuclear factor of
activated T cells (NFAT) transcription factor. Data demonstrating the
role of NFAT signalling in myofibre hypertophy are controversial
(for a review, see Shavlakadze and Grounds48); however, in mdx
muscle, upregulation of the calcineurin/NFAT pathway is protective
against muscle degeneration.63 Furthermore, although deflazacort
(a steroid used to treat DMD boys) did not alter JNK1 activity itself,
it increased activity of the calcineurin phosphatase and upregulated
NFAT-dependent gene expression, which, in turn, negates JNK1
inhibition.64 Taken together, these results suggest that further
evaluation of JNK inhibitors, including JNK inhibitory peptides, and
JNK ATP competitive inhibitors65,66 as new treatments for muscular
dystrophy (with potential clinical application to DMD) should be
considered.
Cross-talk between IGF-1 and TNF is further complicated by a
report that IGF-1 can inhibit TNF signalling involved in protein
catabolism, as shown in human colonic adenocarcinoma cells where
pretreatment with IGF-1 reduced TNF-mediated nuclear localization
of NF-kB.67 It was suggested that muscle-specific elevation of
IGF-1 would also intercept TNF signalling and reduce the loss of
muscle mass (cachexia) in inflammatory conditions; moreover, it has
been demonstrated recently that inhibition of NF-kB signalling
protects against denervation-induced muscle atrophy.68 Experiments
are required to test the in vivo possibility that IGF-1 may play an
inhibitory role in inflammatory mediated wasting of skeletal muscle.
BEYOND DYSTROPHY: CLINICAL
IMPLICATIONS FOR AGEING AND OTHER
MUSCLE CONDITIONS
Maintenance of skeletal muscle mass is governed by a complexity
of signalling interactions48 and age-related muscle weakness and loss
of muscle function (sarcopenia) presents many serious problems.69,70
Human studies show that, in the elderly, systemic low-grade
inflammation associated with increased blood levels of TNF and
interleukin-6 can contribute to loss of muscle mass and strength.71,72
Cytokines are responsible for muscle protein degradation in more
severe cases of inflammation, such as cancer cachexia, sepsis and
AIDS.55 Muscle wasting produced by TNF is associated with
induction of oxidative stress,55 which is considered to be a major
contributor to age-related sarcopenia. It has been suggested that the
effects of TNF on muscle atrophy may also be mediated, in part, via
interference with IGF-1 signalling47 and inhibition of the anabolic
signalling cascade downstream of the IGF-1 receptor, which would
lead to decreased protein synthesis and upregulation of atrophy
related genes. Thus, attempts to minimize muscle wasting in various
clinical conditions have focused on both anti-inflammatory drugs
to block TNF action and the development of strategies to deliver
IGF-1 to skeletal myofibre. Clarification of interactions between
these two opposing pathways presents the possibility of new
therapeutic targets and should provide valuable insight into
molecular events determining the severity of muscular dystrophy
and other muscle disorders.
ACKNOWLEDGEMENTS
Funding support is gratefully acknowledged from Akton Benni
and Co. (Bochum, Germany), Association Française contre les
Myopathies (Evry, France) and the Muscular Dystrophy Association
(Tucson, AZ, USA) for the research group of MDG and from the
National Health and Medical Research Council of Australia (MDG,
MAB, TS).
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70. Lynch GS, Shavlakadze T, Grounds MD. Strategies to reduce agerelated skeletal muscle wasting. In: Rattan S (ed.). Aging Interventions and Therapies. World Scientific Publishers, Singapore. 2005;
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71. Visser M, Pahor M, Taaffe DR et al. Relationship of interleukin-6 and
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526, e9–17.
Author's personal copy
Neurobiology of Disease 31 (2008) 1–19
Contents lists available at ScienceDirect
Neurobiology of Disease
j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / y n b d i
Review
Towards developing standard operating procedures for pre-clinical testing in the mdx
mouse model of Duchenne muscular dystrophy
Miranda D. Grounds,a,⁎ Hannah G. Radley,a Gordon S. Lynch, b Kanneboyina Nagaraju, c and Annamaria De Luca d
a
School of Anatomy and Human Biology, the University of Western Australia, Perth, Western Australia, Australia
Basic and Clinical Myology Laboratory, Department of Physiology, the University of Melbourne, Victoria, Australia
Research Centre for Genetic Medicine, Murine Drug Testing Facility, Children's National Medical Center, Washington, USA
d
Sezione di Farmacologia, Dipartimento Farmacobiologico, Facoltà di Farmacia, Università di Bari, Bari, Italy
b
c
A R T I C L E
I N F O
Article history:
Received 7 February 2008
Revised 20 March 2008
Accepted 24 March 2008
Available online 9 April 2008
Keywords:
Mdx mouse
Muscular dystrophy
Standard operating procedures
Biological variation
Muscle function
Pre-clinical trials
A B S T R A C T
This review discusses various issues to consider when developing standard operating procedures for preclinical studies in the mdx mouse model of Duchenne muscular dystrophy (DMD). The review describes and
evaluates a wide range of techniques used to measure parameters of muscle pathology in mdx mice and
identifies some basic techniques that might comprise standardised approaches for evaluation. While the
central aim is to provide a basis for the development of standardised procedures to evaluate efficacy of a drug
or a therapeutic strategy, a further aim is to gain insight into pathophysiological mechanisms in order to
identify other therapeutic targets. The desired outcome is to enable easier and more rigorous comparison of
pre-clinical data from different laboratories around the world, in order to accelerate identification of the best
pre-clinical therapies in the mdx mouse that will fast-track translation into effective clinical treatments for
DMD.
© 2008 Elsevier Inc. All rights reserved.
Contents
Introduction . . . . . . . . . . . . . . . . . . . . . . . . .
Therapeutic approaches to DMD . . . . . . . . . . . . . .
Scope and limitations of the review . . . . . . . . . . . .
Part I. The mdx mouse (and biological variation) . . . . . . . .
Dmdmdx . . . . . . . . . . . . . . . . . . . . . . . . .
Dmdmdx-2Cv to Dmdmdx-5Cv . . . . . . . . . . . . . . . .
Mdx52 mice . . . . . . . . . . . . . . . . . . . . . . .
Response of different skeletal muscles to muscular dystrophy
Limb muscles. . . . . . . . . . . . . . . . . . . . .
Diaphragm . . . . . . . . . . . . . . . . . . . . . .
Impact of growth parameters . . . . . . . . . . . . .
Biological variation between and within mice . . . . . . .
Factors influencing biological variation . . . . . . . . .
Developmental and in utero influences . . . . . . . . .
Neonatal influences . . . . . . . . . . . . . . . . . .
Gender. . . . . . . . . . . . . . . . . . . . . . . .
Genetics . . . . . . . . . . . . . . . . . . . . . . .
Stress . . . . . . . . . . . . . . . . . . . . . . . .
Good husbandry practises to minimise biological variation .
Cage design . . . . . . . . . . . . . . . . . . . . .
Night and day (light and dark cycles) . . . . . . . . .
Food and water. . . . . . . . . . . . . . . . . . . .
⁎ Corresponding author. Fax: +61 8 6488 1051.
E-mail address: [email protected] (M.D. Grounds).
Available online on ScienceDirect (www.sciencedirect.com).
0969-9961/$ – see front matter © 2008 Elsevier Inc. All rights reserved.
doi:10.1016/j.nbd.2008.03.008
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Author's personal copy
2
M.D. Grounds et al. / Neurobiology of Disease 31 (2008) 1–19
Models of exercise-induced muscle damage to increase pathology in mdx mice . . . . . . .
Voluntary wheel running . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Forced treadmill running . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Part I. Conclusions and recommendations. . . . . . . . . . . . . . . . . . . . . . . . .
Part II. Parameters to measure muscle dystropathology and function in mdx mice. . . . . . . .
Measuring leakiness of myofibres . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Evans Blue Dye (EBD) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Other markers (dyes and proteins) that enter damaged myofibres . . . . . . . . . . .
Blood measurements of CK and other proteins that leak out of damaged myofibres . . .
Whole body imaging to measure histopathology over time . . . . . . . . . . . . . . . .
Histological measurements to evaluate necrosis, regeneration and fibrosis . . . . . . . . .
Young mdx mice (b 4 weeks). . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Adult mdx mice (6 weeks +) . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Older mdx mice (6 months +) and fibrosis . . . . . . . . . . . . . . . . . . . . . .
Frozen vs. fixed/paraffin-embedded muscle tissue sections. . . . . . . . . . . . . . .
Immunohistochemical and molecular analyses . . . . . . . . . . . . . . . . . . . . . .
Immunological analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Molecular analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
In vivo measurements of whole body function and muscle strength in mice . . . . . . . .
Whole animals: behavioural activity and response to exercise . . . . . . . . . . . . .
Whole animals: grip bar and rotarod strength and coordination measurements . . . . .
Whole animals: in situ nerve stimulated contraction with dynamometer to measure function
Terminal physiological measurements of muscle function . . . . . . . . . . . . . . . . .
In situ nerve stimulated contraction of individual whole muscles . . . . . . . . . . . .
Isolated whole muscles: in vitro measurements of function . . . . . . . . . . . . . .
Isolated individual myofibres in vitro . . . . . . . . . . . . . . . . . . . . . . . . .
Calcium regulation and ion channels measurements. . . . . . . . . . . . . . . . . . . .
Determination of calcium ion homeostasis . . . . . . . . . . . . . . . . . . . . . .
Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Other ion channels and biophysical recordings . . . . . . . . . . . . . . . . . . . .
Part II. Conclusions, recommendation and grading of data . . . . . . . . . . . . . . . . .
Scale to grade efficacy of treatment . . . . . . . . . . . . . . . . . . . . . . . . .
Benchmarking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acknowledgments. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Introduction
Therapeutic approaches to DMD
Duchenne muscular dystrophy (DMD) is a lethal X-linked muscle
disease due to a defect in the sub-sarcolemmal protein dystrophin, that
leads to membrane fragility, myofibre death (necrosis) and replacement
of skeletal muscle by fibrous and fatty connective tissue (due to failed
regeneration). This results in extensive wasting, weakness and loss of
muscle function leading to death, often by the early 20s. DMD affects
males although some carrier females can manifest and be severely
affected (depending on the proportion of normal X-chromosomes that
are inactivated during development) (Matthews et al., 1995; Wenger
et al., 1992). While the genetic defect was identified in 1987, there is still
no effective treatment for DMD. The therapeutic research approach that
has received most attention to date is replacement of functional dystrophin by genetic, cell transplantation or molecular interventions, and
there are many exciting developments in this field (Odom et al., 2007). In
parallel, there is increasing interest in administration of exogenous
factors (drugs or food supplements) to reduce the extent of myofibre
necrosis, since promising protective effects have been reported for a
variety of agents (Radley et al., 2007; Tidball and Wehling-Henricks,
2004) and combinations of such interventions present a daunting array
of protocols to be tested. In addition it is feasible that pharmacotherapy
may be necessary to increase the efficiency of genetic or molecular
interventions, a factor that extends the importance of adequate preclinical tests for single or combined approaches.
During the last 20 years, various laboratories world-wide have
focused on clarifying the pathogenic mechanisms and the eventual
compensatory mechanisms, consequent to the primary defect in dys-
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trophic muscles; this approach has led to the identification of new
potential drug targets. Interestingly, different laboratories, using a
variety of independent experimental approaches, generally obtain
similar results in terms of factors aggravating the pathology and the
potential efficacy of various drugs. However, comparing the relative
efficacy of different drugs and interventions between laboratories is
still difficult with consequent delay in data sharing and fragmentation
of efforts. This observation pushes toward a concerted development
of standard operating procedures (SOPs) for experiments in mdx
mice that will simplify and hasten comparisons of data from different
laboratories around the world and assist the research of scientists and
pharmaceutical companies to optimise pre-clinical treatments for
translation into clinical therapies.
Scope and limitations of the review
There are several animal models for DMD and all of these, like the
human counterpart DMD, have defects in the sub-sarcolemmal
protein dystrophin that make the muscle membrane fragile and result
in necrosis of skeletal muscle fibres along with cardiac and other
problems (Collins and Morgan, 2003; McNally and MacLeod, 2005).
Only the skeletal muscle situation is addressed in this review. The mdx
mouse, first identified in 1984, is the most widely used model due to
ease of breeding, genetic uniformity, economy, and convenience for
laboratory experiments. Similar pathology to mdx is seen in mice
lacking alpha-sarcoglycan, another protein in the dystrophin dystroglycoprotein sarcolemmal complex (Duclos et al., 1998). Dystrophic
dog models of DMD were first identified in 1988 (Kornegay et al.,
1988) in the dystrophic golden retriever (Collins and Morgan, 2003)
which has a much more severe pathology than the mdx mouse and
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more closely resembles the human condition: whereas the smaller
dystrophic beagles exhibit a less severe pathology (Shimatsu et al.,
2003; Yugeta et al., 2006). The highly variable phenotype of dystrophic
dogs combined with expense of maintaining colonies has limited their
use for pre-clinical testing. Beyond these mammalian models, complementary use is being made of invertebrate models that are readily
manipulated genetically and are relatively easy and inexpensive to
breed and maintain, such as the dystrophic worm Caenorhabditis
elegans (Collins and Morgan, 2003) and dystrophic zebra fish (Bassett
and Currie, 2004; den Hertog, 2005); although the usefulness of these
models for drug screening for human conditions is debated.
The review is comprised of two main parts. Part I is a description of
the mdx mouse model and the high biological variation. In Part II the
main parameters used to measure specific effects on the dystrophic
muscles are discussed. Some basic protocols are proposed for both
Parts I and II.
Part I. The mdx mouse (and biological variation)
Since the review is focused on the mdx mouse model of DMD, it is
pertinent to first outline the variations of the mdx model that are
available.
Dmdmdx
The classical biochemical and genetic mouse model of DMD is the
mdx mouse discovered in 1981 (Bulfield et al., 1984; Hoffman et al.,
1987). Over the last 25 years, many elegant papers have been published
on the mdx mouse and it is not our purpose to review this literature. In
brief, there is an acute onset of pathology (increased myofibre necrosis
and elevated blood CK) around 3 weeks of age, this reduces to a chronic
low level of damage by 8 weeks which persists throughout life but is
further decreased by 1 year of age (McGeachie et al., 1993). The mdx
mutation occurred spontaneously due to a premature stop codon
resulting in a termination in exon 23 of the dystrophin gene. The mdx
mouse has no detectable dystrophin protein, although sporadic
‘revertant’ myofibres can express dystrophin: this can complicate
interpretation of some studies especially those related to gene or cell
replacement of dystrophin (Yokota et al., 2006).
Higher (~ 3.5 fold) mortality in mdx litters is reported (Torres and
Duchen, 1987) and recent studies show increased (~ 45%) mortality
before 7 days of age in mdx mice: this is not affected by litter size with
some litters being unaffected and others totally lost [Radley and
Grounds; De Luca et al., unpublished data 2007]. Mdx mice seem very
susceptible to stress and this may be a contributing factor since
avoiding all animal handling or routine cage cleaning reduces the
neonatal mortality.
Dmdmdx-2Cv to Dmdmdx-5Cv
Elevated plasma CK levels were used to screen the progeny of
chemical mutagen-treated male mice to identify four new mutations
of Dmd, called Dmdmdx-2-5Cv (Chapman et al., 1989). Preliminary data
showed that mice with mdx2Cv and mdx3Cv mutations have
muscular dystrophic phenotypes that do not grossly differ from the
characterized mdx mutation, and spontaneous revertant fibres occur
less frequently in the Dmdmdx-4Cv and Dmdmdx-5Cv mutants than in
Dmdmdx. These additional mdx mutations have not been widely used.
Mdx52 mice
Since the point mutation in exon 23 of the dystrophin gene in mdx
mice allows the expression of four other shorter isoforms of the
dystrophin gene through differential promoter usage, exon 52 knockout mice (mdx52) were generated to simulate the DMD phenotype
commonly seen in human patients (Araki et al., 1997). A major
3
advantage of this mutation (especially for studies designed to replace
the missing dystrophin gene) is the complete absence of dystrophin
since there are no revertant myofibres. The skeletal muscle pathology
of mdx52 is similar to that of the mdx mouse for limb and diaphragm
muscles, although hypertrophy was reported in mdx52 limb muscles.
Response of different skeletal muscles to muscular dystrophy
Fast-twitch muscles are generally the most susceptible to muscular
dystrophy (and also ageing) in all species (Lynch et al., 2001). The
progress of the dystropathology has been extensively described in mdx
mice in the 1980s (Coulton et al., 1988b; Torres and Duchen, 1987)
(reviewed in Shavlakadze et al., 2004). The absence of dystrophin results
in Z-line streaming of sarcomeres by 1 day post-natal, rare isolated
necrotic myofibres are seen by 5 days, by 10 days necrosis is evident in
rostral muscles such as the head (masseter) and shoulder girdle (parascapular) (Torres and Duchen, 1987) and other muscles may show some
pre-necrotic changes (Coulton et al., 1988b). There is an abrupt onset of
skeletal muscle necrosis around 21 days of age in hind limb and many
other muscles (from 20 to 80% of the muscle can be affected) that
stimulates muscle regeneration (Whitehead et al., 2006b) (Fig. 1). Most
groups report that myonecrosis in limb muscles is rare before 21 days of
age (Shavlakadze et al., 2004), although a slightly earlier onset at 16 to
17 days has been reported in quadriceps muscles (Muntoni et al., 1993):
these differences may reflect divergence between isolated mdx colonies
in different countries. Necrosis peaks (30–60%, occasionally ~90%, of the
TA muscle is affected during this acute damage phase) by 25–26 days
(with many myotubes resulting from regeneration by day 28) and then
decreases significantly to stabilize by 8 weeks of age to a relatively low
level of active damage (~4–6%): the cyclic progression of necrosis (and
regeneration) continues throughout life although reduces by 1 year
(McGeachie et al., 1993). The acute onset of myofibre necrosis provides a
good model to study therapeutic interventions designed to prevent or
reduce necrosis, since a reduction in dystropathology is easily identified
(Grounds and Torrisi, 2004; Radley and Grounds, 2006; Shavlakadze and
Grounds, 2003; Stupka et al., 2001). However, drug interventions that
may be toxic to the post-natal development of neuromuscular apparatus
should be considered for such young mice. In contrast, reduced necrosis
can be difficult to detect in adult mdx mice where there is little myofibre
breakdown (~5%) and cumulative muscle pathology: for this reason
exercise is often used to provoke myofibre damage in adult mdx mice.
While young mdx mice at the acute phase of dystropathology show
muscle weakness and the mdx muscles appear more susceptible to
fatigue in vivo than control mice, overall, adult mdx mice do not show in
vivo functional muscle impairment up to 1 year of age (Coulton et al.,
1988a; Muntoni et al., 1993; De Luca et al., 2003). The symptoms of
dystropathology are cumulative, with fibrosis becoming increasingly
pronounced in older (15 months old) mdx mice (Lefaucheur et al., 1995):
there are several different stages in the severity of the dystropathology
between growing and mature mdx mice (Keeling et al., 2007) and these
are affected by gender (Salimena et al., 2004). The limb and diaphragm
are the 2 main groups of muscle that have been studied in muscular
dystrophy.
In addition, some muscles of the head and chest region, including
extraocular muscles (Fisher et al., 2005), masseter (Muller et al., 2001)
and the laryngeal muscles (Marques et al., 2007), show a very mild
pathology and are relatively spared from myonecrosis. The reasons for
the mild dystropathology are not clear, although it is noted that these
muscles may have an improved ability to regulate calcium homeostasis (Khurana et al., 1995) and selected mechanical properties that
offer resistance to damage (Wiesen et al., 2007).
Limb muscles
The hind limb muscles are the most widely studied and include the
tibialis anterior (TA) and extensor digitorum longus (EDL), the gastrocnemius, quadriceps and the soleus muscle (Parry and Wilkinson, 1990;
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M.D. Grounds et al. / Neurobiology of Disease 31 (2008) 1–19
Fig. 1. Histological features on a transverse section of exercised adult mdx quadriceps muscle stained with Haematoxylin and Eosin (all images are from the same muscle section).
A) Active muscle necrosis characterized by many inflammatory cells which have infiltrated dystrophic myofibres (sarcoplasm is barely visible). B) Active muscle necrosis
characterized by fragmented sarcoplasm of dystrophic myofibres with irregular shape and few myonuclei; inflammatory cells are not conspicuous. C) Recent regeneration shown by
small dystrophic myofibres (sometimes seen as smaller myotubes) with central nuclei. D) Regenerated muscle indicated by large mature dystrophic myofibres with central nuclei.
Scale bar represents 100 μm.
Wang and Kernell, 2001). The TA typically first manifests muscle
necrosis from 21 days after birth and this is more pronounced than in
the quadriceps (Radley and Grounds, 2006; Shavlakadze et al., 2004).
Another study observed more necrosis in soleus than EDL at 24 days
with greater cumulative muscle damage in soleus (~ 86%) than EDL
(~ 36%) muscle at 34 days (Passaquin et al., 2002). The precise reason
for the acute onset of dystropathology at 3 weeks of age is unresolved:
it may relate to adult-type locomotor activity or to striking developmental changes in expression of various genes, including downregulation of utrophin (Khurana et al., 1991) and of key genes involved
in creatine synthesis (McClure et al., 2007) as well as in proteins
involved in excitation–contraction coupling mechanisms (Bertocchini
et al., 1997; De Luca et al., 1990; Schiaffino and Reggiani, 1996).
Exercise has a different impact on various limb muscles (depending
on which muscles are recruited for the specific exercise regimes) and
this needs to be taken into account e.g. 48 h voluntary wheel running
doubles necrosis in quadriceps muscles of adult mdx mice, but causes
less damage to the TA and diaphragm (Archer et al., 2006; Hodgetts
et al., 2006; Radley and Grounds, 2006) and the EDL is more affected
than the plantaris and soleus muscles (Hayes and Williams, 1996).
After 3 downhill running sessions (10 m/min for 10 min at a 15°
decline) the muscles most affected are the diaphragm and triceps
brachii, with little damage seen in either the TA or EDL (Brussee et al.,
1997). A different exercise regimen, such as a protocol of chronic (at
least 4 weeks) forced running on horizontal treadmill, increases
muscle necrosis in the gastrocnemius muscle, with damage also being
observed, although to a lesser extent, in TA and diaphragm (Burdi et al.,
2006; Pierno et al., 2007) [Burdi and De Luca., data under review].
Diaphragm
Over time, the diaphragm shows a more severe pathology than the
limb muscles with extensive replacement of muscle fibres with
fibrous connective tissue in mdx mice, more closely resembling the
severe pathology of DMD where loss of diaphragm function is a major
problem (Lynch et al., 1997; Stedman et al., 1991). Normal diaphragm
muscle is composed mainly of type 2X and type 2A myofibres as
shown by histochemical staining or antibodies specific for type 2A and
2B myosins (Gregorevic et al., 2002; Shavlakadze et al., 2004). Yet in
dystrophic mdx muscles, type 2B myofibres increase over time as a
result of bouts of regeneration in response to necrosis and this is
conspicuous by 12 weeks of age (Shavlakadze et al., 2004). However, at
early stages the pathology of the diaphragm is very mild and quite
different to the severe acute onset in limb muscles: isolated necrotic
myofibres are evident earlier, by 15 days after birth, and the damage is
mild at least up to 30 days (Shavlakadze et al., 2004), with increasing
severity of myofibre degeneration and increasing fibrous connective
tissue over time (Gosselin and Williams, 2006; Krupnick et al., 2003;
Niebroj-Dobosz et al., 1997; Stedman et al., 1991). Intrinsic differences
in collagen metabolism have been demonstrated between functionally different normal skeletal muscles (Gosselin et al., 2007).
Impact of growth parameters
The much less severe dystropathology in mdx mice compared with
DMD boys may be due in large part to vast difference in growth
parameters between mice and humans, specifically to the much shorter
time-scale of growth and maturation of mice (about 3 months compared
with 20 years), much smaller body size (about 30 g compared with
70 kg) and consequent greatly reduced load on the smaller muscles in
mice. In addition, there is stress on different muscle groups due to the
use of 4 legs in mice compared with the vertical bipedal posture of
humans. A comparison of developmental milestones for mice and men
(see Box 1) suggests that as a very rough indication: 2 weeks (for mouse)
may be equivalent to 3 months (for human); 3 weeks to 6 months;
4 weeks to 10 years; 8 weeks to 20 years and 12 weeks to 25 years.
Biological variation between and within mice
Factors influencing biological variation
Great variation in the timing and severity of the dystropathology
can be seen between and within different mdx mouse colonies,
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M.D. Grounds et al. / Neurobiology of Disease 31 (2008) 1–19
Box 1
Developmental milestones in mice and men
Composite data, compiled in close collaboration with Marta
Fiorotto (Baylor College of Medicine, Houston, Texas)
When trying to draw parallels between the mouse and human
for the study of muscle, the lines should be drawn on the basis of
hormonal changes and physiological milestones beginning with
muscle differentiation. It must be recognised that there is no
linear scaling; the proportion of the life-span taken to sexual
maturity is only about 5% for mice, whereas for man it is
approximately 20%. (The long childhood is a unique characteristic of man and the apes; rodents and many other mammals do
not have a “childhood” and they effectively go from baby to
teenager.) There is also a rostro-caudal pattern of development so
that the muscles in the upper part of the body are likely to be at a
slightly more advanced stage than that described below which is
derived mainly from the study of (male) mouse hind limb.
Birth: skeletal muscle is poly-innervated, the tubular systems are
rudimentary, and neonatal myosin heavy chain predominates. At
the same time activity of thyroid hormone and the HPA axis are
suppressed. Human muscle is at a similar stage of differentiation
at about 18–22 weeks of gestation.
By 8–10 days post-natally: the tubular system is more mature,
fibres have become mono-innervated and enable the coordinated
contraction of muscles to promote balance and increased
locomotor activity. With respect to locomotor function, mice
begin weight bearing from 1–2 weeks of age, this contributes to
synapse elimination (Minatel et al., 2003) and maturation
(Missias et al., 1996), whereas in humans this is completed by
birth (Hesselmans et al., 1993). In mice, activity of the thyroid
(McArdle et al., 1998) and hypothalamic pituitary adrenal (HPA)
(Schmidt et al., 2003) hormonal axes are starting to increase, and
replacement of immature MHC by the adult myosin heavy chains
is occurring (Allen and Leinwand, 2001). Mice are suckled on milk
which is high fat/low carbohydrate. This roughly corresponds
with a newborn human (Bronson, 2001; Elmlinger et al., 2001).
[In mdx mice, blood CK levels at 7 days are the same as normal
mice but by 10 days are elevated — indicating early symptoms of
the disease.]
14–16 days: Milk is gradually becoming limiting and insulin levels
decrease substantially. The GI tract is not fully capable of digesting
complex carbohydrates, and there is evidence from the inflexion in
the normal muscle growth curves, that growth capacity is limited.
At this time myostatin is also increasing rapidly. Muscle IGF-II
mRNA and muscle IGF 1 receptor levels are approaching a nadir. In
the human this represents about 3 months of age.
19–21 days: Mice are fully weaned: muscle is mature (Allen and
Leinwand, 2001). Diet is now largely chow, and there is very rapid
growth of the muscle. Growth hormone starts to increase from
21 days and peaks around 28–30 days (Alba and Salvatori, 2004).
This represents approximately 6 months in humans (Butler-Browne
et al., 1990; Mehta et al., 2005). [In mdx mice, acute muscle
necrosis starts in hind limb muscles from 21 days: necrosis is seen
in forelimb and other muscles at earlier ages.]
5
4–5 weeks: pre-adolescence in mouse; gender differences are just
beginning to emerge: equivalent to about 10 years of human age.
5–7 weeks: puberty in mice (Jean-Faucher et al., 1978), they
are fully fertile and growth rate decreases: about 14–18 years of
age in humans (Rodriguez et al., 2007).
Around 10–15 weeks: muscle has reached a maximum size in
mice and growth in general has reached a plateau (BaliceGordon and Lichtman, 1993): N20 years in the human. [In mdx
mice, muscle necrosis stabilizes to a low persistence level of
damage, b10% of muscle affected, that is increased by
exercise.] 12–18 months: in the mouse a gradual diminution in
muscle mass becomes evident from 18 months, i.e. start of
sarcopenia. In humans, this starts significantly after about
50 years of age (Shavlakadze and Grounds, 2003).
24+ months: life-span in the mouse is approximately
30 months (varies between strains), that may correspond to
about 80 years in humans. [In mdx mice, myofibre necrosis in
hind limbs is further decreased and blood CK levels are very low
by 1 year of age.]
Summary of possible parallels for younger mdx mouse (during first 6 months)
Mouse
(weeks)
Human
1
2
3
4
6
8
12
26
Newborn
3
months
6
months
10
years
16–18
years
20
years
25
years
35
years
between littermates and even between 2 legs of an individual mouse.
Parameters affected include histological (e.g. the extent of necrosis
and muscle damage varies from massive [60–80%] to moderate
necrosis [20–40%] at 23 days with some mice showing almost no
damage even at 26 days (Radley and Grounds, 2006); this occurs
between littermates and between TA muscles in both legs of an
individual mouse e.g. 0% compared to 24% for one mouse at 21 days
[Radley and Grounds, unpublished data].); cellular (e.g. number of
satellite cells (Schafer et al., 2005) and age-related increase in
revertant myofibres (Yokota et al., 2006)); biochemical (serum
creatine kinase (CK) levels in sedentary and exercised mice (Burdi
et al., 2006; De Luca et al., 2005; Pierno et al., 2007; Vilquin et al.,
1998) [see Blood measurements of CK and other proteins that leak out
of damaged myofibres]; molecular [gene expression profiles (Turk
et al., 2006)] and functional aspects (participation in voluntary wheel
exercise (Radley and Grounds, 2006) or forced treadmill running (De
Luca et al., 2003; Vilquin et al., 1998) [discussed in Voluntary wheel
running and Forced treadmill running]. Such inherent variation
necessitates constant conditions when grouping the animals, with
large sample sizes (often at least 6–8 mice) to show statistically
significant effects of treatments.
Epigenetic and genetic factors probably contribute to the wide
range of phenotypic expression of muscular dystrophy (as discussed
below). While it is clearly impractical to standardise many of these
variables across laboratories globally, a deeper understanding of the
many reasons for biological variation will help to initiate practises to
minimise this problem.
Developmental and in utero influences
It is now widely recognised that very early events before or at
fertilisation, as well as in utero can have dramatic post-natal effects
that contribute to large biological variation in anatomy, physiology,
behaviour and the onset of disease (Gartner, 1990; Vandenbergh,
2004). Many of these are due to pre-natal hormone exposure;
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however environmental epigenetic also plays an important role in
disease susceptibility (Jirtle and Skinner, 2007; Vandenbergh, 2004).
Some of the in utero factors to consider that may influence the severity
of the dystropathology in individual mdx mice (within a litter and
between litters) are the size of the litter, the position within the
uterus, the number of male siblings and the proximity of female pups
to male littermates (this influences exposure to testosterone) along
with a range of other factors (Vandenbergh, 2003). Some laboratories
standardise litter size (e.g. 4–8 pups) for all experiments to minimise
variation in litter size that can affect initial body weights and thus
biological variation. However, this may be considered wasteful and is
unlikely to become a standard, plus it does not take into account the
consequence of early neonatal mortality of mdx litters. One simple
solution that is strongly recommended to reduce effects of inter-litter
variation is to always select pups from a single litter for both test and
control mice.
Neonatal influences
The early post-natal environment is important with a wealth of
data showing that maternal care mediates further variation in
offspring phenotype and behaviour (Champagne et al., 2007) and
catch-up post-natal growth by low birth weight humans and animals
affects many metabolic and signalling events. Transmission of maternal behaviour (such as grooming and licking) and stress responses can
occur from one generation to another by epigenetic modification of the
chromatin around the glucocorticoid receptor gene (Francis et al.,
1999; Weaver et al., 2004). The net result to the progeny is tighter
regulation of stress hormone levels, an effect that is relayed to the next
generation in subsequent maternal behaviour patterns.
These issues highlight some of the developmental variables that
can influence the phenotype (severity of the dystropathology and
potentially the propensity for voluntary exercise) of genetically
equivalent inbred mice to contribute to the wide biological variation
seem for mdx mice, both within and between litters.
Gender
Major sex differences have been noted in skeletal muscles in
relation to energy metabolism, fibre type composition and contractile
speed. Generally, muscles from males tend to be faster and have
higher maximum power output than from females, whereas muscles
from females are more fatigue resistant, recover faster from repeated
contractions and show less mechanical damage after exercise (Glenmark et al., 2004). Striking differences have been noted in the
dystropathology between male and female mdx mice at different ages
(Salimena et al., 2004), with less susceptibility to muscle damage in
young (6 weeks) female mice but greater fibrosis in old females (1 and
2 years) suggesting a role for female hormones in the pattern of
myonecrosis and repair. Other studies show that levels of blood serum
creatine kinase are higher in older male mdx mice (Yoshida et al.,
2006). Sex differences in muscle membrane damage in response to
intense exercise have been shown in animals and humans, with
females being less susceptible than males: this appears to be due to a
reduced inflammatory response in females, with many aspects being
influenced by gender (Stupka et al., 2000). It is widely recognised that
both the innate and adaptive immune responses of females are
heightened and more robust than in males (Verthelyi, 2006) and
recent evidence confirms that the innate and adaptive arms of the
immune system are different in post-pubertal male and female mice
(Lamason et al., 2006). These gender differences are largely attributed
to oestrogen levels and the regulation of nitric oxide by oestrogen
increasingly appears to play a key role (Verthelyi, 2006). There are also
gender differences in response to drugs (Franconi et al., 2007) and to
diets (see Food and water) and thus gender should be carefully
considered when testing pharmaceutical or nutritional interventions.
While such gender-related issues appear to be important in the
mdx mouse, they have barely been considered. Interpretation and
data comparison is complicated when the gender of the mice used in
experiments is either not specified (Hall et al., 2007; Kaczor et al.,
2007) or mixed males and females are used (Granchelli et al., 2000).
Ideally pre-clinical tests should be performed on groups of mdx mice
of the same sex.
Males. Testosterone has many effects; male mice of some strains are
particularly aggressive and can be difficult to cage as placing males
together may lead to fighting and thus additional muscle damage. This
is generally not a problem if males are either caged together at
weaning, many mice are caged together (since 2 males alone in one
cage are more likely to fight), and the males are not near a stud male or
females (due to pheromones). The male response to female odours can
affect behaviour and is influenced by the major histocompatibility
type of the foetus of pregnant females (Beauchamp et al., 2000). In
addition, age-related changes in pheromones in urine appear to relate
to altered immune function (Osada et al., 2003). The effects of such
odours within an animal house can be minimised by using
individually ventilated cage systems (see Cage design). In addition,
inter-male aggression in grouped housing is influenced by different
kinds of environmental enrichment (Van Loo et al., 2004). Beyond the
sex determining region on the Y-chromosome (SRY) and testosterone,
there are multiple pathways that control sexual differentiation; e.g.
sexual differences in motoneurones may be affected by the testicular
hormone Mullerian Inhibitory Substance that is present in the blood
of pre-pubertal males, but not females (Wang et al., 2005). Since DMD
affects boys, it might be considered that it is more appropriate to use
male mice, although the overall significance of this issue has yet to be
proven.
Females. Hormonal changes during the estrus cycle influence immune
status, stress, activity levels and age-related changes in cognitive
function (Kopp et al., 2006) and this may be a significant additional
variable when using female mdx mice.
Genetics
Inbred mdx mice are homozygous and theoretically there is longterm genetic stability, but there will be a slow sub-line differentiation
between colonies over many years (reviewed in Harris, 1997). Ideally,
colonies should be replaced periodically from an international source.
Furthermore, if inbred mice are not kept under specific pathogen free
(SPF) conditions, their phenotype variability can be higher than
outbred mice (reviewed in Biggers, 1958; Harris, 1997). The genetic
background strain can greatly influence the severity of the dystrophic
phenotype as demonstrated for sarcoglycan deficient mice, although
the gene loci that suppress the dystrophic phenotype remain to be
identified (Heydemann et al., 2005).
While most mdx mice are maintained as homozygous/hemizygous
colonies (where all X-chromosomes carry the gene mutation and thus
all females and males are affected), maintenance as a heterozygous
colony could more closely resemble the human situation and also
provide appropriate negative littermate controls.
Stress
Numerous factors can stress animals and this has effects on the
brain, behaviour, hormones, and the immune system. Some of the well
known stressors that produce physiological effects are social stress
related to housing and dominance/subordination (reviewed in
Bartolomucci, 2007), transport, restraint and handling, in addition to
blood sampling, surgery and anaesthesia. For example, even simply
transferring mice to a different room can increase corticosterone
levels and mice take about 4 days to acclimatize (Tuli et al., 1995). The
rapid endocrine and metabolic response to the stress of handling is
manifested by rapid changes in many blood components leading to
the recommendation that any sampling be completed within 100 s of
first touching an animal's cage (Gartner et al., 1980) since some
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components are altered by 100% within 30 min. Considering that the
in vivo procedures for drug treatment and evaluation cannot avoid a
certain level of stress being experienced by the animals, it is essential
to maintain standard handling procedures for all mice (test and
controls) throughout the experimental procedure. Anecdotal evidence
suggests that mdx mice are especially susceptible to stress.
Good husbandry practises to minimise biological variation
Environmental conditions such as housing and husbandry have a
major impact on the laboratory animal throughout its life and will
thereby influence the outcome of animal experiments. Much of the
biological variation between and within mice (and in different
laboratories) may reflect variation between aspects of animal
husbandry (reviewed in Biggers, 1958; Harris, 1997; Reilly, 1998).
Issues to consider for standard laboratory practise include; caging,
housing systems, space recommendations and microenvironmentenrichment, bedding; temperature and humidity; ventilation: air
quality, relative air pressure and individual cage ventilation; illumination: photoperiod, intensity; noises; food: types of diet, quality
assurance; water; sanitation and cleaning; identification and record
keeping. There is a wealth of literature on these topics and it is well
recognised that they have a major impact on phenotypic variation
(Gartner, 1990). Inbred mice such as mdx are far more susceptible than
outbred animals to phenotypic variation induced by minimal
environmental variability (such as pathogens), simply because the
population lacks genetic diversity (reviewed in Harris, 1997). Some of
these factors are discussed below.
Cage design
Cage design can modify the activity and behaviour of mice
(Wurbel, 2001) with changes in the environments of animals,
including environmental (supplemental) enrichment, having important effects on brain structure, physiology (including recovery from
illness and injury), gene expression in various organs (Benefiel et al.,
2005) and aggression between males (Van Loo et al., 2004).
Individually ventilated cage systems generally help to maintain low
ammonia and carbon dioxide concentrations; they support a low
relative humidity, and reduce spread of allergens and infections.
Additional benefits are that cages need to be cleaned less frequently
(with reduced disturbance of the mice) and airborne pheromones
(due to gender, stress, age) that can affect mouse hormonal responses
are eliminated (Reeb-Whitaker et al., 2001).
Night and day (light and dark cycles)
Traditionally, mice are subjected to forced exercise and experiments are performed during the day (e.g. tissue samples collection
and/or physiology experiments). Yet mice are nocturnal and normally
active at night. Since the diurnal rhythms of mice and humans are out
of phase, the nocturnal mouse more accurately corresponds to the
daytime human (McLennan and Taylor-Jeffs, 2004). It is well
documented that circadian rhythms profoundly influence many
molecular, metabolic, endocrine, immunological and behavioural
parameters (McCarthy et al., 2007; McLennan and Taylor-Jeffs, 2004)
and thus it is important, within each laboratory, to maintain strictly
the same time of day for exercising, treating, sampling or conducting
physiological experiments on mice. One strategy to consider is the use
of sodium lights to shift the day/night cycle in order and allow
observations and interventions during the nocturnal phase with many
scientific and welfare advantages (McLennan and Taylor-Jeffs, 2004).
Food and water
Diet can have a dramatic and rapid impact on cellular responses
and gene expression, as illustrated by effects of soy or casein on
cardiac hypertrophy (Stauffer et al., 2006) and how a diet enriched
with omega-3 fatty acids can have potent post-natal effects on fetal
7
programming (Wyrwoll et al., 2006): both of these studies emphasise
the strong influence of gender. There is a huge literature on nutritional
effects but these two examples illustrate the need for strictly
standardised diets. Indeed nutritional interventions to ameliorate
muscular dystrophy are being trialled in mdx mice and humans
(Radley et al., 2007). It can be a challenge to rigorously control the
quality of a well characterized diet (e.g. mouse chow) from a supplier,
since even the standard ingredients may vary depending on the
market source and the season. A further variable to consider is the
impact of short-term (e.g. overnight) fasting on metabolic and cellular
parameters. Awareness of such issues is important when evaluating
variation between results from different laboratories and even within
one laboratory over time.
Tap water that is traditionally used as drinking water in animal
houses can vary very markedly in its mineral content and different
additives such as chloride and fluoride, between different cities and
may vary throughout the year (e.g. summer and winter) for the same
location. Such differences may chemically interfere with some drugs.
Whether such variations significantly influence the dystropathology is
not known. To avoid such possible complications some laboratories,
such as pre-clinical drug testing facility at CNMC in Washington
(Nagaraju), now use purified water for all experimental mice.
Models of exercise-induced muscle damage to increase pathology in mdx
mice
Endurance training produces many physiological, metabolic and
vascular adaptations in skeletal muscle. This adaptive response may
involve myokines, such as IL-6 and other cytokines, which are
produced and released by contracting skeletal muscles and affect
other organs of the body (Febbraio and Pedersen, 2005) combined
with improvements in cardiac function. Beneficial effects of regular
exercise on dystrophic mdx muscle are reported for free wheel
running (Dupont-Versteegden et al., 1994; Hayes and Williams, 1996)
and swimming that is a non-weight-bearing, low-intensity exercise
(Hayes and Williams, 1998). The implications of such adaptation for
strengthening dystrophic muscle, although still debated, are of much
interest for physical therapy of DMD patients. However, here we will
focus on exercise as an intervention to increase the severity of the
phenotype in mdx mice, to enable more effective evaluation of drug
treatment efficacy.
In adult mdx mice the muscle pathology is normally relatively mild
and does not closely resemble the severity of DMD. The low level of
damage in adult mdx mice can be elevated by exercise that increases
myofibre necrosis and decreases muscle strength (Brussee et al., 1997;
De Luca et al., 2003; Okano et al., 2005; Vilquin et al., 1998) enabling
potential therapeutic interventions to be evaluated more rigorously
throughout the in vivo treatment (Archer et al., 2006; De Luca et al.,
2005; Granchelli et al., 2000; Payne et al., 2006; Radley et al., 2008).
Muscles differ in their susceptibility to exercise-induced muscle
damage, with fast fibres (Type 2) more likely to be damaged than slow
(Type 1) fibres. Various models of exercise-induced muscle damage
have been used to exacerbate the disease in mdx mice, including
voluntary wheel running, treadmill running and swimming (also see
Whole animals: in situ nerve stimulated contraction with dynamometer to measure function of muscle groups), but only two widely
used in vivo running models are described here.
Voluntary wheel running
The simplest model is voluntary spontaneous exercise and this is
generally well tolerated (Hayes and Williams, 1996). Voluntary wheel
running allows the distance run by individual mice to be measured
accurately and so relative activity can be related to the severity of the
resultant muscle damage. Mice are voluntarily exercised using a metal
mouse wheel placed (often suspended) inside the cage. Exercise data
are collected via a small magnet attached to the mouse wheel and a
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sensor from a bicycle pedometer attached to the back of the cage. The
sensor records single wheel revolutions, allowing total distance (km)
and speed run by an individual mouse to be determined (Archer et al.,
2006; Dupont-Versteegden et al., 1994; Hayes and Williams, 1996;
Radley and Grounds, 2006). The majority of voluntarily running is
done at night (12 h dark cycle) (Hayes and Williams, 1996; Radley and
Grounds, 2006). This exercise has the advantage that mice can be left
with the exercise equipment continuously and their activity measured
over many months. There can be wide variations in the amount of
running between individual mice, one possible disadvantage is that
mice are caged individually (even though the cages may have clear
sides so they can see each other) and this lack of socialising may affect
their behaviour. Another issue that is rarely considered is that dirt (e.g.
urine and faeces) can accumulate and increase resistance of the wheel
rotation. This can also be influenced by the design of the wheel, and
thus the amount of effort required to turn the wheel will vary as will
the impact on the muscles. Such variations in wheel resistance may
contribute to different results between mice and between laboratories
and should be considered: indeed some designs deliberately increase
the wheel resistance to increase the work load (Konhilas et al., 2005).
More sophisticated computerised monitoring systems can be used to
collect precise data on the patterns of running and stopping (Hara
et al., 2002; Radley and Grounds, 2006) and have revealed that mdx
mice run more intermittently than wild type mice (Hara et al., 2002).
Muscle necrosis is roughly doubled (increases from ~6 to 12%) in
quadricep muscle after 48 h of voluntary exercise, although other
muscles such as the TA are barely affected (Radley and Grounds, 2006)
a finding reported by others even after a single night of running
(Archer et al., 2006). Histological analysis after 48 h allows induced
necrosis to be assessed without the ensuing complication of new
muscle formation (myotubes appear by 3 days after injury). Adaptation to voluntary wheel running occurs over about 2 months in normal
adult C57Bl mice and varies between muscles (Konhilas et al., 2005).
The capacity for voluntary wheel running over a long-period (Brunelli
et al., 2007; Dupont-Versteegden et al., 1994; Radley et al., 2008)
probably reflects the relative health of mdx mice. Therefore, voluntary
exercise can provide an additional measure of the protective benefits
of interventions. (The use of voluntary or treadmill running to
measure improvements in exercise capacity is also mentioned in In
vivo measurements of whole body function and muscle strength in
mice). When mdx mice voluntarily run greater distances, this puts
additional strain on their muscles and so an improved muscle
pathology under these conditions may represent an even greater
protection than is immediately evident (Radley et al., 2008). Voluntary
wheel running combined with forced treadmill running twice a week
is another regimen to consider.
differences in capability for treadmill running include slight divergence in mdx colonies, the age of the mdx mice, their husbandry and
even calibration of treadmills. As chronic treadmill exercise is used
mainly to exacerbate the pathology, the essential requirement is to
maintain constant conditions during the protocol and to confirm
muscle impairment.
When the protocol is started at 4–5 weeks of age, mdx mice (and
rarely also wild type mice) may be reluctant to participate and
sporadically stop running and rest for a few minutes (before being
encouraged to start again) during a 30 min session of forced exercise
at 12 m/min (De Luca et al., 2003, 2002). Slight increases or decreases
in speed do not greatly change this attitude (at slower speeds all mice
can lose interest in the activity and stop more frequently). The
repeated stopping of the mdx mice may be due to increased fatigue or
avoidance behaviour. A short warm-up at a slow speed (8 m/min) for
10 min is a strategy to produce more consistent running over the
30 min run at 12 m/min (Payne et al., 2006): this helps to increase the
proportion of mdx mice that complete the 30 min standard protocol,
although 25% of 12 week old mdx mice may still fail to meet this
requirement [Radley and Grounds, unpublished data]. A low-intensity
electric shock is sometimes used to stimulate mouse running:
however, this protocol is stressful and is not recommended for the
mdx mice. A disadvantage of the treadmill exercise protocol is that it is
time consuming, since dedicated and continuous supervision is
required twice a week for a time depending on the total number of
mice to be exercised. In mdx mice, the sustained forced exercise
protocol significantly damages the triceps, gastrocnemius and quadriceps muscles. A significant increase in plasma CK is also generally
observed in exercised mdx after 4–8 weeks, corroborating sarcolemmal damage in response to muscle work load (De Luca et al., 2005;
Pierno et al., 2007).
Eccentric exercise (where activated muscles are forcibly lengthened) is most damaging to muscles of mdx mice, with more severe
damage resulting from downhill running in comparison to horizontal
running. With running downhill, mdx mice rapidly tire at a speed of
10 m/min after 30 s, are unable to run continuously and often need
encouragement after 2 min, with such exercise being limited to 5 min
bouts. In some cases this exhaustion can lead to death of mdx mice
(Vilquin et al., 1998). Uphill running is less damaging and an example
protocol of chronic running for mdx mice employed a speed of 15 m/
min, for 60 min twice a week for 5 weeks, followed by speeds of 23 m/
min for a further 5 weeks (Okano et al., 2005).
Clearly a consistent exercise regime (either voluntary exercise, or
forced exercise such as that of (De Luca et al., 2003; Granchelli et al.,
2000; Okano et al., 2005; Payne et al., 2006) needs to be used across
laboratories for accurate comparisons.
Forced treadmill running
Voluntary exercise produces mild damage in mdx mice but the
severity of damage can be increased by forced exercise running on a
treadmill (Burdi et al., 2006; De Luca et al., 2005; Granchelli et al.,
2000; Nakamura et al., 2003; Vilquin et al., 1998). This is usually done
for convenience during the day. A protocol of 30 min running on a
horizontal treadmill at a speed of 12 m/min, twice a week for at least
4 weeks, starting at 4 weeks of age, i.e. soon after the first major bout
of degeneration, causes significant weakness in the limb strength as
measured by a grip strength meter (Granchelli et al., 2000). The in vivo
weakness produced by such a protocol is observed exclusively in mdx
mice with no similar effects in wild type mice: thus providing a
reliable in vivo index with which to rapidly monitor potential drug
efficacy (De Luca et al., 2003; De Luca et al., 2002; Granchelli et al.,
2000). Due to concern about mdx mice being reluctant to run at the
high speed of 12 m/min, others have used slower speeds (6 m/min or
9.5 m/min twice weekly) but also reported muscle damage and
exhaustion with this forced running (Hudecki et al., 1993; Payne et al.,
2006; Vilquin et al., 1998). Factors that can contribute to such
Part I. Conclusions and recommendations
Part I has outlined the factors that influence biological variation
and emphasised the need to be aware of these and to standardise
conditions where possible (e.g. related to breeding, husbandry and
gender). It seems that the 3 aspects that need to be addressed in order
to help establish Standard Operating Procedures are:
1. Specify basic core experiments e.g. age of sampling, gender, exercise
regime, onset of treatment (see Recommendation I in Box 2).
2. Define basic core methods of analysis (discussed in Part II).
3. Develop a scale to grade the efficacy of treatment (see Part II).
Part II. Parameters to measure muscle dystropathology and
function in mdx mice
A range of histological analyses on muscle tissue sections, blood
measurements and physiological parameters are used to assess the
impact of various interventions on the pathology and function of
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Box 2
Recommendation I: Basic standard experimental regimes for preclinical testing
Basic regimes to assist with global standardisation of studies in
mdx mice are suggested. (Other issues of standardisation
related to animal husbandry and developmental and neonatal
influences can be more challenging to address). Key issues to
consider are: age of onset of treatment and sampling (influenced
by the experimental aim); sampling at standard ages to facilitate
comparison of much data; gender (specify gender; males may be
more suitable for testing some drugs); each litter should be
divided into test and control mice to help reduce variation;
exercise — either voluntary wheel (the total distance run by each
mouse must be measured) or forced treadmill running. (Protocols for analysis are outlined in Part II.)
Regime A To test the effects of interventions on the acute early
stage of the disease treatments are started before the onset of
necrosis (from about 14–17 days): two sampling regimes are
outlined.
A1 Sample at 28 days. Treatment is started by day 17 with
tissues sampled at 28 days (4 weeks) initially. This specifically targets the initial acute phase of myofibre necrosis.
A2 Sample at 12 weeks or later. Treatment can easily be
extended to further sampling and analysis at 12 weeks (as
for B1). Half of the mdx mice ideally should be exposed to
exercise from 4 weeks of age (either voluntary wheel
running or treadmill twice a week — or a combination) to
increase the severity of the disease. Note: young mice
before 4 weeks of age do not exercise well. (Longer term
studies can build on this).
Advantages: treatment starts before the major initial bout of
damage to mdx limb muscles. Interpretation is simplified due to
absence of pre-existing background pathology. This young age
relates to childhood events in DMD.
Disadvantages: this is an active period of growth and some drugs
may have adverse effects or be difficult to administer to very
young mice. Biological variation can be high due to differences in
the time of onset and severity of pathology between young mice.
Regime B To test the effects of interventions on adult and older
mice where there is a relatively low background level of pathology.
These regimes all involve exercise
B1 Short term (2 day) studies to specifically test the impact
on myonecrosis. This is a quick ‘proof-of-concept’ test.
Treatment of mice exercised with a single bout of
treadmill exercise (day 0), or overnight wheel running
and sampled on day 2. Use of 12 week old mice
corresponds to a standard sampling time.
B2 Short term (1 month) studies in young adult mice.
Treatment of young adult mice (8 weeks) where active
necrosis is low (but much pre-existing pathology exists).
One month of treatment/exercise with sampling at
12 weeks is a simple protocol.
B3 Long term studies. Treatment/exercise starting at 4, 8 or
12 weeks and sampling at 12, 24 or 52 weeks.
9
B4 Fibrosis and later aspects of the disease. In some cases the
treatment/exercise might start at 6 months. However,
chronic exercise started at earlier stage can increase
fibrosis, and thus some of these aspects can be investigated during B3 protocols.
Advantages: easy to obtain and work with older mice.
Pathology has stabilized and is more standardised. Exercise
exacerbates severity of the disease and can shorten the time
required to show effects.
Disadvantages: the pre-existing background pathology can
complicate interpretation of results.
muscles of mdx mice. Issues associated with these different measurements are discussed. It is noted that some of the measurements
e.g. whole body imaging and functional and physiological assessments
(see Whole body imaging to measure histopathology over time, In vivo
measurements of whole body function and muscle strength in mice
and Whole animals: in situ nerve stimulated contraction with dynamometer to measure function of muscle groups) are done on intact
animals, and often repeated throughout the study, prior to sacrifice.
Measuring leakiness of myofibres
Lack of functional dystrophin renders dystrophic myofibres
susceptible to mechanical stresses, such as exercise-induced damage
that result in small disruptions of the muscle sarcolemma. These
membrane lesions may be rapidly resealed or lead to further myofibre
breakdown and necrosis. The exact mechanisms determining whether
initial lesions result in resealing of the damaged sarcolemma (Doherty
and McNally, 2003; McNeil and Kirchhausen, 2005) or alternatively
result in myofibre necrosis are of considerable interest and underpin
much current therapeutic research. There is good in vitro and in vivo
evidence to support the hypothesis that the initial sarcolemmal
damage is exacerbated by inflammatory cells and cytokines that result
in further damage leading to myofibre necrosis (Brunelli et al., 2007;
Grounds and Torrisi, 2004; Hodgetts et al., 2006; Radley et al., 2008;
Radley and Grounds, 2006; Spencer et al., 2001; Tidball and WehlingHenricks, 2005). Two main approaches are used to measure the extent
of damaged (leaky or necrotic) myofibres. One uses labels (such as
dyes or proteins) that rapidly enter through damaged sarcolemma into
myofibres and remain in the sarcoplasm, and the other measures
blood levels of various proteins that diffuse out of damaged myofibres.
Evans Blue Dye (EBD)
Evans Blue Dye (EBD) is used to identify blood vessel and cell
membrane permeability in vivo since it is non-toxic and can be
injected systemically as an intravital dye (Hamer et al., 2002). EBD
uses albumin as a transporter molecule and diffuses into cells through
membrane discontinuities to readily identify myofibres with permeable (leaky or necrotic) sarcolemma. Damaged myofibres which stain
positive for EBD can be viewed in two different ways. Macroscopic
examination of the whole mouse once the skin is removed shows dark
blue stained myofibres that demonstrate the extent and pattern of
damage for all muscles in the body (Matsuda et al., 1995; Straub et al.,
1997). Microscopic examination of frozen muscle sections under green
fluorescent light (Hamer et al., 2002) identifies EDB positive myofibres
by red auto-fluorescence and has been widely used to quantitate
myofibre damage (as a proportion of total myofibres) in many studies
using mdx mice (Archer et al., 2006; Brussee et al., 1997; Shavlakadze
et al., 2004; Straub et al., 1997). Once EBD enters the myofibre it
diffuses along the length of the myofibre and thus may be visible at
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some distance (~150 μm) from the initial site of damage (Hamer et al.,
2002; Straub et al., 1997): this is an important consideration when
viewing transverse muscle sections. EBD persists in vivo for at least
4 days after intraperitoneal injection (Hamer et al., 2002).
Administration of EBD is by intraperitoneal (IP) or intravenous (IV)
injection. IV is widely used as it ensures rapid availability to all tissues,
although IV injections through the tail vein of (black) mdx mice can be
difficult for the inexperienced: therefore the easier route of administration is IP injection. Recommended protocols for EBD administration are: an IP injection, 16–24 h prior to tissue sampling, of a 1% dye
solution injected at 1% volume relative to body mass (Hamer et al.,
2002) or IV injection (tail vein) of EBD that can be done within 3–6 h
prior to sampling (Straub et al., 1997). The choice between these
protocols is probably not critical for many experiments.
It is agreed that myofibres with histologically distinct necrosis
always stain positive for EBD (Archer et al., 2006; Brussee et al., 1997;
Matsuda et al., 1995; Straub et al., 1997) and that myofibres with an
intact sarcolemma do not stain (Matsuda et al., 1995; Straub et al.,
1997). Some studies report that hypercontracted myofibres stain EBD
positive (Brussee et al., 1997; Matsuda et al., 1995) whereas others do
not support this conclusion (Straub et al., 1997). EBD can be present
within myofibres that appear morphologically normal in H&E stained
sections (Brussee et al., 1997; Hamer et al., 2002) and EBD uptake (into
leaky cells) does not always reflect severe myofibre damage (necrosis)
that will provoke regeneration and require myogenesis (Archer et al.,
2006). Thus, the number of EBD positive myofibres can exceed and not
accurately reflect the number of necrotic myofibres (Shavlakadze
et al., 2004; Straub et al., 1997). This must be considered when quantitating spectrophotometrically the total EBD (in a section or extracted
from dystrophic muscle) as a measure of overall damage (Hamer et al.,
2002).
Other markers (dyes and proteins) that enter damaged myofibres
Other techniques to measure sarcolemmal damage (reviewed in
Hamer et al., 2002) include identification of proteins from the blood
that have entered damaged myofibres, such as albumin (less sensitive
than EBD bound to albumin but avoids the need for EBD administration) or fibronectin (Palacio et al., 2002); fluorescent labelled dextrans
and Procion orange dye (POD). POD (FW 631 g mol− 1) is a smaller
molecule than EBD and can be administered in vitro or in vivo
(although there is some uncertainty about toxicity in the latter) (Pagel
and Partridge, 1999; Palacio et al., 2002). POD is especially useful for
post-sampling labelling of damaged myofibres by immersion of
muscles in POD solution for 30–60 min (2% wt/volume in Ringer's
solution), which conveniently avoids the need for in vivo administration (Wehling et al., 2001). However, in vitro immersion is limited by
the possibility of additional myofibre injury during the procedure
(Consolino and Brooks, 2004; Palacio et al., 2002).
Blood measurements of CK and other proteins that leak out of damaged
myofibres
In blood samples (serum or plasma), a high level of activity of the
enzyme creatine kinase (CK) (measured in a spectrophotometric
assay) is another index of sarcolemmal fragility widely used as a
diagnostic marker for muscular dystrophy (Zatz et al., 1991). It is
assumed that the enzyme activity is a direct measure of the CK protein
level in blood. The muscle (MM) isoform of CK is produced by both
skeletal and heart muscle and total CK measurement in serum can
include isoenzymes of CK derived from other tissues. Normally this is
not a major issue and can be addressed by measuring the specific CK
isoforms if required. The relationship between muscle damage and
serum CK is not always straightforward and can be influenced by
many factors. Exercise generally increases serum CK, suggesting a
direct correlation between mechanical stress and sarcolemmal
damage especially in dystrophic muscles (De Luca et al., 2005) and
lower serum CK levels are an indication of reduced pathology and
drug efficacy. CK levels in mdx mice increase between 7 and 10 days
post-natally indicating the onset of muscle leakiness, are higher in
older males than females (Yoshida et al., 2006) and decrease by 1 year
of age (Coulton et al., 1988b). CK measurements at rest range from
about 1000–7000 U/L for females and up to17,000 U/L for males and in
exercised mdx mice range from 1500–30,000U/L. Problems with CK
measurements relate to high variability between individual mdx mice
and changes related to age and the stage of disease. Humans studies
show that after a single bout of strenuous exercise (that damages the
sarcolemma), blood CK levels can increase immediately, peak by
24 hours and return to control values by 48 hours (McBride et al.,
2008). However, many factors, including training and type and
duration of excercise, affect the extent to which blood CK levels
increase and persist (Brancaccio et al., 2008) and the kinetics of blood
CK levels might be affected by the dystrophic phenotype. There is also
variability between assay runs and possible interference of chemicals
used for plasma preparation with the diagnostic kits, so it is
recommended that control normal mouse blood is included for all
assays. Collection of a sufficient volume of blood (about 200–500 μl
blood is required to yield approximately 100–200 μl serum and at least
5–50 μl is required per assay) is invasive and stressful. Thus, rather
than multipoint evaluation in the same animals, most studies collect
blood from the heart under terminal anaesthesia. Due to the high
variation, CK analysis requires quite large number of mice (n = 6–8) for
unequivocal statistical analysis. Increased blood CK levels may also
reflect increased muscle mass or increased CK within myofibres; it is
noted that up-regulation (about double) of the creatine synthetic
pathway is reported in mature muscles of mdx mice (McClure et al.,
2007). Despite these issues, dramatic changes in blood CK levels can
be a very useful measure of the severity, or correction, of dystropathology and this blood marker is used widely.
Other proteins that leak from damaged myofibres into the blood
and have been used as a measure of muscle damage include the
enzymes pyruvate kinase (Coulton et al., 1988b), aldolase, enolase,
aspartate aminotransferase, and lactate dehydrogenase isoenzyme 5,
as well as the muscle proteins myoglobin, troponin and alpha-actin
(Martinez Amat et al., 2007) and some of these have been examined in
mdx mice. In addition, leakage into serum of the soluble Ca(++)binding protein parvalbumin that is very high in fast myofibres has
been proposed as a useful diagnostic tool in mdx mice (Jockusch et al.,
1990). To date, CK measurements remain the most widely used for
monitoring muscle diseases but other more specific markers may
emerge.
Whole body imaging to measure histopathology over time
Image capture technology aims to provide routine imaging of
whole animals, body parts or whole muscles without the need for
tissue biopsy or animal sacrifice, thus allowing repeat imaging from an
individual mouse over time: this would be a great advantage.
However, these techniques are still new and not yet established for
routine laboratory use and they will face the same problems of
standardisation to allow comparison of results between laboratory
groups. A number of biomedical imaging modalities have been used to
examine muscle tissue in vivo, including ultrasonography, magnetic
resonance imaging (MRI) and confocal and multi-photon microscopy,
with the potential of Optical Coherence Tomography (OCT) just
starting to be investigated for mdx mice (Pasquesi et al., 2006)
(reviewed in Klyen et al., 2008).
MRI can distinguish between healthy and damaged dystrophic
muscles of mdx mice (McIntosh et al., 1998). The resolution is
enhanced by combination with albumin-targeted contrast agents
(taken into damaged cells) (Amthor et al., 2004), although another
study in mice concluded that endogenous MR contrast was sufficient
and did not require combination with a gadolinium based MRI
contrast agent (Walter et al., 2005). MRI (without and with contrast
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agent) has recently been used in the dystrophic dog model where very
high biological variation and the expense of dogs (Thibaud et al., 2007)
makes 3-dimensional in vivo tissue analysis of an individual animal
over time particularly attractive, especially for evaluating the effects of
pre-clinical trials. The acquisition times for MRI are long and so
motion due to cardiovascular-induced or breathing-induced artefacts
can be an issue. The resolution of MRI is not very high (the resolution
of 125 μm is insufficient to image the individual myofibres with an
average diameter of 30–50 μm) but the development of MRI scanners
specifically for mice, combined with enhanced image detail, may
result in this technique becoming more widely used experimentally.
MRI can also be used to monitor, non-invasively, transplanted stem
cells pre-labelled by incubation with ferumoxide-polycation complexes that provide images with high spatial resolution (Cahill et al.,
2004).
Confocal and multi-photon microscopy, with their superior
resolution, can readily image individual myofibres in vivo under
physiological and pathological conditions but depend on detection of
fluorescent signals. For example, two-photon microscopy was used to
characterize the topology and metabolic function of mitochondria
within skeletal muscle of a living mouse (Rothstein et al., 2005).
Whole animal imaging via fluorescent or luminescent labelling is a
rapidly evolving and promising technique (reviewed in Ntziachristos,
2006). The power of this approach was elegantly demonstrated using
transgenic α-sarcoglycan null mice that emit a fluorescent calpainactivated signal from damaged muscle (Bartoli et al., 2006), but this is
not readily applied to conventional mdx mice (that lack the vital
transgenic label).
Whether non-invasive imaging will become a routine procedure to
repeatedly monitor the status of muscles in an individual animal over
time in response to drug and other therapeutic treatments, remains to
be demonstrated.
Histological measurements to evaluate necrosis, regeneration and
fibrosis
Dystrophic skeletal muscle pathology in mdx mice is typically
assessed on haematoxylin and eosin (H&E) stained transverse
sections, with the tibialis anterior muscle of the lower hind limb
being widely used, as it is readily accessible. When undertaking
analysis it is important to consider the age of the mdx mice as
different histological features change with age (discussed in Response
of different skeletal muscles to muscular dystrophy).
Young mdx mice (b4 weeks)
The acute onset of myofibre necrosis occurs from around 21 days of
age. The acute onset of dystropathology with high levels of necrosis
provides a very sensitive assay to specifically evaluate therapeutic
interventions designed to prevent or reduce myofibre necrosis. Normal (pre-necrotic) myofibres have peripheral nuclei, intact sarcolemma and non-fragmented sarcoplasm. Necrotic muscle is identified
by the presence of infiltrating inflammatory cells (basophilic staining),
hypercontracted myofibres and degenerating myofibres with fragmented sarcoplasm. Regenerating (recently necrotic) muscle is
identified by activated myoblasts and, 2–3 days later, small basophilic
myotubes. These myotubes subsequently mature into plump myofibres with central nuclei (regenerated myofibres). Cumulative skeletal
muscle damage in young mdx mice consists of active myofibre
necrosis plus the areas of subsequent regeneration (new myofibres)
(Fig. 1). It is calculated that myonuclei of newly regenerated myofibres
of mdx mice remain in a central location for about 50–100 days and
thereafter 3–4% of myonuclei move to a peripheral sub-sarcolemmal
position; i.e. numbers of central myonuclei may decline after 100 days
of age (McGeachie et al., 1993). Myofibre size can be measured as the
cross sectional area but, while accurate for true transverse sections,
values are distorted by myofibres cut obliquely (this is a major
11
problem for clinical biopsies with variable myofibre orientation): this
problem is avoided by instead measuring the minimal Feret's
diameter of myofibres (Briguet et al., 2004).
Adult mdx mice (6 weeks +)
After the acute onset of myofibre necrosis in young mice, skeletal
muscle from adult mdx mice consists of a low level of necrotic and
regenerating (recently necrotic) tissue, regenerated myofibres (with
central nuclei) and some unaffected (intact) myofibres. As described
previously, necrotic and regenerating and regenerated muscles have
distinct histological features (Fig. 1). Unlike regenerated human
myofibres, the nuclei of regenerated mouse myofibres stay central
for many months. Therefore mdx myofibres with central nuclei are a
reliable indicator of previously necrotic/regenerated tissue (although
in other situations they might instead represent denervated myofibres). However, central nuclei do not indicate the ‘number’ of times
that an individual myofibre has undergone necrosis and subsequent
regeneration. In studies of older mdx mice, the area of muscle that has
not succumbed to necrosis can be a useful measure, since this indicates
resistance of the myofibres to damage: such unaffected intact myofibres
look normal with peripheral nuclei (although it is noted that at a lower
level the same myofibre might also contain central nuclei). Standardisation of key sampling times (e.g. 4 and 12 weeks and 6 months of age)
greatly facilitates comparison of data between laboratories.
Older mdx mice (6 months +) and fibrosis
Fibrosis and fatty connective tissue and myofibre atrophy can be
pronounced in older mdx mice. Fibrosis is readily observed in H&E
stained sections but can be emphasised by routine histochemical
stains such as Van Gieson's or Masson's Trichrome. The onset of
progressive replacement of muscle by fibrous connective tissue (mild
fibrosis) is reported in limb muscles from 10–13 weeks, with extensive
fibrous connective tissue (fibrosis) and some calcification from 16 to
20 months of age (Keeling et al., 2007; Lefaucheur et al., 1995;
Salimena et al., 2004). Intrinsic differences in collagen content
(measured by amount of hydroxyproline) and metabolism have
been demonstrated between functionally different normal and mdx
skeletal muscles (Gosselin et al., 2007). It is well documented that the
progression of dystropathology is different in the mdx diaphragm
with significant fibrosis by 9 months (discussed in Diaphragm). With
the diaphragm, extra care must be taken to cut sections at equivalent
locations for histological comparisons, due to the complex structure
and varying width of the diaphragm muscle (illustrated in Shavlakadze et al., 2004).
Frozen vs. fixed/paraffin-embedded muscle tissue sections
Frozen sections are routinely used as they avoid problems of
shrinkage due to fixation, can provide excellent histology and have the
major advantage that the same tissue can be readily used for
immunohistochemistry (since many antibodies do not work well on
fixed muscle sections). Muscles are routinely frozen in isopentane
cooled in liquid nitrogen, since the isopentane reduces surface tension
and avoids trapping air around the muscle that can slow the freezing
process. Disadvantages of frozen tissues are that skill is required to
prepare (to avoid ice artefact) and to cut sections, the tissues must be
stored at −80 °C and they can deteriorate over time. In contrast,
muscles that are fixed (in paraformaldehyde) and processed into
paraffin blocks can simply be stored on the shelf indefinitely. Muscle
sections from paraffin blocks are ideal for H&E analysis and for other
routine histochemical stains, but can be limiting with respect to
enzymatic or antibody staining.
Morphological features are usually identified manually by the
researcher and quantified using various image analysis software. The
analysis of dystropathology on histological muscle sections is highly
interpretive and thus can vary slightly between individuals and
laboratories: a standard set of reference images to emphasise the
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precise features that are measured (Fig. 1) would help to reduce this
variation. Scientific analysis that involves any degree of interpretation
should be carried out as ‘blind’ analysis.
Immunohistochemical and molecular analyses
Immunological analysis
Immunological determination can help to gain insight into tissue
events accounting for the histological changes and a wealth of
different antibodies can be used depending on the specific question
being addressed (e.g. as a consequence of drug or other treatment);
only two specific antibodies are discussed below. Determination of
dystrophin levels may help to evaluate the percentage of spontaneously and/or therapy induced revertant myofibres (Yokota et al.,
2006) and similar approaches apply for components of the dystrophin–glycoprotein complex. Other than for gene therapies, these
approaches are useful when considering drugs able to force premature
stop codon mutations, or to exert exon skipping, or to inhibit
proteasome activity (Alter et al., 2006; Bonuccelli et al., 2007; Welch
et al., 2007): ideally this results in uniform distribution of dystrophin
in most myofibres. Similarly, immunological determination of utrophin expression is an important assay to evaluate compensatory
mechanism in dystrophic muscle induced by experimental protocols
and/or drugs (Moghadaszadeh et al., 2003; Nowak and Davies, 2004).
One cautionary note is that digital imaging of fluorescently labelled
proteins or cells, in the hands of the inexperienced can sometimes
result in false positives due to confusion with background fluorescence (that can be high in muscle tissue) and this can be exacerbated
by image manipulation.
Molecular analysis
Treatment of animals with compounds can change gene expression profiles. Gene expression can be measured at the mRNA level,
using real-time quantitative RT-PCR or microarray, or at the protein
level by measuring changes in levels of specific proteins using
Western blot, Elisa, proteomics (Ge et al., 2003) or phospho-protein
profiling. Global gene expression profiling using microarrays is
increasingly popular to monitor many mRNAs of target tissues before
and after drug therapy. Microarray analysis of mdx mice at different
stages of the disease and muscle biopsies from DMD patients, provide
considerable new insights into muscular dystrophies (Chen et al.,
2005; Haslett and Kunkel, 2002; Porter et al., 2003), have identified
pathways that are amenable for therapeutic intervention early in the
disease process and large muscle biopsy microarray data sets are now
in the public domain (Bakay et al., 2002).
The powerful high-throughput tools of systems biology analysis
(for RNA and protein) combined with biochemical techniques allow
verification of the possible involvement of specific pathways in
normal and diseased muscle (Hittel et al., 2007). However, gene
expression patterns are not always easy to interpret, a change in gene
expression does not always result in a change in protein expression
and function, sub-threshold changes in both the gene and the protein
may have a great impact for tissue function, and the relative (rather
than absolute) amount of a protein may be the critical factor. Overall,
the gene, protein and emerging non-protein-coding RNA (Pheasant
and Mattick, 2007) array methods, which are difficult and expensive,
require additional functional or biochemical analysis to detect real
changes in gene product function.
In vivo measurements of whole body function and muscle strength in
mice
Body weight is usually monitored (weekly or monthly) throughout
chronic experiments as an index of general health, in addition to
numerous tests on whole animals to evaluate overall functional
capacity, muscle strength, muscle endurance and ability to fatigue and
adapt. Apart from the measurements of activity (outlined in Whole
animals: behavioural activity and response to exercise), there are
several simple non-invasive methods to evaluate muscle function in
intact whole animals (Whole animals: grip bar and rotarod strength
and coordination measurements). The techniques based on behavioural testing (open field, exercise, grip meters etc.) may be biased by
effect of drugs on tissues other than skeletal muscle, modulating
either animal motivation or animal metabolism that can modify
strength and capacity to participate in the test, with the possibility of
false positive or false negative results. Therefore detailed analysis of
functional parameters is required for validation of a benefit. Muscles
can be further tested in whole animals by invasive in situ procedures
with some possibility of repeated measurements on an individual
mouse (Whole animals: in situ nerve stimulated contraction with
dynamometer to measure function of muscle groups) and, finally,
many detailed measurements are made in vitro on muscles removed
from animals in terminal experiments (see Terminal physiological
measurements of muscle function).
Whole animals: behavioural activity and response to exercise
Since activity (i.e. exercise) affects the amount of damage of
dystrophic muscle, it is very important to determine whether a drug
or treatment has any effect on mouse activity. The Digiscan open field
apparatus measures exploratory locomotor activity of the animal, via a
grid of invisible infrared light beams, with the position of the animal
being determined when the beam is interrupted (Crawley, 1999;
Hamann et al., 2003; Nagaraju et al., 2000). Such locomotion and
behaviour is also clearly influenced by some drugs acting on cardiac
and neurological systems. Behavioural tests are prone to variability
and significant variation in absolute values can easily occur between
different laboratories and between different experimenters within a
same laboratory: therefore standard protocols must be used for these
tests. Mdx mice at certain ages (e.g. between 10 and 28 weeks) show
reduced locomotor activities in comparison to age and sex matched
control normal mice [Nagaraju; unpublished data].
Other common ways to measure activity include monitoring
voluntary wheel running or treadmill running (as outlined in
Voluntary wheel running and Forced treadmill running). The ability
of mdx mice to run in either of these situations (measured as distance run/day or week, or the time taken to cover a particular
distance) is an indication of their general well-being and muscle
function (Brunelli et al., 2007; Dupont-Versteegden et al., 1994;
Radley et al., 2008).
Whole animals: grip bar and rotarod strength and
coordination measurements
Functional strength in mice has been widely measured by
exploiting the animals' tendency to grasp a horizontal metal bar
while suspended by its tail. The bar is attached to a force transducer
and the force produced during the pull on the bar can be measured
regularly (e.g. weekly). This is a relatively simple way of measuring
body strength and repeated measurement can be made on the same
individual throughout the life of the mdx mouse. This grip bar
strength dynamometer is the most commonly used (for simplicity and
economy) in vivo test for monitoring impaired limb strength caused
by chronic exercise in mdx mice and whether a specific intervention
can reduce muscle weakness (Anderson et al., 2000; Connolly et al.,
2001; De Luca et al., 2005; De Luca et al., 2003; Granchelli et al., 2000;
Payne et al., 2006; Smith et al., 1995). Some studies suggest that
strength of mdx mice decreases after 3 months of age, but there is
some controversy (Keeling et al., 2007).
Grip strength determination has to be performed under strict
experimental condition as it may be affected by many variables (e.g.
volition, cognition and fatigue) independent of muscle dysfunction.
Therefore, as for the other behavioural approaches, the benefit of an
intervention may not be through direct effects on muscle per se. It is
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M.D. Grounds et al. / Neurobiology of Disease 31 (2008) 1–19
important that mouse strength is determined always by the same
operator, using a constant protocol, i.e. a fixed number of determinations spaced by a fixed time, and possibly in a blind-fashion.
The Rotarod measures overall motor coordination; this is a
motorised rotating treadmill which requires mice to maintain their
grip and balance on a rotating drum, or simply lose their balance and
fall off (Bogdanovich et al., 2002; Ozawa et al., 2006; Payne et al.,
2006). Other tests for motor coordination involve coaxing an animal to
walk along a narrow beam between two cages, to examine walking
performance over successive weeks of treatment, or the wire hangtest where a mouse is placed on a wire cage lid, then held upsidedown and latency to fall is recorded (Hamann et al., 2003).
Electromyographic studies are invasive but allow direct in vivo
monitoring of electrical properties of muscles and disease progression in mdx mice (Han et al., 2006a). Other invasive procedures
that directly measure muscle strength (without behavioural complications) are discussed below (Whole animals: in situ nerve stimulated contraction with dynamometer to measure function of
muscle groups and Terminal physiological measurements of muscle
function).
Whole animals: in situ nerve stimulated contraction with dynamometer
to measure function of muscle groups
Measuring muscle function in situ in anaesthetized laboratory mice
usually involves measuring the strength of an entire muscle group, such
as the ankle plantarflexors or dorsiflexors using a dynamometer
(Ashton-Miller et al., 1992; Brooks et al., 2001; Hamer et al., 2002;
Miller et al., 1998). This is an invasive technique since the (peroneal or
tibial) nerve innervating the muscle group of the leg is stimulated via
surface, hook, or needle electrodes, with the foot of one leg secured to a
force plate. When the muscle group is electrically stimulated, the
apparatus measures the moment developed about the ankle during
isometric, isovelocity shortening, or isovelocity lengthening contractions (Ashton-Miller et al., 1992). One advantage of this precise in vivo
measurement is that changes in the functional properties, e.g. adaptation, of a muscle group can be repeatedly evaluated in an individual
mouse over time [Ridgley, Grounds et al., paper under review]. This
technique involves minimal surgery and there are no complicating
factors such as muscle injury/repair that could affect force production.
The major disadvantage is that such measurements require the use of
elaborate custom-built hardware that is not widely available.
Terminal physiological measurements of muscle function
In many physiological studies, muscles are removed from the
animal and various parameters measured in vitro: therefore these
experiments are terminal, as are many in situ studies (below).
Histological analysis can subsequently be carried out on whole muscles
and these data reconciled with the physiological measurements.
In situ nerve stimulated contraction of individual whole muscles
For in situ analysis the distal tendon of (usually) a hind limb muscle
such as EDL, soleus, medial gastrocnemius, or tibialis anterior is
isolated and the tendon is sutured directly to the lever arm of a force/
position controller while still attached to the muscle or muscle group
(with the possibility of repeated experiments over time) (Brooks,
1998); alternatively the tendon is severed and then secured to the
lever arm with suture (this is usually a terminal experiment). After the
tendon is attached to the force-recording apparatus, the knee and
body of the animal is secured, and the muscle in question stimulated
by its nerve, e.g. the sciatic nerve (Brooks, 1998; Consolino and Brooks,
2004; Dellorusso et al., 2001; Schertzer et al., 2006; Stupka et al.,
2006). The advantage of this technique (as for the in vivo evaluation in
Whole animals: in situ nerve stimulated contraction with dynamometer to measure function of muscle groups) is that the muscle
13
sarcolemma, basement membrane and extracellular matrix connections remain intact, and the nerve and blood supply are undamaged
which permit evaluation of larger muscles (e.g. tibialis anterior,
gastrocnemius) that cannot be evaluated in vitro due to difficulties
with perfusion of such large muscles. A potential disadvantage of the
in situ approach is that since the muscle is stimulated via the nerve,
the accuracy of the measurements relies upon there being no
interference with normal innervation. Some neuromuscular conditions can obviously affect peripheral nerves which may thus interfere
with normal neurotransmission and invalidate the in situ approach.
Isolated whole muscles: in vitro measurements of function
Evaluation of muscle function in vitro requires careful surgical
tendon-to-tendon excision of muscles from anaesthetized animals;
these techniques take time to perfect. The slightest damage to muscle
fibre integrity during surgery compromises muscle force-producing
capacity. The isolated muscle (e.g. usually the whole EDL), is tied to a
force recording apparatus which includes a fixed pin at one end and a
lever arm of a dual-mode (force-length) servomotor or simply an
isometric force transducer. The isolated muscle is stimulated by platinum plate electrodes that flank, but do not touch the preparation. The
accurate measurement of maximum muscle force-producing capacity
(and power output) in vitro is dependent upon many factors including
surgical dissection, the use of accurate force recording equipment (to
ensure that the muscle is stimulated adequately to recruit all motor units
within the muscle) and adequate muscle perfusion to prevent any part of
the muscle becoming anoxic (Lynch et al., 2001). Muscles can be
stimulated to contract with muscle length either maintained (isometric
contractions), or shortened (“concentric” contractions), or lengthened
(eccentric or lengthening contractions). Such in vitro analysis also allows
physiological parameters of diaphragm muscle to be measured (Lynch
et al., 1997; Petrof et al., 1993; Stedman et al., 1991). In most cases,
evaluation of muscle function capacity in vivo (Whole animals: in situ
nerve stimulated contraction with dynamometer to measure function of
muscle groups) or ex vivo (Terminal physiological measurements of
muscle function) should produce very similar maximum forces. If there
is a significant discrepancy between these values, there are deficiencies
in the methodology and/or equipment. If any of these parameters are
overlooked, the measurement of the muscle's true functional capacity
will be inaccurate and therefore evaluation of the efficacy of the
pharmaceutical or nutritional intervention will be compromised. The
relative advantages and disadvantages of the in vitro preparation for
assessing muscle function, depend on whether an intact nerve and blood
supply is desirable for a specific evaluation. The in vitro preparation can
be advantageous for assessing whether a treatment affects the
contractile apparatus directly, without contributions from the nerve
and blood supply. Ideally, measuring muscle function in situ and in vitro
(using a contralateral muscle) would provide the most comprehensive
assessment of treatment efficacy. In practise, however, such an approach
is beyond the scope of most studies both in terms of the time constraints
for investigation and the technical expertise required.
Isolated individual myofibres in vitro
This technique uses isolated individual myofibres extracted from
whole muscles, rather than assessment of isolated muscle fibre
bundles or intact whole muscles. Experiments can be performed on
membrane intact myofibres (Yeung et al., 2003) or on “skinned”
myofibre preparations where the sarcolemma has been either
mechanically peeled (Plant and Lynch, 2003) or chemically permeabilized (Lynch et al., 2000). This preparation allows for direct testing of
the sensitivity of contractile filaments to activating Ca2+ (or other
divalent cations, such as Sr2+) and an assessment of the effects of
treatment on the contractile apparatus. Isolated myofibres can be
attached directly between sensitive force transducers (at one end) and
a length controller (at the other end) so that fibres can be activated
(electrically or chemically) and fibre length either maintained,
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shortened or lengthened, as per assessments on intact muscle
preparations. These experiments can be informative for assessing
the cellular mechanisms of action of novel treatments since they
permit examination of the contractile apparatus and of excitation–
contraction coupling. As for all studies using single myofibres, it is
important to recognise that only a limited number of myofibres can be
sampled and these may or may not be representative of the entire
population of fibres within a muscle. In dystrophic muscles, the
incidence of myofibres with atypical morphologies such as complex
branching also needs to be considered for these physiological
assessments (Williams et al., 1993). This helps to ensure that the
myofibre sampling methods do not skew findings due to the inclusion
of only less damaged myofibres. The single fibre assessment of muscle
function provides a more physiological approach than examination of
myotubes formed in culture (Han et al., 2006b).
Calcium regulation and ion channels measurements
Determination of calcium ion homeostasis
Altered Ca2+ homeostasis, linked to altered activity of various ion
channels, has been proposed as a key consequence of dystrophindeficiency contributing to the dystrophic pathology and to calciumdependent proteolytic events (Alderton and Steinhardt, 2000; De Luca
et al., 2003; Whitehead et al., 2006b). However, it is still debated
whether dysregulation of Ca2+ is a primary consequence, or just
another downstream player in the complex pathophysiological
cascade. Changes in excitation–contraction coupling detected by
electrophysiology are a functional integrated monitoring of altered
calcium homeostasis (De Luca et al., 2001; Fraysse et al., 2004).
A variety of methods including the use of radiolabelled calcium and
Ca2+-specific fluorescent dyes (like Fura-2 and FFP-18) in conjunction
with microspectrofluorimetry or fluorescence digital imaging microscopy, have been used to measure alterations in resting intracellular
free [Ca2+] ([Ca2+]i), sarcolemmal permeability and the events
controlling Ca2+ homeostasis. These techniques are usually applied
ex vivo on collagenase dissociated individual myofibres, myotubes
(often of the C2C12 cell line) or intact tendon-to-tendon bundles of
muscle fibres, but similar methods can be used to visualize Ca2+
events in peripherally located myofibres within intact muscles in
vivo. The development of newer generation genetically encoded
calcium biosensors offer the promise of improved Ca2+ sensitivity
both in vitro and in vivo and will allow for powerful two-photon Ca2+
imaging of normal and dystrophic myofibres. A detailed discussion of
these techniques is beyond the scope of this review although the
determination of calcium homeostasis is an important end-point in
pre-clinical testing. Despite different laboratories finding either
dysregulation or no change in cytosolic Ca2+ levels in dystrophic
muscle fibres (De Backer et al., 2002), attributed primarily to
differences in fibre preparations and the dyes employed, there is a
general consensus that dystrophic myofibres exhibit an increased
sarcolemmal permeability to Ca2+, especially in response to osmotic
and/or mechanical stress (Allen et al., 2005; Fraysse et al., 2004; Han
et al., 2006b).
Channels
Altered Ca2+permeability and homeostasis in dystrophic muscle
may result from a greater activity/expression of specific subset of
voltage-independent ion channels, rather than from physical breaks in
the sarcolemma (Fraysse et al., 2004; Whitehead et al., 2006b).
Attention has been focused on various ion channels involved in
sarcolemmal permeability including stretch-activated and transientreceptor-potential (TRP) channels, as this may lead to a better
understanding of the pathogenetic events as well as identification of
possible target for pharmacological intervention (Iwata et al., 2003;
Rolland et al., 2006; Whitehead et al., 2006a). A variety of techniques
are used for characterization of the candidate channels, such as
biophysical (see below), immunofluorescent and biochemical
approaches (RT-PCR, Western blotting etc.) in parallel with toxins,
antibodies and drugs able to target specific channels (Rolland et al.,
2006; Suchyna and Sachs, 2007; Yeung et al., 2005).
Fig. 2. Example of a flowchart for a typical 3–6 months pre-clinical drug trial in mdx mice based on protocols used by the authors. The items shown in bold (indicated by A) are
essential for all pre-clinical trials. Many of the remaining procedures (indicated by the letter B) are highly recommended, with additional analyses also included.
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M.D. Grounds et al. / Neurobiology of Disease 31 (2008) 1–19
Other ion channels and biophysical recordings
Sarcolemmal ion channels, gated by voltage/mechanical/chemical
stimuli, are important in determining excitation patterns and a proper
contractile response. In dystrophic myofibres alterations in ion
channels can result either directly from dystrophin-related sarcolemmal defects, or from downstream mechanisms modulating their function and/or expression. Electrophysiological techniques can measure
ion channel function (voltage-clamp recordings) and membrane
excitability characteristics (current clamp recordings), by electrodemediated recordings in isolated muscles or myofibres. A detailed
description of these specialised techniques is beyond the scope of this
review. However, patch clamp recordings in freshly isolated myofibres
and/or cultured myotubes are highly versatile allowing detection of a
single channel current for any sort of ion channel. Although a large
number of native freshly dissociated myofibres have to be sampled to
be representative of all muscles, alteration in Ca2+ permeable channels
have been clearly described in mdx myofibres by this approach
(Rolland et al., 2006; Vandebrouck et al., 2002; Yeung et al., 2005).
Apart from over-activity of voltage-independent Ca2+ channels, the
muscle chloride (Cl−) channels (CIC-1) are most affected by muscular
dystrophy. These channels are responsible for the large conductance
of Cl− (gCl), that provides electrical stability of the sarcolemma, and are
monitored by the intracellular microelectrode current clamp method
in intact isolated muscle. A decrease in gCl is a functional cellular sign
of spontaneous (as in diaphragm) or exercised-induced (as in EDL)
damage in mdx muscles, and leads to an altered excitability pattern
(De Luca et al., 1997, 2003; Pierno et al., 2007). This biophysical
impairment can account for an increased excitability-induced stress of
dystrophic muscle as well as for the EMG abnormalities detected in
both mdx mice and DMD patients (Han et al., 2006a).
Part II. Conclusions, recommendation and grading of data
The many techniques that can be used to measure specific aspects
of the pathophysiology of muscular dystrophy (e.g. cellular changes
and function), to assess the efficacy of an intervention, have been
outlined. A suite of core methods for analysis of pre-clinical
experiments in the mdx mouse model are outlined in Recommendation II (see Box 3). Additional techniques will be used depending on
this initial analysis and the specific questions being addressed by
various experiments. A flow chart outlining a broader range of
analyses is shown in Fig. 2. One result of a recent Workshop on Preclinical testing for Duchenne dystrophy: End-points in the mdx mouse
held in Washington, USA in October 2007, is that details of many
methods for analysis will be collated and available in 2008 on the
websites for TREAT-NMD (http://www.treat-nmd.eu/) and WellstoneDC (http://www.wellstone-dc.org/) to help standardise the use of
these procedures.
Scale to grade efficacy of treatment
To enable efficient comparison of data a universal grade to rank the
relative efficacy of each treatment is needed. Since different
interventions may affect one aspect (e.g. histology) more than another
(e.g. physiology) it seems appropriate to grade the outcome for each
parameter, rather than simply use one overall final score. A scale with
5 grades is proposed for each parameter ranging from 1 (best) to 5
(worst), with ‘1’ corresponding to the measurement in normal control
C57Bl/10Scsn mice and ‘5’ to that in untreated mdx mice. (This grading
might also be applied retrospectively to many published papers to try
and facilitate some meaningful comparison of existing data).
Benchmarking
Benchmarking of reference data for mdx and normal C57Bl/10Scsn
mice is required to help standardise measurements. While it is
appreciated that measurements may vary slightly between different
laboratories (and this may be more of an issue for some techniques),
15
Box 3
Recommendation II. Basic methods to analyse the pre-clinical
experiments.
A core of basic measurements that are possible for most
laboratories is proposed. Clearly many other measurements are
possible routinely in various laboratories but the core measurements should be made by all laboratories to help standardise
basic comparison of data.
If an effect is found then many additional analyses can be
undertaken often using the existing sampled tissues. Therefore,
need to ensure that tissues are collected in the correct way for
such further analyses e.g. fresh myofibres for electrophysiology
or calcium determination and frozen sections for immunohistochemistry, frozen tissues for protein and RNA. Physiological
measurements are very informative but may not be possible in all
laboratories (therefore this is shown in italics). However some
measure of muscle function and strength such as the capacity
for exercise or grip strength should be included.
The minimal measurements are indicated below in the order
in which they would be done for sampling. Items 3 and 4 are
essential. However, it is highly recommended to try and include
items 1 and 2. This simple suite of procedures is based on
standard analyses along with the opportunity for more in-depth
investigations. (An example of a flow chart for a pre-clinical drug
trial is shown in Fig. 2).
1. Ongoing measurements. During the experiment, some simple
non-invasive measurement of muscle function such as
monitoring the exercise capacity and/or grip strength or
Rotarod activity can be undertaken regularly (see In vivo
measurements of whole body function and muscle strength
in mice). In addition, body weights should be measured
regularly, at least at the beginning and end of the experiment.
Ideally, whole body imaging would also be performed but this
is not yet optimised and specialised equipment is required (see
Whole body imaging to measure histopathology over time).
2. Muscle strength. Study of isolated EDL and other muscles ex
vivo to measure specific force and strength is done
immediately after sacrifice (see In situ nerve stimulated
contraction of individual muscle cells).
3. Blood. At the termination of the experiment, blood should be
taken, serum prepared and stored frozen, for analysis of
serum CK enzyme activity (see Blood measurements of CK
and other proteins that leak out of damaged myofibres).
4. Muscle histology. Analysis of H&E stained transverse muscle
sections (frozen or paraffin) is essential to quantitate areas of
active muscle necrosis and regeneration, previous regeneration and (sometimes) unaffected myofibres, along with
fibrosis and other features as required (see Histological measurements to evaluate necrosis, regeneration and fibrosis).
Use of EBD (that involves injection of the dye prior to
sampling) or post-sacrifice immersion of samples in Procion
orange to indicate leakiness are both optional (see Measuring
leakiness of myofibres).
such reference data for both strains of mice would be very useful to
help validate a technique and to establish the range of values for the
grading scale. Much of this information is already published in
research papers, but needs to be collated and readily available on
websites (as outlined above). Reference data for all parameters for
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both male and female mdx and control C57Bl/10ScSn mice at one
standard age (e.g. 12 weeks), plus a range of other ages, including
measurements after exercise, would be most helpful.
The global availability of such detailed background information on
various parameters for analysis that will emerge from new collaborative networks, combined with agreement on basic core standard
procedures for analysis, should rapidly advance the comparison of data
between laboratories world-wide and accelerate research progress.
Summary
This review describes many factors that can influence the variation
between similar experiments in different laboratories (Part I) and
critically evaluates the technical and methodological approaches for
analysis (Part II) to help coordinate pre-clinical tests in the mdx
mouse. Although the mdx mouse is far from being the ideal model, its
use allows us to gain more insight into the pathology and potential
therapies for Duchenne muscular dystrophy. It is hoped that a more
coordinated effort and the joining of different expertise may help to
achieve greater and faster success in this important field. This review
makes two major recommendations. The first relates to basic standardisation of experimental approaches (Recommendation I) and the
second to minimal parameters for analysis to assist with comparison
between laboratories (Recommendation II). These guide-lines are not
intended as a limitation to experimental approach, but rather as a
suggestion to help provide more precise answers in a field with so
many expectations. It is hoped that this review and its recommendations will serve as a basis for further discussion and refinements and
as a starting point for the implementation of standard approaches for
evaluating pre-clinical studies in mdx mice.
Acknowledgments
This review arose from discussions at The first Brazilian International
Workshop on preclinical tests for drug therapies for muscular dystrophy
held in Ribeirao Preto, Brazil in late 2006. MG thanks Ian McLennan
(Dunedin, New Zealand) for stimulating discussions on animal husbandry. Current research support from various funding agencies is gratefully
acknowledged with grants to: MG and HR, from the National Health &
Medical Research Council (NH&MRC, Australia), L'Association Francaise
contre les myopathies (France) and Parent Project, Germany (Akton
Benni & Co.); KN, the US Department of Defence, National Institute of
Health, Foundation to Eradicate Dystrophy and the Jain Foundation; G L,
the Muscular Dystrophy Association (USA), NH&MRC (Australia) and the
Australian Research Council; and ADL, Telethon-Italy.
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DMD_M.1.2.007
TREAT-NMD Activity A07: Accelerate preclinical phase of new therapeutic treatment
development
Work package 7.4: Develop standardised protocols and procedures for harmonising and
accelerating pre-clinical studies (including standardised data analysis)
SOP Title
Quantification of histopathology in Haemotoxylin and Eosin
stained muscle sections
SOP (ID) Number
DMD_M.1.2.007
Version
1.0
Status
Final / May 18th, 2010
Last reviewed
May 18th, 2010
Author
Miranda Grounds
School of Anatomy and Human Biology, the University of
Western Australia, Perth, Australia
Working group
members
Hannah Radley-Crabb (School of Anatomy and Human Biology,
the University of Western Australia, Perth, Australia)
Kanneboyina Nagaraju (Research Center for Genetic Medicine,
Children's National Medical Center
111 Michigan Avenue, N.W. Washington, DC 20010)
Annemieke Aartsma-Rus (Leiden University Medical Center,
Department of Human Genetics, Leiden, The Netherlands)
Maaike van Putten (Leiden University Medical Center,
Department of Human Genetics, Leiden, The Netherlands)
Markus A. Rüegg (Biozentrum, University of Basel, Basel,
Switzerland)
Sabine de La Porte (Laboratoire de Neurobiologie Cellulaire et
Moléculaire, CNRS UPR 9040, Gif sur Yvette, France)
SOP responsible
Miranda Grounds
Official reviewer
Nagaraju Kanneboyina
Page 1 of 13
DMD_M.1.2.007
TABLE OF CONTENTS
OBJECTIVE.............................................................................................................................. 3
SCOPE AND APPLICABILITY .............................................................................................. 3
CAUTIONS............................................................................................................................... 3
MATERIALS ............................................................................................................................ 3
METHODS................................................................................................................................ 4
EVALUATION AND INTERPRETATION OF RESULTS.................................................... 6
REFERENCES.......................................................................................................................... 9
APPENDIX ............................................................................................................................. 10
Page 2 of 13
DMD_M.1.2.007
1
OBJECTIVE
This document describes a method and provides reference values for the histological
characterisation of dystrophic muscle from mdx mice.
2
SCOPE AND APPLICABILITY
Dystrophic skeletal muscle pathology can be easily assessed on haematoxylin and
eosin (H&E) stained transverse muscle sections. This SOP is written as a guide for those who
wish to quantitate the amount of skeletal muscle necrosis (as an indication of the extent of
dystropathology) and subsequent muscle regeneration in histological muscle sections from
dystrophic mdx mice.
3
CAUTIONS
As with all histological staining, great care must be taken in preparation of the
samples:
- Accurate removal of the muscle(s) from the mouse will ensure that the appropriate
portion of the appropriate muscle is actually being consistently analysed.
- Fixation (or freezing) should be done carefully to minimize shrinkage of tissues (or
ice-artifact). It is important to identify areas of ice-artifact in frozen sections and to
distinguish this from actual areas of muscle necrosis.
- Accurate embedding is essential as all sections must be embedded the same way.
Longitudinally embedded muscle samples cannot be quantitated and compared to
transverse embedded muscle samples.
- Bad cutting (e.g. a blunt blade which leaves knife scores in the section) or folding of
the muscle section can make it hard to analyse a section accurately.
4
MATERIALS
Animals: C57Bl/10mdx/mdx, Animal Resource Centre Murdoch (Western Australia
http://www.arc.wa.gov.au/) or a similar animal supplier such as Jackson Laboratories
(http://jaxmice.jax.org) in the USA.
For all: Superfrost glass slides (Lomb SF41296), Harris Haematoxylin (BDH 35194.6T),
Eosin yellowish (BDH 341973), 100% ethanol, 70% ethanol, Xylene, DPX mountant (BDH
36029.4H).
For frozen histology: Gum Tragacanth (Sigma G1128), Isopentane (BDH 103616V), Liquid
Nitrogen, 1cm3 cork squares.
For paraffin histology: 4% Paraformaldehyde (Sigma P6148).
Page 3 of 13
DMD_M.1.2.007
5
METHODS
Animal treatment:
Dystrophic mice do not necessarily need to be treated prior to assessing their
histology. However, if treatments are involved it is important to have at least 2 groups in
order to get interpretable information: for example, typically treated and untreated groups for
drug testing, exercised and unexercised groups for exercise interventions, or combinations,
depending on the experimental design.
Tissue preparation:
For paraffin histology, muscles (e.g. the tibialis anterior, TA) are removed from the
mouse and immediately placed into 4% paraformaldehyde for fixation. If possible, placing an
entire leg into paraformaldehyde will allow the muscles to hold their shape and minimise
curling of the muscles. The muscles are left in paraformaldehyde for up to 48 hours
(depending on the size of the muscle/leg) and once fixed moved into 70% ethanol. As an
alternative to paraformaldehyde, various commercially available formalin solutions (Confix –
Australian BioStain) can be used for basic muscle histology although these may not be
suitable for antibody staining. After fixation, muscles are cut transversely, caged and
processed overnight in an automatic processor e.g. Shandon, (ethanol through to paraffin wax)
and embedded into paraffin blocks.
For frozen histology, the fresh muscles are bisected transversely and the 2 pieces
mounted side-by-side on cork squares using tragacanth gum. The muscles are routinely frozen
in a slurry of isopentane cooled in liquid nitrogen. Isopentane reduces surface tension and
avoids trapping air around the muscle (which can slow the freezing process) thus producing a
better frozen sample and excellent histology (with little or no freeze artefact).
Sectioning:
Skeletal muscle tissue sections are generally cut at a width of approximately 5µm for
paraffin blocks, collected onto uncoated glass slides and stored in the dark at room
temperature until stained. Frozen sections are cut at 8µm (on a cryostat) directly onto
uncoated or silinated glass slides: ideally these are stained immediately but can be stored at 20ºC until stained.
Staining:
Dystrophic skeletal muscle pathology can be accurately assessed on sections stained
with Haematoxylin and Eosin (H&E). Haematoxylin stains eosinophilic structures (e.g.
muscle sarcoplasm) pink and Eosin stains basophilic structures (e.g. nuclei) dark purple, with
high RNA producing paler purple staining in the cytoplasm (e.g. in young myotubes) (Fig. 1).
Basic H&E staining protocol for paraffin sections. For frozen sections begin at ‘f’
a)
b)
c)
d)
e)
Dewax paraffin sections – incubate slides at 60ºC for 20minutes.
Dewax sections – Xylene wash for 3 minutes (repeat x3).
Rehydrate sections – 100% ethanol for 3 minutes (repeat x2).
Rehydrate sections – 70% ethanol for 3 minutes.
Rinse – double distilled water for 1-3 minutes.
Page 4 of 13
DMD_M.1.2.007
f)
g)
h)
i)
j)
k)
l)
Stain - Haematoxylin for 30 seconds. (Start frozen sections here)
Stain - Remove excess stain in tap water (until nuclei turn blue).
Stain - 70% ethanol for 3 minutes.
Stain - Eosin for 15 seconds.
Dehydrate – 100% ethanol for 3 minutes (repeat x3).
Clear – Xylene wash for 3 minutes (repeat x3).
Mount - DPX mountant and coverslip.
Image analysis and quantitation:
The morphological features of the muscle are identified using digital images of the
stained section. Non-overlapping images of a transverse muscle section can be tiled together
to provide a single digital image of the entire muscle cross sectional area. Morphological
features appear to be equally divided through-out the length of the muscle (e.g. from tendon to
tendon) in unexercised mdx mice (Figure 2) so it is sufficient to take pictures at one location
within the muscle i.e. the middle (van Putten et al 2010). For consistency it is necessary to
keep the location constant in all muscles and to mention which part of the muscle is analyzed.
Images are acquired using a bright-light microscope, digital camera and image capture
software (e.g. Leica DM RBE microscope, a personal computer, a Hitachi HVC2OM digital
camera, Image Pro Plus 4.5.1 software and Vexta stage movement software). More
sophisticated digital slide scanners (e.g. Aperio Scanscope) can be used to collect images
although not all researchers will have access to these machines.
Some researchers perform histological analysis on multiple single frame images taken
from the same muscle section (fields of view) and do not analyse the entire cross sectional
area. However, this method assumes that histological features are homogenous throughout the
muscle cross section, which may not always be true, in particular when mice have been
subjected to exercise.
Histological features are identified manually by the researcher and quantified using
image analysis software (e.g. Image Pro Plus). To quantitate the abundance of a specific
histological feature (e.g. myofibre necrosis) per muscle cross sectional area, first open the
digital image in your image analysis software (e.g. Image Pro Plus) and measure (draw
around) the entire cross-section area of the muscle. This can be done in pixels (sufficient
when determining a percentage) or the software can be calibrated and features measured as an
absolute value (e.g. µm2). Second, identify and manually measure (draw around) all
histological features of interest (e.g. all areas of infiltrating inflammatory cells and
fragmented muscle sarcoplasm).
Once all the appropriate histological features have been measured in the muscle cross
section, data can be exported directly into Microsoft Excel and a percentage (e.g. % myofibre
necrosis) can be calculated (e.g. Total area of all histological features/ Total area of muscle
cross section x 100). The data are then subjected to the appropriate statistical analyses.
Analysis of all morphological features at once on whole cross sections can also be
performed using the H&E colour deconvolution plugin of ImageJ (free software) which
separates the Hematoxylin component and the Eosin component. Normal, undamaged
myofibers are represented by the Eosin component and this area can be measured by
thresholding. This can distinguish undamaged from damaged/regenerating myofibres and also
Page 5 of 13
DMD_M.1.2.007
identify areas of fibrosis. The total area can be determined on the original picture (van Putten
et al. 2010).
6
EVALUATION AND INTERPRETATION OF RESULTS
Which muscle to analyse?
In all dystrophic mdx mice (both treated and untreated) it is vital to consider the age of
the mouse when assessing skeletal muscle dystropathology and to compare age-matched mice
(treated vs. untreated), since the dystrophic characteristics of muscle change with age. The
tibialis anterior (TA) muscle of the lower hindlimb is widely used, since it is readily
accessible, transverse sections are easily obtained and it contains mainly fast myofibres. The
soleus is also of interest as a model of slow myofibres. There is also a wealth of information
on other large hindlimb muscles such as the quadriceps and gastrocnemius, although these can
be more complex to interpret since a true transverse section through all myofibres is not
possible due to the architecture of some muscles. In addition, the plane of section and analysis
is complicated since these muscles are composed of several groups of muscles. e.g. the
gastrocnemius muscle has 2 heads that attach to the femur and the quadriceps muscle is
composed of 4 individual muscles, the rectus femoris, vastus lateralis, vastus intermedius and
vastus medialis.
The forelimb muscles such as triceps and biceps are also sometimes measured and
generally show similar changes with age as the hindlimb muscles.
The diaphragm is of special interest since it shows striking dystropathology, especially
in older mice (Fig. 2). Due to the complex structure and varying width of the diaphragm
muscle, extra care must be taken to cut sections at either equivalent locations for histological
comparisons or at various intervals through the diaphragm.
Exercise will also influence the muscle analysed as some muscles are severely
damaged by exercise (e.g. forced treadmill running, voluntary wheel running, swimming etc.)
whereas other muscles may be relatively unaffected by a specific exercise regime. For
example, voluntary wheel running almost doubles the amount of necrosis in the quadriceps,
whereas the TA is almost unaffected (Radley et al. 2008); similarly, various muscles in the
forelimbs and hindlimbs are differentially affected by treadmill running (Grounds et al, 2008).
The specific age of the mdx mice will greatly influence the histological features that
are seen and analysed. For example a high level of muscle necrosis (e.g. 30%) in the TA
muscle of a 23 day old mdx mouse is expected and the acute onset of muscle necrosis around
21 days of age is well documented (Grounds and Torrisi, 2004; Radley and Grounds, 2006).
However, a high level of muscle necrosis in a sedentary adult (6 weeks+) mdx mouse is not
expected. Histological features in early and later stages of the disease can be very different.
Reference data for 3 ages are shown in Tables 1 and 2 and Fig. 2.
Morphological features to be identified:
A) Normal / undamaged myofibres: Normal myofibres have peripheral nuclei, intact
sarcolemma and non-fragmented sarcoplasm (Fig 1A). In young mice these myofibres
Page 6 of 13
DMD_M.1.2.007
represent areas of muscle that have not undergone the process of necrosis and regeneration. In
adult mice these myofibres represent areas of muscle that have either never undergone
necrosis (this is the usual interpretation), or have previously undergone necrosis and
regeneration with long enough time for the myonuclei to have moved to the periphery.
B) Necrotic myofibres: Necrotic muscle is identified by the presence of infiltrating
inflammatory cells (Fig 1B) (basophilic staining) and/or hypercontracted myofibres and
degenerating myofibres with fragmented sarcoplasm (Fig 1C). Measurements of muscle
necrosis in mdx mice are of much value in (i) young mice before and after the acute onset of
myofibre necrosis, which occurs in limb muscles around 3 weeks of age and (ii) in older mice
where necrosis is induced by exercise. The measurements of active muscle necrosis are
generally of less value in sedentary adult mdx mice since the levels are very low in some limb
muscles (e.g. around 5% of whole TA muscle affected and even lower after about 1 year of
age) – thus it is difficult to detect a relative decrease in such already low values. The acute
onset of dystropathology with high levels of necrosis provides a very sensitive assay to
specifically evaluate therapeutic interventions designed to prevent or reduce myofibre
necrosis.
C) Regenerating (recently necrotic): Regenerating muscle is identified by activated
myoblasts and, 2 -3 days later, small basophilic myotubes (Fig 1D). These myotubes
subsequently mature into plump myofibres with central nuclei (regenerated myofibres) (Fig
1E).
Cumulative skeletal muscle damage in young mdx mice consists of active myofibre
necrosis plus the areas of subsequent regeneration (new myotubes/myofibres), see Table 2. It
is noted that in mdx mice up to at least one year of age, there appears to be excellent capacity
for new muscle formation in damaged limb muscles.
In older mdx mice, quantitation of undamaged myofibres, that look normal with
peripheral nuclei (Fig 1A), is a useful measure of muscle that has NOT been subjected to
necrosis (in the last 100 days)(McGeachie et al. 1993), since this indicates resistance of the
myofibres to damage. This is a quicker parameter to measure (since it is smaller, often only 510% in untreated adult mdx mice) compared with measuring the remaining bulk of tissue that
contains myofibres with central nuclei (as an indicator of previously necrotic/regenerated
tissue). If a treatment/intervention has significantly reduced the amount of necrosis then the
area of unaffected myofibres will be greater. As indicated above, in older mice where the
dystropathology has stabilised, measurements of active muscle necrosis and regeneration are
often not very informative since the background level is usually very low (however, the
specific question being addressed needs to be considered for such analyses).
D) Fibrosis and fatty connective tissue: Non-muscle tissue can be pronounced in older mdx
mice (Fig 1F). Fibrosis and fatty tissue are readily observed in H&E stained sections but
fibrosis can be emphasised by routine histochemical stains such as Van Gieson’s or Masson’s
Trichrome, and lipids by specific stains such as Oil red O. Extensive fibrous connective tissue
and some calcification are reported in limb muscles of mdx mice after about 16 months of
age: in contrast the mdx diaphragm shows significant fibrosis much earlier and this increases
Page 7 of 13
DMD_M.1.2.007
in severity with age. Fibrosis, measured biochemically by hydroxyproline content, can also be
quantitated in tissue extracts [see TREAT-NMD SOP: DMD_M.1.2.006].
E) Myofibre size: Muscle size is not widely measured for mdx mice although increased
muscle size (hypertrophy) is an initial feature of this disease and decreased muscle size
(atrophy) is seen. Muscle size can be measured as the cross sectional area of the entire muscle
or of the individual myofibres but, while accurate for true transverse sections, values are
distorted by myofibres cut obliquely (this is a major problem for clinical biopsies with
variable myofibre orientation). This problem is avoided by instead measuring the minimal
Feret’s diameter of immunohistochemically stained frozen myofibres (See SOP
DMD_M.1.2.001).
Interpretation:
The analysis of dystropathology on histological muscle sections is highly interpretive
and thus can vary slightly between individuals and laboratories: a standard set of reference
images to emphasise the precise features that are measured would help to reduce this variation
globally (Fig 1A-F). Scientific analysis that involves any degree of interpretation should be
carried out as ‘blind’ analysis using coded slides, to avoid any bias.
Potential advantages/disadvantages of methodology:
Muscles that are fixed (usually in freshly prepared 4% paraformaldehyde) and
processed into paraffin blocks can simply be stored on the shelf indefinitely. Muscle sections
from paraffin blocks are ideal for H&E analysis and for other routine histochemical stains, but
can be limiting with respect to enzymatic or antibody staining, plus poor fixation can result in
shrinkage of the muscle fibres (which is undesirable).
Frozen sections are routinely used as they avoid problems of shrinkage due to fixation
(and can thus be used to measure myofibre size), can provide excellent histology and have the
major advantage that the same tissue can be readily used for immunohistochemistry (since
many antibodies do not work well on paraffin fixed muscle sections). Disadvantages of frozen
tissues are that skill is required to prepare (to avoid ice artefact damaging the muscle sample)
and to cut sections, the tissues must be stored at -80oC and they can deteriorate over time.
Page 8 of 13
DMD_M.1.2.007
7
REFERENCES
Grounds MD and Torrisi J (2004). Anti-TNFα (Remicade) therapy protects dystrophic
skeletal muscle from necrosis. FASEB J. 18: 676-682.
Grounds MD, Radley H, Lynch GS, Nagaraju K, De Luca A (2008). Towards developing
standard operating procedures for pre-clinical research using the mdx mouse model of
Duchenne Muscular Dystrophy. Neurobiol. Disease 31: 1-19.
McGeachie JK, Grounds MD, Partridge TA, Morgan JE (1993). Age-related changes in
replication of myogenic cells in mdx mice: quantitative autoradiographic studies. J. neurol.
Sci. 119: 169-179.
Radley HG, Davies M, Grounds MD (2008). Reduced muscle necrosis and long-term benefits
in dystrophic mdx mice after cV1q (blockade of TNF) treatment. Neuromusc. Disord. 18: 227
– 238
Radley HG and Grounds MD (2006). Cromolyn administration (to block mast cell
degranulation) reduces necrosis of dystrophic muscle in mdx mice. Neurobiol. Dis. 23: 387397.
Spurney CF, Gordish-Dressman H, Guerron AD, Sali A, Pandey GS, Rawat R, Van Der
Meulen JH, Cha HJ, Pistilli EE, Partridge TA, Hoffman EP, Nagaraju K. (2009) Preclinical
drug trials in the mdx mouse: Assessment of reliable and sensitive outcome measures. Muscle
Nerve. 39:591-602.
Van Putten M, de Winter CL, van Roon-Mom W, van Ommen GJB, ’t Hoen PAC, AartsmaRus AM. (2010) A 3 months mild functional test regime does not affect disease parameters in
young
mdx
mice.
Neuromuscular
Disord.
4:
273-280.
Page 9 of 13
DMD_M.1.2.007
8
APPENDIX
Figure 1. Histological features in transverse sections of various mdx quadriceps muscles
stained with Haematoxylin and Eosin.
A) Normal myofibres (undamaged/pre-necrotic), characterised by peripheral nuclei, intact
sarcoplasm and regular shape are mainly visible. A few very small basophilic myofibres
(myotubes) with a central nucleus are also present.
B) Necrotic myofibres (inflammation) characterised by many inflammatory cells which have
infiltrated dystrophic myofibres (sarcoplasm is barely visible).
C) Necrotic myofibres (degeneration) characterised by fragmented sarcoplasm of dystrophic
myofibres with irregular shape and few myonuclei; inflammatory cells are not conspicuous.
D) Recent regeneration, shown by many small dystrophic myofibres (sometimes seen as
smaller myotubes) with central nuclei: most of these are basophilic (stain slightly purple) due
to large amounts of RNA in these actively differentiating and growing cells.
E) Regenerated myofibres, indicated by large plump mature dystrophic myofibres with central
nuclei.
F) Fat deposition in dystrophic muscle, between myofibres in the interstitial space.
Pictures A - C were all taken from a 22 day old male mdx mouse. Picture D was taken from a
28 day old male mdx mouse. Pictures E & F were taken from a 12 week old treadmill
exercised male mdx mouse. Scale bar represents 100µm.
Page 10 of 13
DMD_M.1.2.007
Page 11 of 13
DMD_M.1.2.007
Table 1. Myofibre necrosis in male mdx mice. Values shown are average percentage (%) of
whole cross sectional area +/- s.e.m. Myofibre necrosis is identified as the presence of
infiltrating inflammatory cells (Fig 1B) (basophilic staining) and/or hypercontracted
myofibres and degenerating myofibres with fragmented sarcoplasm (Fig 1C).
Age/Muscle
Quadriceps
Gastrocnemius
3 months
(12 wks)
6 months
(24 wks)
10 months
(40 wks)
6.12%
(+/- 0.6)
3.32%
(+/- 0.4)
2.23%
(+/- 1.3)
2.5%
(+/- 0.4)
N/A
(Not available)
N/A
Tibialis
Anterior
6.89%
(+/- 1.4)
N/A
Triceps
Diaphragm
Reference
8.5%
(+/- 1.4)
N/A
2.32%
(+/- 0.2)
N/A
N/A
N/A
N/A
Radley-Crabb
2010 Submitted
Radley-Crabb
unpublished
Radley-Crabb
unpublished
Table 2. Cumulative muscle damage in young mdx litters. Values shown are average
percentage (%) of whole cross sectional area +/- s.e.m. Cumulative skeletal muscle damage in
young mdx mice consists of active myofibre necrosis plus the areas of subsequent
regeneration (new myofibres).
Age/Muscle
24 days
28 days
Necrosis
5.11
(+/- 1.45)
4.56
(+/- 1.1)
Tibialis Anterior
Regeneration Cumulative damage
28.03
33.16
(+/- 7.7)
(+/- 6.7)
36.25
40.84
(+/- 9.7)
(+/- 9.4)
Reference
Radley et al 2008
Neuromus Dis 18; 227-238
Radley et al 2008
Neuromus Dis 18; 227-238
Page 12 of 13
DMD_M.1.2.007
Figure 2. Quantification of histopathology between different areas within a muscle in 16
week old mdx males. In 4 different skeletal muscles and the diaphragm, the percentage of
damage (necrosis and fibrosis) was determined at five locations (A-E) from tendon to tendon.
No significant difference was found between different areas within one muscle. Significant
differences in histopathological severity were found between different muscles; the tibialis
anterior and the diaphragm were the least and most severely affected muscles respectively.
Based on (van Putten et al. 2010).
45
Percentage fibrosis/necrosis
40
35
30
25
20
15
10
5
0
A
B
C
Qua
D
E
A
B
C
Gas
D
E
A
B
C
TA
D
E
A
B
C
Tri
D
E
A
B
C
D
E
Dia
Page 13 of 13
Neuromuscular Disorders 18 (2008) 227–238
www.elsevier.com/locate/nmd
Reduced muscle necrosis and long-term benefits in dystrophic
mdx mice after cV1q (blockade of TNF) treatment
Hannah G. Radley, Marilyn J. Davies, Miranda D. Grounds
*
School of Anatomy and Human Biology, The University of Western Australia, Anatomy and Human Biology, Perth, WA 6009, Australia
Received 18 May 2007; received in revised form 10 October 2007; accepted 1 November 2007
Abstract
Tumour necrosis factor (TNF) is a potent inflammatory cytokine that appears to exacerbate damage of dystrophic muscle in vivo.
The monoclonal murine specific antibody cV1q that specifically neutralises murine TNF demonstrated significant anti-inflammatory
effects in dystrophic mdx mice. cV1q administration protected dystrophic skeletal myofibres against necrosis in both young and
adult mdx mice and in adult mdx mice subjected to 48 h voluntary wheel exercise. Long-term studies (up to 90 days) in voluntarily
exercised mdx mice showed beneficial effects of cV1q treatment with reduced histological evidence of myofibre damage and a striking decrease in serum creatine kinase levels. However, in the absence of exercise long-term cV1q treatment did not reduce necrosis
or background pathology in mdx mice. An additional measure of well-being in the cV1q treated mice was that they ran significantly
more than control mdx mice.
2007 Elsevier B.V. All rights reserved.
Keywords: DMD; Mdx mouse; TNF; Myofibre necrosis; Voluntary exercise
1. Introduction
Duchenne muscular dystrophy (DMD) is a lethal muscle
wasting disorder, affecting approximately 1/3500 male
births [1,2]. Complete absence or impaired function of
the skeletal muscle protein dystrophin leaves dystrophic
myofibres susceptible to damage during mechanical contraction [3,4]. Consequently this initial damage progresses
to myofibre necrosis. Repeated cycles of necrosis ultimately
result in replacement of myofibres by fat and fibrotic connective tissue and loss of muscle function [2,5]. It is hypothesised that the initial myofibre damage is exacerbated by
the endogenous inflammatory response [6–9] and that
inflammatory cells and cytokines further damage the sarcolemma resulting in myofibre necrosis rather than the repair
of minor membrane lesions. There is strong evidence to
suggest that inflammatory cells and cytokines play a role
in skeletal muscle damage (reviewed in [8]) and dystrophic
*
Corresponding author. Tel.: +61 08 6488 7127.
E-mail address: [email protected] (M.D. Grounds).
0960-8966/$ - see front matter 2007 Elsevier B.V. All rights reserved.
doi:10.1016/j.nmd.2007.11.002
muscle tissue has a considerably different gene expression
pattern compared to non-dystrophic muscle with up-regulation of multiple genes involved in both the inflammatory
response and muscle regeneration [10]. Blockade or depletion of resident T cells [11], neutrophils [9], macrophages
[12,13] and mast cells [14,15] in mdx mice in vivo reduces
the severity of dystropathology.
These cells all produce tumour necrosis factor (TNF)
that is a potent pro-inflammatory cytokine that induces
chemokine expression and upregulates adhesion protein
expression on endothelial cells, resulting in cell infiltration
to sites of inflammation [16,17]. TNF was previously
referred to as TNFa, however as discussed in a recent
review [18] the renaming of TNFb as lymphotoxin (LTa
and LTb) leaves TNFa an orphan term and thus the appropriate term for use is now TNF. TNF is elevated in both
DMD and mdx mouse muscles [19–21]. Antibody blockade
of TNF with the human/mouse chimeric antibody infliximab (Remicade) in young mdx mice, results in a striking
protective effect on dystrophic myofibres and suppresses
the early acute phase of myofibre necrosis [7]. A similar
228
H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
protective effect in young dystrophic muscle was demonstrated with etanercept (Enbrel – a soluble TNF receptor)
[9]. These results strongly support a key role for TNF in
both inflammation and necrosis in dystrophic muscle. In
addition, both infliximab and etanercept treatment prevented the inflammatory response normally seen at 5 days
in whole muscle autografts in non-dystrophic C57BL/10
mice, confirming the efficacy of these drugs in mice [22].
The monoclonal antibody infliximab was generated to
block human TNF [23], and is currently very effectively
used in the treatment of Crohn’s disease and rheumatoid
arthritis [24]. There is controversy regarding the cross-species binding of infliximab based on in vitro tests, yet at least
four papers report anti-inflammatory effects in vivo in mice
[7], rats [25,26] and pigs [27]. To increase the efficacy of
TNF blockade in mice and to avoid potential problems
of immune response to the human constant domain
sequences of infliximab, a rat monoclonal antibody specific
for mouse TNF [28] and chimerized using mouse kappa
light chain and mouse 1gG2a heavy chain constant domain
sequences (cV1q) was identified [29]. The present paper
tests the effectiveness of the cV1q (mouse-specific antiTNF) antibody in both dystrophic and non-dystrophic
mice, in both short and long-term (up to 90 days) studies
combined with voluntary exercise. The use of the species
specific antibody is considered important for long-term
studies to minimise immune problems.
The mdx mouse, an animal model for DMD [30], undergoes a spontaneous onset of acute necrosis and subsequent
regeneration in limb and paraspinal muscles around 3
weeks of age [7,31]. The high level of muscle necrosis
between 21 and 28 days provides an excellent model to
study therapeutic interventions designed to prevent or
reduced muscle necrosis, as a reduction in dystropatholgy
is easily observed [7,15,32,33]. Myofibre necrosis markedly
decreases and stabilises by 6 weeks of age [34,35]. The low
level of dystropathology in adult mice can be made significantly worse by exercise that increases myofibre necrosis
and reduces muscle strength [14,36,37] enabling potential
therapeutic interventions to be evaluated in adult mdx mice
[15,38–41].
2. Experimental overview
The cV1q antibody was tested in four in vivo models of
inflammation. (1) Whole muscle autografts in adult nondystrophic C57BL/10ScSn mice (the non-dystrophic parental strain for mdx), (2) young dystrophic mdx mice (male
and female littermates), (3) adult dystrophic (mdx) mice
subjected to 48 h voluntary exercise and (4) long-term (up
to 90 days) cV1q treatment in both exercised and unexercised adult dystrophic (mdx) mice. Voluntary wheel running has several advantages compared to forced treadmill
exercise; mice are able to run at night when they are normally active [42,43] which avoids the stress associated with
exercising mice during the day when they are normally
inactive, also voluntary exercise is less stressful to the ani-
mal than forced high intensity exercise [44,45]. We also
hypothesise that the amount of voluntary exercise undertaken by an individual mouse may reflect the overall health
of the mouse and serve as an additional measure of wellbeing to test drug interventions. The effects of cV1q are
compared with infliximab and etanercept for three of the
experimental models [7,9,22].
3. Material and methods
Mice. Experiments were carried out using dystrophic
mdx litters (male and female littermates), dystrophic female
mdx (C57BL/10ScSnmdx/mdx) mice and non-dystrophic
female C57BL/10ScSn mice (Table 1). All mice were
obtained from the Animal Resources Centre (ARC) Murdoch, Western Australia, housed under a 12 h day–night
cycle and allowed access to food and water ad libitum. Mice
were treated in strict accordance with the Western Australian Prevention of Cruelties to Animals Act (1920), the
National Health and Medical Research Council and the
University of Western Australia Animal Ethics.
cV1q treatment. Intra-peritoneal (IP) injections of cV1q
and the isotype-matched, negative control antibody
(cVAM) both provided by Centocor were given at a concentration of 20 lg/g/mouse/week (Table 1). For the whole
muscle autografts (experiment 1) cV1q was routinely
injected 24 h prior to surgery (unless otherwise stated). In
young mdx mice (experiments 2 and 4) injections began
at 19 days of age. In the 48 h voluntarily exercised adult
mice (experiment 3) a single injection was given 24 h prior
to voluntary exercise.
Whole muscle autograft surgery. Whole muscle autografts were used as an in vivo bio-assay to assess the antiinflammatory properties of cV1q in non-dystrophic mice.
Six-weeks-old female C57BL/10 mice were anaesthetised
using 2% (v/v) Rodia Halothane. The extensor digitorum
longus (EDL) muscle with both tendons was removed form
the anatomical bed and transplanted onto the surface of
the tibialis anterior (TA) muscle, the EDL tendons were
sutured to the TA, the skin closed and wound left to heal,
as described in [22,46,47]. It is well-documented that this
Table 1
Animal treatment summary
Experiment
Strain
Age
(1) Whole muscle
autografts
(2) Necrosis onset
in young mdx litters
(3) 48 h voluntary
exercise
(4) Long-term cV1q
treatment
C57BL/10
6 weeks Female 1 day prior
to surgery*
d19–28 Mix
d19–d28
(weekly)
6 weeks Female 1 day prior
to exercise
d19–90 Female d19–d90
(weekly)
Mdx
Mdx
Mdx
Sex
cV1q
treatment
*
Indicates that cV1q injections were usually given 1 day prior to whole
muscle graft surgery; however some injections occurred at 1 week prior to
surgery and some at 1 week prior to surgery plus again at 5 days after
surgery.
H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
procedure severs the nerve and blood supply to the EDL
muscle and results in necrosis and subsequent inflammation and regeneration of the graft. Regeneration that
results in new muscle formation begins with infiltration
of inflammatory cells and revascularization [48–50]. This
inflammatory zone is well-established by 5 days after surgery and provides a good model to observe anti-inflammatory interventions. To test the duration of biological
efficacy, cV1q was administered at (A) 24 h prior to surgery
( 1 day), (B) one week prior to surgery, or (C) as two injections, one 24 h prior to surgery and again on day 5. Whole
muscle autografts were sampled at either 5 or 7 days after
surgery and paraffin processed for histological analysis.
Voluntary exercise. Mice were voluntarily exercised
using a metal mouse wheel (300 mm) placed inside the cage.
Exercise data were collected via a small magnet attached to
the mouse wheel with a sensor from a bicycle pedometer,
attached to the back of the cage, that recorded single wheel
revolutions, allowing total distance (km) run by an individual mouse to be determined, as per [9,15]. While an
extended period of voluntary exercise might result in additional muscle necrosis, it also allows myofibres that became
necrotic at the beginning of the exercise regime to commence regeneration resulting in new myofibre formation;
however this makes interpretation difficult against the
background pathology. We tested specifically for the effects
of exercise on myofibre necrosis in adult mdx mice and
therefore exercise was limited to only 48 h (experiment 3)
to allow for analysis shortly after the onset of exerciseinduced necrosis (and before new myotube formation –
that normally starts about 2.5 days after necrosis) [51].
Long-term voluntary exercise. Young mdx mice were
placed in cages containing exercise wheels from 28 days
of age. Before 28 days of age mice are too small to run
on the exercise wheels and mice did not run any considerable distance until 33 days of age; therefore, exercise measurements were recorded only from 33 days through to 90
days of age (8 weeks on the wheel – experiment 4). These
mice were all injected weekly with cV1q, control cVaM
or sterile water from 19 days of age.
Tissue collection and image acquisition. All mice were
sacrificed by cervical dislocation while under terminal halothane (3%v/v) anaesthesia. The TA, grafted EDL or
quadriceps muscles were sampled and prepared for paraffin
processing. Transverse sections (5 lm) were cut through
the mid-region of each muscle. Slides were stained with
haematoxylin and eosin (H&E) for morphological analysis.
Non-overlapping tiled images of transverse muscle sections
provided a picture of the entire muscle cross-section.
Images were acquired using a Leica DM RBE microscope,
a personal computer, a Hitachi HVC2OM digital camera,
Image Pro Plus 4.5.1 software and Vexta stage movement
software. Tiled images were taken at 10· magnification.
Histological image analysis. Histological analysis was
carried out on whole muscle cross-sections. Muscle morphology was drawn interactively by the researcher using
Image Pro Plus 4.5.1 software and specific histological fea-
229
tures measured as a percentage (area) of the whole muscle
section. All section analysis was done ‘blind’. Different
morphological features are quantitated in specific experiments and are therefore covered in the results section for
each individual experiment. For each individual experiment, both the left and right leg from each mouse was analysed, the numeric value (e.g., % muscle necrosis) for each
mouse (n) thus represents an average of both legs.
Serum creatine kinase assay. While under terminal
anaesthesia blood from the mdx mice (experiments 2–4)
was collected via cardiac puncture. Blood was refrigerated
overnight, centrifuged for 3 min (1200 rpm) and serum
removed. Blood serum creatine kinase (CK) analysis was
completed at the Murdoch Veterinary Hospital, Murdoch,
WA.
Statistical analysis. All statistical analysis was completed
in SPSS (SPSS 15.0 for windows). Statistical analysis for
direct comparison between two groups was performed by
unpaired Student’s t-tests. Multiple comparisons between
groups were made using a General Linear Model, univariate analysis of variance (ANOVA). A Least Significant Difference (LSD) posthoc t-test was used to identify
differences between groups. For experiments 3 and 4 there
was no difference (P > 0.7) between data from cVaM
injected and untreated control exercised mdx mice, therefore both groups were pooled to form one single exercised
‘control’ group. Significance was set at P < 0.05 for all
comparisons.
4. Results
4.1. Experiment 1: whole muscle autografts
Morphological analysis was carried out on H&E stained
transverse muscle sections of the autografted EDL at either
5 or 7 days after transplantation (Fig. 1). The area within
the centre of the graft (persisting necrotic tissue) was quantitated and expressed as a percentage (%) of the whole graft
area. The remainder of the graft consists of inflammatory
cell infiltration, surviving myofibres and regenerating muscle (myoblasts and myotubes) (Fig. 1). The extent of persisting necrotic muscle tissue is an inverse reflection of
inflammatory cell activity and new muscle formation; with
a larger area of persisting (central) necrotic tissue indicating less inflammatory cell infiltration (from the periphery).
TNF is a key pro-inflammatory cytokine involved in
inflammation that is essential for the removal of necrotic
tissue, stimulation of revascularization and activation of
muscle precursor cells for regeneration [48,50]. Therefore
it was hypothesised that blockade of TNF activity via the
administration of cV1q would delay or prevent the inflammatory process and thus subsequent regeneration of the
graft.
The average area of persisting necrotic tissue in autografts from cV1q treated mice (71.3%) sampled at 5 days
after surgery, is significantly higher than in control grafts
(35%) (Fig. 2, Table 2), therefore cV1q administration at
230
H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
Table 2
Extent of persisting necrotic muscle tissue (% of whole muscle graft)
Treatment
n
Sample day
(after
surgery)
Persisting
necrotic tissue
(% of graft)
Comment on extent
of new muscle
formation
cV1q
( 1d)
cV1q
( 1wk)
Control
cV1q
( 1d)
cV1q (·2)
Control
N=5
Day 5
71.3*
Strongly inhibited
N=6
Day 5
50.2
Inhibited
N=5
N=8
Day 5
Day 7
35
10.4
Well-established
Advanced
N=6
N=7
Day 7
Day 7
15
15.8
Advanced
Advanced
Persistent necrotic tissue inversely reflects the amount of inflammatory cell
infiltration and hence new muscle formation. cV1q treatment at 1 day
prior to surgery resulted in significantly (*) more persisting necrotic tissue
in comparison to controls grafts (P = 0.001). New muscle formation, seen
as myotubes, was also much lower in d5 cV1q treated grafts compared to
controls, reflecting the absence of the strong inflammatory response that
precedes new muscle formation.
Fig. 1. Whole muscle autografts sampled at 5 days after surgery in adult
C57BL/10 mice. Haematoxylin and eosin stained transverse sections of
grafted EDL muscle and the underlying TA muscle. The cV1q treated
graft (A) contains a much larger area of persisting necrotic muscle tissue
(NM) compared to the cVaM treated control graft (B). The active cellular
zone (periphery) of both grafts contains new blood vessels and some
surviving myofibres (SM). The cVaM treated control graft (B) contains
many small multinucleated myotubes (MT) in comparison to the cV1q
treated graft where very few myotubes are present. Bar indicates 200 lm.
1 day (P = 0.01) prior to surgery significantly impaired
inflammatory cell infiltration and subsequent muscle regeneration. cV1q intervention resulted in an approximate 2fold increase in persisting necrotic autograft tissue sampled
at 5 days after surgery. However after this time, inflamma-
tion does occur and at 7 days after surgery, autografts in
cV1q treated have a similar appearance to untreated control mice with little necrotic tissue and extensive new muscle formation in all grafts (Fig. 2 and Table 2). To test how
long the effects of cV1q persist, one single injection of cV1q
was given one week prior to surgery, resulting in a slight
improvement in day 5 grafts but this was not significant,
in striking contrast with cV1q when given 24 h prior to
surgery.
4.2. Experiment 2: young mdx mice (litters)
The onset of muscle necrosis in mdx mice shows high
levels of biological variation between both individual mice
Persisting necrotic tissue (% graft)
90
80
*
d5 control
70
d5 cV1q (-1day)
60
d5 cV1q (-1 wk)
50
d7 control
40
d7 cV1q (-1 day)
30
d7 cV1q (x2)
20
10
0
d5 grafts
d7 grafts
Fig. 2. Persistent necrosis (as% of graft area) in grafts sampled at 5 or 7 days after surgery. The extent of persisting necrotic tissue inversely reflects the
amount of inflammatory cell infiltration and new muscle formation. cV1q treatment 1 day (*) significantly (P = 0.01) reduced inflammation in grafts
sampled at 5 days after surgery. Error bars represent standard error. N = 5, 6, 5, 8, 6 and 7 grafts (1 graft per mouse), respectively.
H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
and litters [15]. To help reduce biological variation, cV1q
injections were performed on half a litter of mdx pups with
the remaining half of the litter used as controls (cVaM
injected). This ensures that comparisons (cV1q vs. cVaM)
are made between age matched mice of one litter to reduce
inter-litter variation [15]. Mice were sampled on either day
24 or day 28: mice sampled on day 24 were injected once
(d19) and mice sampled on day 28 were injected twice
(d19 and d26).
Muscle necrosis was identified and measured on H&E
stained transverse sections of TA muscles by the presence
of infiltrating inflammatory cells (basophilic staining) and
degenerating myofibres with fragmented sarcoplasm.
Regeneration occurs in response to necrosis and results
(2–3 days later) in myotubes that, over time, mature into
myofibres with central nuclei. Cumulative skeletal muscle
damage in young mdx mice (up to 28 days of age) consists
of active myofibre necrosis (usually only necrosis present at
24 days) plus the areas of subsequent myotube formation
(present by 28 days).
cV1q treatment significantly reduced (P = 0.03) cumulative muscle damage in the TA of 24 and 28-day-old mdx
mice (Fig. 3), with cV1q treatment, cumulative muscle
damage in 24-day-old mice was half that seen in control
(14.59%) vs. (33.16%) in an entire cross-sectional area
(CSA) of the TA. Further examination of cumulative muscle damage at day 24 (Fig. 3) indicates a delayed onset of
necrosis after cV1q treatment, as reflected by a significantly
higher proportion of necrosis (P = 0.01) and a lower proportion of regeneration (P = 0.01) in cV1q treated mice.
That is, given there are a more regenerating myofibres
(new myotubes) in the control mice and since myotubes
first form from about 2.5 days after necrosis [51]; the onset
of necrosis may have occurred earlier in the control mice.
231
While the process of muscle necrosis and hence regeneration was delayed after cV1q administration, there were
no adverse effects on new muscle formation. Similarly no
adverse effects on muscle formation were seen after the
administration of infliximab [7] or etanercept [9].
Analysis of serum CK levels showed a clear trend for
reduced (roughly halved) serum CK levels in young
cV1q-treated mdx mice compared with controls [day 24
cV1q treated, 3330 U/L (±378); cVaM treated, 6300
(±3676). Day 28 cV1q treated, 1200 U/L (±608); cVaM
treated, 2950 (±1484)], although due to high variation
the effect of cV1q treatment on serum CK levels was not
statistically significant.
Due to the well-documented high variation in severity of
dystropathology between individual mdx mice, a direct
comparison of infliximab and cV1q data for mdx mice treated at the onset of myofibre necrosis (i.e., percentage muscle necrosis) is inconclusive, that is absolute numbers
cannot be compared. A comparison of ‘relative change’ is
more appropriate with both infliximab [7] and cV1q producing 2-fold reduction in myofibre necrosis in 24-dayold mdx mice. cV1q treatment also demonstrated a strong
protective effect (2-fold) at 28 days of age (Fig. 3) whereas
infliximab treatment showed less benefit at this later timepoint. It is concluded that both treatments were highly
effective in reducing the extent of myofibre necrosis in
young mdx mice.
4.3. Experiment 3: 48 h voluntary exercise of adult mdx mice
After 48 h of voluntary wheel, the quadriceps muscle of
6-week-old female mdx mice were analysed histologically.
Measurements were calculated for: the area occupied by
necrotic myofibres, regenerating (small myotubes) and
Muscle damage (% TA)
60
50
cV1q necrosis
40
cV1q
regeneration
30
* !#
*
cVaM (control)
necrosis
20
cVaM (control)
regeneration
10
0
24 cV1q
24 control
d28 cV1q
d28 control
Fig. 3. Cumulative muscle damage in cV1q treated mdx mice aged 24 and 28-days-old compared with control (cVaM) littermates. cV1q treatment
significantly reduced (P = 0.03) cumulative muscle damage in the TA muscle of both 24 and 28-day-old dystrophic mice, as indicated by *. ! shows
significantly more necrosis. # indicates significantly less regeneration in cV1q treated 24-day-old mice, this reflects the delay in muscle damage and thus
absence of new muscle formation (short-term up to day 28). Each time-point consists of 2 litters (cV1q treated and untreated litter mate controls). Error
bars represent standard error and n = 7, 6, 6 and 5 mice, respectively, across the groups.
232
H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
regenerated (myofibres with central nuclei) muscle and
unaffected/intact myofibres as described previously [15].
100% of the muscle cross-sectional area (CSA) is made
up of necrotic tissue, regenerating tissue (recently necrotic),
regenerated tissue (central nuclei), unaffected tissue (never
been necrotic) and connective tissue. As reported previously, exercise-induced necrosis in the TA muscle of all
mice was minimal and therefore only the quadriceps muscle
was analysed [9,15,41]. No differences were found between
control cVaM treated or untreated control exercised mdx
mice, therefore for the purpose of statistical and graphical
analysis both groups were pooled and are presented as one
single exercised ‘control’ group (n = 6).
Voluntary exercise (48 h) resulted in a significant
(P = 0.02) increase (approximately 2-fold) in muscle necrosis in the quadriceps muscle of control mdx mice (Fig. 4) as
reported previously [9,15,41]. cV1q treatment significantly
reduced (P = 0.04) the extent of exercise-induced myofibre
necrosis from 12.93% (control) down to 7.58%; this low
level of necrosis is similar (not significantly different) to
the necrosis (6.22%) in unexercised quadriceps muscle of
adult mdx mice. cV1q administration did not prevent an
increase in serum CK levels after 48 h exposure to voluntary exercise (data not shown).
The mdx mice ran between 4.13 and 14.45 km over the
48 h; the average distance run by the control mice was
9.03 km and the average distance run by the cV1q treated
mice was 8.27 km. It is important to note that the cV1q
treatment had no significant affect on the distance run by
the mice over 48 h, since reduced activity might result in
reduced muscle damage (% necrosis). However, the distance run by an individual mouse does not directly corre-
Unexercised
late to either the percentage of myofibre necrosis in the
quadriceps muscle or serum CK (data not shown) this is
also documented in previous studies [9,15].
4.4. Experiment 4: long-term cV1q treatment in exercised
and unexercised dystrophic (mdx) mice
The main objective of this study was to assess long-term
treatment with an antibody to TNF in muscles of dystrophic mdx mice. cV1q treatment began at 19 days of age
(before the onset of myofibre necrosis) and adult female
mdx mice were sampled at 90 days of age (approximately
3 months). No differences were found between cVaM treated or untreated mice, therefore for the purpose of statistical analysis both groups were pooled and are presented as
one single ‘control’ group (n = 6 unexercised mice or n = 8
exercised mice). Histological analysis was performed as per
experiment 3.
cV1q treated mdx mice ran more than control mdx mice
over the 8 week period (35–90 days of age) of voluntary
exercise. cV1q treated mice ran significantly more
(P = 0.03) in total distance (383.7 km n = 8) compared
with control mdx mice (236.6 km n = 8) (Fig. 5, Table 3)
and cV1q treated mice also ran more on average each week
(Fig. 6) with the largest difference seen in weeks 2–5.
In unexercised mdx mice cV1q treatment had no effect
on the amount of active muscle necrosis in the TA (data
not shown) or in the quadriceps muscles (Fig. 7). Voluntary
exercise over the 8 week period caused a significant
(P = 0.017) increase in myofibre necrosis in the quadriceps
muscle of mdx mice (Fig. 7) in comparison to unexercised
mdx mice. cV1q treatment in long-term exercised mice
Control + EX
cV1q + EX
18
Muscle necrosis (% of Quad)
16
**
14
12
*
10
8
6
4
2
0
6 week old mdx mice
Fig. 4. Muscle necrosis in exercised adult mdx mice. Muscle necrosis in the quadriceps muscle of unexercised (6.22%), exercised control (12.93%) and cV1q
treated exercised adult mdx mice (7.58%). The analysis of variance test for multiple comparisons between groups showed statistical difference (P = 0.04).
Posthoc LSD T-tests showed a significant (P = 0.02) increase in skeletal muscle necrosis in the quadriceps muscle of exercised control mdx mice in
comparison to unexercised mice (**) and a significant (P = 0.04) reduction in necrosis in cV1q treated exercised mdx mice in comparison to control
exercised mice (*). Exercised ‘control’ refers to 3 cVaM treated and 3 untreated mice. Error bars indicate standard deviation, horizontal bar indicates
exercised mice and n = 5, 6 and 6 mice, respectively.
H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
Control + EX
233
cV1q + EX
Avg Total km run (9 weeks on wheel)
500
*
450
400
350
300
250
200
150
100
50
0
90 day old mdx mice
Fig. 5. Total distance run (km) over 8 weeks (between 35 and 90 days of age). The analysis of variance test for multiple comparisons between groups
showed statistical difference (P = 0.03). Posthoc LSD T-tests showed a significant increase (P = 0.03) in the total distance run by cV1q treated mice in
comparison to control mice (*). ‘Control’ refers to pooled data from cVaM treated (n = 4) and untreated (n = 4) exercised mdx mice. Error bars represent
standard error and n = 8 mice for both groups.
Table 3
Summary – total distance run (km) over 8 weeks by cV1q treated and control exercised mice, combined with the data for % necrosis in the quadriceps
muscle and serum creatine kinase (CK) levels
Number
Treatment
Distance (Avg. km/wk)
Distance (total km/8wk)
Necrosis (quad% area)
CK (U/L)
Unaffected tissue (quad% area)
1
2
3
4
5
6
7
8
cVaM
cVaM
cVaM
cVaM
Untreated
Untreated
Untreated
Untreated
34.4
54.5
35.2
12.7
30.2
19.4
29.2
21.1
275.1
435.7
282.0
101.6
241.7
154.7
233.5
168.8
15.85
12.9
6.15
3.75
8.38
10.27
13.25
11
39900
9850
33600
24150
17350
16100
3600
6250
2.1
1.7
2.6
2.4
3.4
1.15
1.2
1.2
Average
Control
29.6 km/week
236.6 km total
10.19%
18850 U/L
1.95%
1
2
3
4
5
6
7
8
cV1q
cV1q
cV1q
cV1q
cV1q
cV1q
cV1q
cV1q
21.2
42.0
71.6
72.8
36.3
62.2
41.15
36.3
169.6
336.0
572.6
582.75
290.8
497.9
329.2
290.9
2.74
11.75
4.13
11.95
4.41
8.07
6.65
8.74
7250
5300
2100
4550
2300
2150
2500
3600
4.8
2.7
6.8
5.4
2.65
5.7
4.4
3.8
Average
cV1q
47.9*
383.7*
7.31
3718*
4.53*
*
Indicates a significant difference (P < 0.05) between control and cV1q treatment.
caused a reduction in the percentage of active myofibre
necrosis in the quadriceps muscle, however this effect was
not significant. It is interesting to note that the percentage
of active muscle necrosis seen after 8 weeks of exercise is
similar to that seen after 48 h voluntary exercise (10.2%
vs. 12.9%) indicating no adaptation to the repeated exercise
regime.
In unexercised adult mdx mice cV1q treatment had no
effect on serum CK (Fig. 8). In striking contrast, voluntary
exercise over the 8 week period (day 35–90) caused a
dramatic (10-fold) and significant (P = 0.01) increase in the
level of serum CK (Fig. 8, Table 2) whereas in cV1q treated
mdx mice serum CK after exercise was significantly lower
(P = 0.01) and, in fact, was the same as unexercised mdx
mice. These data suggest that long-term voluntary exercise
significantly increases myofibre ‘leakiness’ of control mdx
muscle (although has no striking impact on the amount of
myofibre necrosis) and this is prevented by cV1q treatment.
It was originally hypothesised that cV1q treatment
would reduce the overall extent of dystropathology as
234
H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
Control + EX
cV1q + EX
100
Avg KM run per group per week
90
80
*
70
60
50
*
40
30
20
10
0
wk 0-1
wk 1-2
wk 2-3
wk 3-4
wk 4-5
wk 5-6
wk 6-7
wk 7-8
Fig. 6. Average distance run (km) per week for cV1q treated and control mice. (*) indicates a significant difference in average distance run during a specific
week between cV1q treated and control mdx mice. ‘Control’ refers to pooled data from cVaM treated (n = 4) and untreated (n = 4) exercised mdx mice.
Error bars represent standard deviation and n = 8 mice for both groups.
Control
cV1q
Contro + EX
cV1q + EX
Muscle necrosis (% of Quad)
15
*
10
5
0
90 day old mdx mice
Fig. 7. Muscle necrosis in 90-day-old mdx mice. Muscle necrosis in the quadriceps muscle of unexercised (4.65%), unexercised + cV1q (5.27%) exercised
control (10.19%) and exercised + cV1q (7.3%) adult mdx mice. The analysis of variance test for multiple comparisons between groups showed statistical
difference (P = 0.04). Posthoc LSD T-tests showed a significant increase (P = 0.017) in muscle necrosis after exercise (*), yet cV1q treatment did not
significantly reduce muscle necrosis in exercised mdx mice. ‘Control’ refers to pooled data from cVaM treated and untreated mice. Error bars represent
standard deviation, horizontal bar indicates exercised mice and n = 6, 6, 8 and 8 mice, respectively.
measured by quantitating the number of unaffected (normal) myofibres without central nuclei – an inverse representation of disease severity. Long-term voluntary
exercise significantly (P = 0.016) reduced the amount of
unaffected myofibres indicating a more severe dystropathology in comparison with unexercised mdx muscle. In
contrast, cV1q treatment significantly maintained
(P = 0.001) the area of unaffected myofibres in exercised
adult mdx mice in comparison to control exercised mice
(Fig. 9, Table 3). This suggests that cV1q treatment
reduced the severity of dystropathology and protects
against myofibre damage in long-term exercised mdx mice.
5. Discussion
These data from short and long-term studies demonstrate that cV1q antibody treatment, which neutralises
murine TNF, reduces the extent of muscle necrosis in both
young and exercised adult dystrophic mice and supports
the original hypothesis that TNF contributes to the necrosis of dystrophic muscle. cV1q treatment also significantly
increased the amount of voluntary exercise completed by
dystrophic mice over 8 weeks, strongly indicating longterm functional benefits combined with reduced disease
severity in the mdx mice.
H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
Control
cV1q
control + EX
235
cV1q + EX
Serum creatine Kinase (U/L)
28000
**
24000
20000
16000
12000
8000
*
4000
0
90 day old mdx mice
Fig. 8. Blood serum creatine kinase level in 90-day-old mdx mice. Creatine kinase in unexercised control (2900U/L), unexercised + cV1q (2200 U/L),
exercised control (18850 U/L) and exercised + cV1q (3718 U/L) adult mdx mice. The analysis of variance test for multiple comparisons between groups
showed statistical difference (P = 0.02). Posthoc LSD T-tests showed a significant increase (P = 0.012) in CK levels after exercise (**) and no significant
increase (P = 0.013) in CK level after exercise when treated with cV1q (*). Error bars indicate standard error, horizontal bar indicates exercised mice and
n = 6, 6, 8 and 8 mice, respectively.
Control
cV1q
Control + EX
cV1q + EX
8
Unaffected tissue (% of quad)
7
*
6
5
4
**
3
2
1
0
90 day old mdx mice
Fig. 9. Unaffected muscle tissue in 90-day-old mdx mice. Unaffected myofibres in the quadriceps muscle of unexercised control (7.32%), unexercised cV1q
(6.97%), exercised control (1.95%) and exercised cV1q (4.53%). The analysis of variance test for multiple comparisons between groups showed statistical
difference (P = 0.05). Posthoc LSD T-tests showed a significant (P = 0.016) reduction in unaffected myofibres after voluntary exercise in comparison to
unexercised mice (**) and a significant (P = 0.001) increase in unaffected myofibres with cV1q treatment in exercised mice in comparison to ‘control
exercised mice (*). ‘Control’ refers to pooled data from cVaM treated and untreated exercised mice. Error bars represent standard deviation, horizontal
bar indicates exercised mice and n = 6, 6, 8 and 8 mice, respectively.
The anti-inflammatory property of cV1q in mice was
assessed using the whole muscle autograft model. Transplantation of whole muscle autografts produces a very
large mass of avascular necrotic tissue and a situation of
extreme pathology, which does not directly compare to
the small foci of damage in dystrophic muscle, but the
autograft model is useful to test the ability of cV1q to block
TNF and the inflammatory response in vivo. The use of
such an assay is also important to confirm the bioactivity
of different batches of anti-inflammatory drugs in mice
in vivo. The cV1q effect on autografts at 5 days (71.3% persistent necrosis) is similar to that seen with infliximab
(64%) and etanercept (77.4%) [22]. The anti-inflammatory
effects of infliximab and cV1q appear indistinguishable in
this autograft model and TNF blockade by both drugs
(and also etanercept) prevents this acute inflammatory cell
response in vivo. cV1q treatment significantly inhibits the
early stages of inflammatory cell infiltration (up to day 5)
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H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
but does not prevent the long-term response of inflammation that precedes regeneration (day 7 onwards): this is
presumably due to some activation of a secondary proinflammatory pathway that is unaffected by TNF inhibition. Infliximab and etanercept treatment similarly showed
no effect in day 7 grafts compared with controls [22]. This
bio-assay (at day 5) confirms an inhibitory effect of cV1q
on the inflammatory response in mice in vivo as anticipated.
The inhibitory effect was reproducible across multiple
grafts and it was specific – in that no effects were seen with
control cVaM administration.
In dystrophic mdx mice, cV1q treatment significantly
reduced the severity of skeletal muscle necrosis in both
the TA of young mdx mice and the quadriceps muscle of
exercised adult mdx mice. The voluntary wheel exercise significantly (P = 0.02) increases (approximately 2-fold) myofibre necrosis in the quadriceps of adult mdx mice and this
was prevented by pre-treatment of the mice with cV1q.
These short-term studies further validate the important
principle that necrosis of dystrophic myofibres can be
reduced by exogenous manipulation of the potent proinflammatory cytokine TNF.
A striking 10-fold increase in serum CK levels in mdx
mice after 8 weeks of voluntary exercise was completely
prevented by cV1q treatment, dramatically emphasizing
the benefits of this long-term treatment. The serum CK levels in the long-term exercised mdx mice (18,850 ±
12,933 U/l) are very high; but do fall within the range
reported by Granchelli (2000) (17,200 ± 11,600 U/l) after
2 · week 30 min exercise sessions on a horizontal treadmill
for 4 weeks [38].
Long-term treatment with cV1q in the absence of
exercise had no significant effect on myofibre necrosis
or extent of undamaged myofibres in the quadriceps
muscle, or on serum CK levels. This indicates that the
already low level of background dystropathology in adult
mdx mice is not further reduced by cV1q blockade of
TNF, whereas cV1q clearly protected against exerciseinduced acute myofibre damage. The lack of effect in
the unexercised mice is in striking contrast to the cV1q
effects seen in exercised mdx mice: whether this reflects
differences in the cellular responses in these two situations is unknown. cV1q treatment in long-term exercised
mdx mice showed (1) Improved muscle function – demonstrated by an increase capacity for exercise. (2)
Reduced myofibre leakiness – demonstrated by a marked
reduction in serum CK levels. (3) Reduced disease severity (myofibre necrosis and regeneration cycles) – demonstrated by a greater area of unaffected myofibres at 90
days of age.
It is calculated that myonuclei of newly regenerated
myofibres remain in a central location for about 50–100
days in mdx mice and thereafter 3–4% of myonuclei move
to a peripheral subsarcolemmal position every 100 days
i.e., numbers of central myonuclei may decline after 100
days of age [34]. In this study, we presume that the central
nucleus of a regenerated myofibre takes more than 70 days
to move to the periphery (from day 21 with the acute onset
of necrosis to day 90 when mice are sampled): thus it is
considered that myofibres with only peripheral nuclei have
never undergone myonecrosis and are deemed unaffected.
Counting such non-centrally nucleated (unaffected) myofibres provides a converse indication of disease severity and
supports the notion of reduced myofibre damage in longterm cV1q treated mdx mice. In contrast, myofibres with
central nuclei must have regenerated at least once. If an
individual myofibre undergoes multiple cycles of necrosis
and regeneration the appearance is similar to a myofibre
that has only regenerated once during the period of the
experiment and thus this analysis does not give any indication of how many times an individual myofibre (with central nuclei) may have undergone cycles of necrosis and
regeneration [51]. It seems likely that the cycles of necrosis
and regeneration may have occurred more often in the control mice compared with cV1q treated mdx mice, although
this cannot be determined from this basic histology.
In summary, the results achieved with cV1q administration in mdx mice are highly promising and similar to the
results achieved by both infliximab and etanercept administration and are strongly supported by the long-term studies combined with exercise. The optimal regime and
dosage, as well as the most appropriate humanised antiTNF drug needs to be selected for possible trials in
DMD boys. The development of new TNF drugs such as
TNF-TeAb (miniantibodies) further complicate the decision [52]. The study confirms that a drug intervention
may not show benefits in unexercised mdx mice, thus exercise appears an essential part of the ‘standard operating
procedure’ to effectively test the effects of anti-inflammatory drugs in mdx mice in vivo.
Direct comparisons of cV1q treatment with other
drugs such as HCT 1026; which appears to be more beneficial than prednisolone [53] is highly desirable as a prelude to selecting the most promising drug for potential
clinical trials. There are many differences between the
protocols used in our mdx study and the long-term study
of HCT 1026 in mdx mice [53]: these include the duration of voluntary exercise; time (age) of animal sampling;
and onset the of drug administration – with the crucial
issue being that HCT 1026 treatment began after the
acute onset of myofibre necrosis that starts at 21 days
in mdx mice [7]. It appears that the gender of mdx mice
also affects the severity of the dystropathology at different ages [54] although this has not been widely recognised and the impact on drug evaluations at different ages
is unclear. We used female adult mdx mice in the present
study; however other experiments show a similar level of
background necrosis and the same doubling of exerciseinduced myonecrosis in male mdx mice at 8 weeks of
age, with similar high biological variation (Radley and
Grounds unpublished data). It is noted that there are
numerous studies where the gender of the mice is not
stated [53,55,56] or a mix of both male and female mice
are used [38]. Such fundamental differences emphasise the
H.G. Radley et al. / Neuromuscular Disorders 18 (2008) 227–238
need for standard operating protocols to facilitate important comparisons of promising therapies.
Acknowledgements
We gratefully acknowledge Centocor (USA) for providing the cV1q anti-mouse TNF antibody and the negative
control antibody (cVaM) and Dr. David Shealy for his support and helpful comments regarding this work.
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1
A single 30 minute treadmill exercise session is suitable for ‘proof-of concept studies’ in
adult mdx mice: a comparison of the early consequences of two different treadmill
protocols.
Hannah Radley-Crabb1*, Jessica Terrill1,2, Thea Shavlakadze1, Joanne Tonkin1, Peter
Arthur2, Miranda Grounds1.
1
School of Anatomy and Human Biology. The University of Western Australia, Crawley,
Australia.
2
School of Biomedical, Biomolecular and Chemical Sciences. The University of Western
Australia, Crawley, Australia.
*Corresponding author:
Hannah Crabb,
Mail Delivery M309
School of Anatomy and Human Biology
The University of Western Australia
35 Stirling Highway, Crawley
Western Australia, 6009
Tel: +61 8 6488 7127
Fax: +61 8 6488 1051
E-mail: [email protected]
2
Abstract
The extent of dystropathology in sedentary adult mdx mice is very low and exercise is
often used to increase myofibre necrosis. The early events in dystrophic muscle and blood
in response to exercise (leading to myofibre necrosis) are unknown. This study describes
in detail two standardised protocols for the treadmill exercise of mdx mice and profiles
changes in molecular and cellular events after a single 30 minute treadmill session
(Protocol A) or after 4 weeks of treadmill exercise (Protocol B). Both treadmill protocols
increased multiple markers of muscle damage. We conclude that a single 30 minute
treadmill exercise session is a sufficient and conveniently fast screening test and could be
used in ‘proof-of-concept’ studies to evaluate the benefits of pre-clinical drugs in vivo.
Myofibre necrosis, blood serum CK and oxidative stress (specifically the ratio of oxidised
to reduced protein thiols) are reliable markers of muscle damage after exercise; many
parameters demonstrated high biological variation including changes in mRNA levels for
key inflammatory cytokines in muscle. The sampling (sacrifice and tissue collection) time
after exercise for these parameters is critical. A more precise understanding of the
changes in dystrophic muscle after exercise aims to identify biomarkers and new potential
therapeutic drug targets for Duchenne Muscular Dystrophy.
Key Words
mdx mouse, treadmill exercise, skeletal muscle damage, myofibre necrosis, creatine
kinase, inflammation, oxidative stress.
3
Introduction
Duchenne Muscular Dystrophy (DMD) is an X-linked, lethal muscle wasting disorder that
affects mainly boys [1, 2]. Impaired function or absence of the sub-sarcolemmal protein
dystrophin, renders dystrophic myofibres susceptible to sarcolemma damage in response
to contraction [3-7]. This initial damage can progress to myofibre necrosis and subsequent
regeneration; repeated cycles of necrosis ultimately result in the replacement of
myofibres with fat and/or fibrotic connective tissue [8]. A progressive loss of muscle mass
and function in DMD leads to premature death often due to respiratory or cardiac failure
[9]. While the genetic defect was identified over 20 years ago the specific cause of
myofibre necrosis is still unknown, although increased levels (or dysregulation) of
inflammation, oxidative stress and intracellular calcium are all heavily implicated [7, 1016].
Mdx mice (C57Bl/10ScSnmdx/mdx) which also lack dystrophin, are an animal model for DMD
and are widely used in pre-clinical research [11, 17]. In sedentary adult mdx mice, the
extent of dystropathology is relatively mild, with usually <6% myofibre necrosis in the
quadriceps muscle (expressed as % of cross sectional area [CSA]) and relatively low serum
creatine kinase (CK) activity – a marker of myofibre leakiness. Exercise is routinely used to
increase mdx myofibre damage and increase serum CK levels [18-21] thus enabling
potential therapeutic interventions to be more rigorously evaluated in vivo [22-27].
4
In the past, our laboratory has used voluntary wheel exercise over 48 hours to increase
myofibre necrosis and histology to demonstrate the benefits of anti-inflammatory drugs
on dystrophic muscle in vivo [26-28]. Muscle necrosis is roughly doubled (~6 to 12% CSA)
in quadriceps muscle after 48 hours of voluntary exercise, although other muscles such as
the tibialis anterior (TA) are barely affected by voluntary exercise [11, 24, 26].
A widely used alternative to voluntary (usually nocturnal) exercise is controlled treadmill
running (experiments usually conducted during the day). This occurs at a controlled speed
for a pre-determined length of time, thus eliminating some of the behavioural variables
experienced with voluntary exercise. A protocol of 30 minutes treadmill running on a
horizontal treadmill at a speed of 12 m/min, twice a week for at least 4 weeks, causes a
significant increase in the dystropathology of adult mdx mice and is widely used in preclinical research [20, 22, 25, 29, 30]. There is however some concerns regarding treadmill
exercise because mdx mice can have problems coping and thus be reluctant to run,
although a short warm-up period at a slower speed appears to help with treadmill running
[23].
A single 30 minute treadmill exercise session represents a precise amount of controlled
exercise that allows the time-course of early cellular and molecular events to be
measured. It is of fundamental interest to determine the extent of the initial skeletal
muscle damage and associated molecular changes in response to a single 30 minute
exercise session (Protocol A) in unexercised mdx mice, compared with mice exercised for
5
4 weeks (Protocol B) which is a widely used exercise regime [20, 22, 25, 29, 30]. In the
present study parameters measured included; histological quantification of myofibre
necrosis, circulating blood CK activity, quadriceps muscle gene expression levels (mRNA)
of the pro-inflammatory cytokines interleukin-1 β (IL-1β), interleukin-6 (IL-6) and tumour
necrosis factor (TNF) and quantification of oxidative stress (protein thiol oxidation ratio
and malondialdehyde (MDA) quantitation) in the quadriceps muscle.
The aims of the present study were to: 1) develop a short (30 minute) and repeatable in
vivo treadmill protocol to increase myofibre necrosis in adult mdx mice; 2) profile the time
course of multiple indicators of muscle damage immediately after a single exercise
session; 3) compare these responses after a single treadmill exercise session to responses
after 4 weeks of treadmill exercise; 4) establish if a single 30 minute exercise session is an
appropriate protocol to increase muscle damage in adult mdx mice and thus be used in
pre-clinical ‘proof-of-concept’ studies; 5) identify key parameters with potential as
diagnostic biomarkers to rapidly monitor efficacy of pre-clinical drug treatments.
Materials and Methods:
1) Animal procedures:
All experiments were carried out on 8 - 12 week old (adult) male non-dystrophic control
C57Bl/10 and dystrophic mdx mice; mice were obtained from the Animal Resource Centre,
Murdoch, Western Australia. They were maintained at the University of Western Australia
6
on a 12-h light/dark cycle, under standard conditions, with free access to food and
drinking water. Mice of each strain were caged in groups of 3-4. All animal experiments
were conducted in strict accordance with the guidelines of the National Health and
Medical Research Council Code of practice for the care and use of animals for scientific
purposes (2004) and the Animal Welfare act of Western Australia (2002) and were
approved by the Animal Ethics committee at the University of Western Australia.
1A) Establishing the 30 minute treadmill protocol:
Based on previous research [20, 22, 29] and as per the TREAT-NMD recommended
standard protocol “Use of treadmill and wheel exercise for impact on mdx mice
phenotype M.2.1_001” http://www.treat-nmd.eu/research/preclinical/SOPs/ a treadmill
exercise regime consisting of 30 minutes treadmill running at a speed of 12m/minute was
used. The rodent treadmill was an Exer 3/6 from Columbus Instruments (USA). Treadmill
Setup: Individual running lanes were separated by clear Perspex dividers so that the mice
could see each other while exercising. The treadmill was horizontal (0° incline) and mdx
mice were run in groups of 3 or 4 as it is time consuming and inefficient to run mdx mice
individually. Exercise Protocol: Groups of 3 or 4 mdx mice were all (1) settled for 2 mins
with the treadmill belt stationary, (2) then acclimatized with gentle walking for 2 mins at a
speed of 4m/min, followed immediately by (3) a warm–up of 8 minutes at 8m/min and
then (4) the main exercise session for 30 minutes at 12m/min. If during the 30 minutes
exercise session a mouse fatigued and could no longer run, the procedure was as follows:
turn the treadmill belt off and give all mice a 2 minute rest, turn the belt on at 4m/min for
7
2 minutes, increase the speed to 12m/minute and run for the remainder of the 30
minutes. Repeat this process if fatigue occurs again (up to 5 times for an individual mdx
mouse).
1B) Treadmill regime and Animal Sample Groups:
All experiments (exercise and sampling) were started at 8am and completed by 11am each
day.
Exercise protocol A (a single 30 minute treadmill exercise session): 12 week old
(completely untrained) mdx and control C57Bl/10 mice were exercised for a single session
on the rodent treadmill. All male mice were sampled at 12 weeks of age. The following 3
groups of male mdx mice were used for histological analysis: 1) unexercised, 2) mice
subjected to 30 minutes treadmill exercise and sampled 24hrs or 3) 48hrs post exercise.
Numerous treadmill exercise experiments have been conducted in our laboratory over the
last year using 12 week old male mdx mice and the histological data from all experiments
were pooled to provide large group numbers (see Table 1). The time course study was
conducted on a total of 32 mdx and 16 C57Bl/10 12 week old mice representing 6
different groups with n=8 for each group. The following groups were used: 1) unexercised
C57Bl/10, 2) exercised C57Bl/10 sampled immediately (0 mins) post exercise, 3)
unexercised mdx, 4) exercised mdx sampled immediately (0 mins) or 5) 2hrs exercise or 6)
24hrs post exercise (see Table 1).
8
Exercise protocol B (4 weeks of treadmill exercise): mdx mice were exercised on the
treadmill twice a week for 4 weeks, with a consistent 72 or 96hr break between each
exercise session. The treadmill exercise started when mice were 8 weeks old and
therefore all mice were 12 weeks old at time of sampling. For consistency all mice had a
72hr (3 day) break before the final (8th) exercise session and subsequent sampling. The 4
week treadmill exercise protocol was conducted on a total of 32 male mdx mice
representing 4 different groups with n=8 for each group. The following groups were used:
1) unexercised mdx, 2) 4 week exercised mdx sampled immediately (0 mins) or 3) 24hrs or
4) 96hrs post exercise (see Table 1).
1C) Forelimb grip strength: Mice from Protocol B were also assessed throughout the study
for forelimb grip strength (measured 24hrs prior to 1st, 5th and 8th treadmill exercise
session). Grip strength was measured using a Chatillon Digital Force Gauge (DFE-002) and
a triangle metal bar, as per the TREAT-NMD recommended standard protocol “Use of grip
strength meter to assess limb strength of mdx mice – M.2.2_001” http://www.treatnmd.eu/research/preclinical/SOPs/. In brief, the mouse was placed on the front of the
triangle bar (attached to a force transducer) and pulled gently until release. Each mouse
underwent 5 consecutive grip-strength trials; the grip strength value for each mouse was
recorded as the average of the 3 best efforts. Average grip strength was then normalized
for body weight [Force (kg)/BW (g)]. Change in normalised grip strength was determined
by subtracting normalised grip strength (8 weeks) from normalised grip strength (12
weeks) [25].
9
2) Tissue collection and image acquisition:
All mice were sacrificed by cervical dislocation while under terminal anaesthesia (2%v/v
Attane isoflurane Bomac Australia). Various muscles were collected, some were
immediately snap frozen in liquid nitrogen for molecular analysis (quadriceps) and some
were prepared for histology (quadriceps, triceps, gastrocnemius, diaphragm, tibialis
anterior and extensor digitorum longus). Limb muscles were fixed immediately in 10% BFS
(Confix Australian Biostain AB1020) and remained in solution for at least 72hrs. Tissues
were placed into 70% ethanol, processed in a Shandon automatic tissue processor
overnight, and paraffin embedded for sectioning. Transverse sections (5μm) were cut
through the mid-region of each muscle. Slides were routinely stained with Haematoxylin
and Eosin (H&E) for morphological analysis of the histology. Non-overlapping tiled images
of transverse muscle sections provided a picture of the entire muscle cross section.
Images were acquired using a Leica DM RBE microscope, a personal computer, a Hitachi
HVC2OM digital camera, Image Pro Plus 4.5.1 software and Vexta stage movement
software. Tiled images were taken at 10x magnification.
3) Histological image analysis:
Histological analysis of muscle necrosis was carried out on whole muscle cross sections.
Muscle morphology was drawn manually by the researcher using Image Pro Plus 4.5.1
software.
The area occupied by necrotic myofibres (i.e myofibres with fragmented
sarcoplasm and/or areas of inflammatory cells) was measured as a percentage (area) of
10
the whole muscle section. All section analysis was done ‘blind’. Histological analysis was
completed as per the TREAT-NMD recommended standard protocol “Histological
measurements
of
dystrophic
muscle
-
M.1.2_007”
http://www.treat-
nmd.eu/research/preclinical/SOPs/.
4) Blood collection and Serum Creatine Kinase (CK) assay:
While mice were under terminal anaesthesia, whole blood (approx 0.5ml) was collected
via cardiac puncture using a 27.5 gauge tuberculin syringe (Sigma Z192082), into a 1.5ml
tube. Extensive experimentation revealed that storage of blood samples overnight at 4ºC
to enable clotting leads to a false increase in serum CK levels and therefore blood samples
were immediately spun down in a refrigerated centrifuged for 5 minutes (12000g), serum
was removed and aliquoted. Blood serum CK activity was determined in duplicate using
the CK-NAC kit (Randox Laboratories) and analysed kinetically using a BioTek Powerwave
XS Spectrophotometer using the KC4 (v 3.4) program. A minimum of 10µl serum is
required to complete this assay.
5) Measuring cytokine gene expression by RNA extraction and RT-PCR:
Levels of mRNA for 3 inflammatory cytokines (IL-1β, IL-6 and TNF) was measured in the
quadriceps muscle since this appeared to have the greatest amount of exercise-induced
muscle damage, as indicated by the extent of myofibre necrosis (Figure 4 and 5). RNA was
extracted from one half of a snap frozen quadriceps muscles using Tri-reagent (Sigma
11
T9424) and quantitated using a Nano Drop Spectrophotometer (ND 1000) and ND 1000
software version 3.5.2. The RNA was DNAse treated using Promega RQ1 RNAse free
DNAse (M610A), RQ1 RNAse free 10x buffer (M198A) and RQ1 DNAse stop solution
(M199A). RNA was reverse transcribed into cDNA using Promega M-MLV Reverse
Transcriptase (M3682), random primers (C1181) and 10mM dNTPs (U1515) and the cDNA
was purified using a MoBiol Clean up kit (12500-250). RT-PCRs were run on a Corbett 3000
(Corbett Research) using QIAGEN quantifast SYBR green PCR mix (204054) and QIAGEN
Quantitect Primer Assays for IL-1β (QT01048355), IL-6 (QT00098875) and TNF
(QT00104006), and standardised to a house-keeping gene; ribosomal protein L-19
(QT01779218) as per [31]. mRNA expression levels were calculated and standardised using
Rotor-gene 6.1 and Microsoft Excel software.
6) Quantitation of Oxidative stress:
Oxidative stress in the quadriceps muscle was measured in two different ways:
i) As a ratio of oxidised (di-sulphide) to reduced (sulfhydryl) protein thiols – 2-Tag
technique. Frozen quadriceps muscles were crushed under liquid nitrogen, before protein
extraction with 20% trichloroacetic acid/acetone. Protein was solubilised in 0.5% sodium
dodecyl sulphate 0.5 M tris (SDS buffer), pH 7.3 and thiols were labelled with the
fluorescent dye BODIPY FL-N-(2-aminoethyl) maleimide (FLM, Invitrogen).
Following
removal of the unbound dye using ethanol, protein was resolubilised in SDS buffer, pH 7
and oxidised thiols were reduced with tris(2-carboxyethyl)phosphine (TCEP, Sigma) before
the subsequent unlabelled reduced thiols were labelled with a second fluorescent dye
12
texas red maleimide (Invitrogen). The sample was washed in pure ethanol and
resuspended in SDS buffer. Samples were read using a fluorescent plate reader (Fluostar
Optima) with wavelengths set at excitation 485, emission 520 for FLM and excitation 595,
emission 610 for texas red. A standard curve for each dye was created using ovalubumin
and all results were expressed per mg of protein, quantified using Detergent Compatible
protein assay (BioRad), as per (Armstrong et al 2010, submitted [32].
ii) Malondialdehyde (MDA), a product formed via the decomposition of lipid peroxidation
products [33] was quantitated using High Performance Liquid Chromatography (HPLC).
Quadriceps muscles were ground under liquid nitrogen, homogenized in 10 x 5%
perchloric acid and 150µl of supernatant mixed with 150µl of 40mM Thiobarbituric acid.
Samples were then incubated at 50ºC for 90 minutes and cooled on ice for 15 minutes.
Butanol (250µl) was added, before vortexing and centrifuging for 5 minutes. 20µl of the
upper butanol layer was injected into a C18 HPLC column (5µl, 4.6 x 150 mm, Dionex) with
an isocratic mobile phase of 60:40 50mM KH2PO4: methanol. Samples were run at a flow
rate of 800µl/min for 7 minutes; the retention time was approximately 4.5 minutes.
Fluorescent detection was achieved using the bandpass filters of 515 for excitation and
553 for emission. Tetraethoxypropane (Sigma) was used as a standard for absolute
calculation. Approximately 30mg of tissue was required for this assay.
13
7) Statistics:
Statistical analysis was performed using Microsoft excel and SPSS 16.0. Data were checked
for equal distribution and normality using Q-Q plots. Multiple variables were analysed by
ANOVAs (one, two or three-way to account for exercise, sampling time and strain). All
data are expressed as mean +/- S.E.M unless otherwise stated.
Results:
1) Ability of mdx mice to run on the treadmill
Protocol A: Preliminary studies revealed that approximately 45% of 12 week old untrained
mdx mice could not complete a full 30 minutes of treadmill exercise at 12m/min (despite
being rested 5 times during the 30 minute exercise session), some exhibited severe fatigue
after 10 minutes exercise. This inability to exercise on a treadmill is similar to previous
reports [23, 29, 34]. For this reason the additional ‘warm-up’ period (8 minutes at
8m/min) was included in the treadmill protocol to help the mdx mice to complete the
treadmill exercise session, as per [23]. It is extremely important, especially when only
conducting one single exercise session, to minimise biological variation in the
experimental exercise protocol. Adding a ‘warm-up’ period significantly increased the
ability of mdx mice (92%) to complete the treadmill exercise session. Out of the 24 mdx
mice exercised in protocol A, 2 mice did not fully complete the 30 minutes treadmill
exercise; these 2 mice each completed approximately 26 minutes exercise but, since
neither produced any outlying results, the data were included in all analyses.
14
Protocol B: The average number of rests required to complete each exercise session
reduced throughout the 4 week exercise period, particularly after the 3rd week, suggesting
that mdx mice can improve their running ability with exercise training (Figure 1).
Untrained 12 week old male mdx mice (Protocol A) required significantly (P<0.01) more
rests to complete a 30 minute exercise session compared to 12 week old male mice that
had been ‘trained’ for 4 weeks (Protocol B) (Figure 1). However, it must be noted that the
running protocol used in this study involved resting all the mdx mice on the treadmill
when only one was experiencing fatigue and this may have had an influence on mdx
running ability overtime. There was no significant difference in the running ability of
unexercised 8 week old (1st session of Protocol B) and unexercised 12 week old (Protocol
A) mdx mice (Figure 1).
2) Forelimb grip strength (Protocol B only):
After 4 weeks of treadmill exercise (Protocol B) the forelimb grip strength of 12 week old
mdx mice was significantly weaker than unexercised mice (Figure 2) as shown by a
significant (P<0.01) decrease in both absolute forelimb strength (0.168 kg +/- 0.007
unexercised vs 0.124 kg +/- 0.005 exercised) and a significant (P<0.01) decrease in
normalised change (normalised strength at 12 wks minus normalised strength at 8 wks) in
forelimb strength (0.002 kg/g +/- 0.0004 unexercised vs 0.00038 kg/g +/- 0.0002 exercised)
(Figure 2). This decrease in forelimb grip strength is similar to previous reports [29, 30]
and falls within the expected general range discussed in the TREAT-NMD recommended
15
standard protocol “Use of grip strength meter to assess limb strength of mdx mice –
M.2.2_001” http://www.treat-nmd.eu/research/preclinical/SOPs/.
3) Histological analysis of muscle necrosis.
Biological variation:
Protocol A: One striking feature of histological analysis was the high variation in the
amount of myofibre necrosis (fragmented sarcoplasm and inflammatory cell infiltration)
from both sedentary and exercised (age, sex and muscle matched) mdx mice. The
variation in myofibre necrosis (% area) in quadriceps muscle from 12 week old sedentary
male mdx mice is demonstrated in Figure 3 and ranges from 1.04 – 23.1% for each
individual quadriceps muscle and 2.97 -17.15% average per experimental group. When
results from 9 separate experiments were pooled together (n=60 quadriceps) the average
amount (% CSA) of myofibre necrosis in the quadriceps muscle of an unexercised 12 week
old male mice is 6.12%. Pooled histological data were also used for myofibre necrosis in
unexercised triceps muscle - 8.5% (n=28) and unexercised gastrocnemius muscle - 6.89%
(n=11).
Exercise induced myofibre necrosis:
Protocol A: High variation in myofibre necrosis was again seen in response to a single 30
minute treadmill exercise session and pooled histological data were also used for the
various muscles sampled 24 hours after exercise from 4 experiments (n= 15-25) and at 48
16
hours after exercise from 6 experiments (n= 13-43) (Figure 4). The quadriceps muscle
showed the highest level of myofibre necrosis (15.06 +/- 6.01%) after a single bout of
treadmill exercise when sampled 24hrs, compared with triceps, gastrocnemius, (Figure 4)
tibialis anterior (TA) and extensor digitorum longus (EDL) (data not shown). Necrosis was
significantly increased in treadmill exercised (compared with unexercised) quadriceps
muscles when sampled at either 24hrs (P<0.01) or 48hrs (P=0.04) post exercise. Both the
TA and the EDL muscle from 12 week old mdx mice had a very low level of background
myofibre necrosis (average <3%) and both appeared unaffected by the single treadmill
exercise session with no consistent or significant increase in myofibre necrosis (data not
shown), in accordance with previous reports [24, 26, 27].
Protocol B: Myofibre necrosis was significantly elevated in the quadriceps (P=0.04), triceps
(P=0.05), diaphragm (P=0.04) and TA (P=0.05) muscles after 4 weeks of treadmill exercise
training when mdx mice were sampled 24hrs after the last exercise session. Necrosis was
also significantly elevated in the diaphragm muscle (P=0.02) when sampled immediately
(0mins) after the last exercise session (Figure 5), suggesting prolonged myofibre necrosis
after the penultimate exercise session or a particular sensitivity to exercise induced
damage in the diaphragm.
Necrosis was most elevated in the quadriceps (2x fold) and diaphragm (3x fold) muscle at
24 hours after the last exercise session. The consistently elevated necrosis in the
quadriceps is similar to that seen after a single exercise session (Protocol A). No significant
17
increase in myofibre necrosis was seen in the exercised gastrocnemius muscle possibly
due to high level of variation in this parameter (Figure 5). Myofibre necrosis (fragmented
sarcoplasm and inflammation) returned to unexercised levels (or below) in all muscles,
within 96hrs after exercise (i.e. when the next exercise session would be due). This
indicates that dystrophic myofibres can regenerate muscle to replace necrotic sarcoplasm
and inflammation in between each treadmill exercise session. This also emphasises the
importance of sampling time when quantitating myofibre necrosis after treadmill exercise.
4) Blood Serum CK levels as a measure of muscle leakiness.
Protocol A: No change in serum CK level was seen after 30 minutes treadmill exercise in
control C57Bl/10 mice (unexercised 183.3U/L +/- 53.8 vs exercised 267.7U/L +/- 98.7).
However, serum CK levels were rapidly elevated in response to treadmill exercise in mdx
mice and were significantly higher (P=0.01) when blood was collected immediately
(0mins) after exercise. The exercise induced increase in serum CK was transient and CK
levels dropped rapidly down to unexercised level within 24hours (Figure 6).
Protocol B: In mdx mice subject to 4 weeks treadmill exercise, blood serum CK levels were
significantly (P<0.01) elevated at 24hrs after the last exercise session and decreased to
unexercised level within 96 hours (Figure 6). This is in marked contrast to the rapid
elevation of CK in mdx mice subjected to a single exercise session (Protocol A).
18
5) Inflammatory cytokine gene expression in the quadriceps muscle.
Protocol A: IL-6 mRNA levels were significantly increased in the quadriceps muscle from
C57Bl/10 mice after a single 30 minute treadmill session (Figure 7ii); although there was
no significant change in IL-1β or TNF mRNA. Expression of all 3 inflammatory cytokines
was significantly higher in unexercised mdx mice compared to unexercised C57Bl/10 mice
(Figure 7i-iii). In mdx mice, mRNA levels for both IL-1β (2hrs post exercise P=0.03) and IL-6
(0mins post exercise, P=0.05 and 2hrs post exercise P=0.05) were significantly elevated
after a single 30 minute exercise session compared to unexercised mice (Figure 7i & 7ii).
This rapid increase in gene expression returned to the unexercised level for both genes
within 24 hours. In contrast, levels of mRNA for TNF were significantly reduced after
exercise (0mins post exercise P<0.01, 2hrs post exercise P=0.02 and 24hrs post exercise
P<0.01) compared to unexercised mdx mice (Fig. 7iii).
Protocol B: In mdx mice there was no change in mRNA for IL-1β or IL-6 immediately after
4 weeks of treadmill exercise, although mRNA for both IL-1β and IL-6 was significantly
(P=0.05) decreased for muscles sampled 96hrs post exercise, compared to unexercised
mdx mice (Figure 8i and 8ii). TNF mRNA was significantly decreased in muscles sampled
immediately (0mins) after exercise compared to unexercised mdx mice (Figure 8iii), but
returned to unexercised level within 24 hours.
19
6) Oxidative stress measurement in the quadriceps muscle:
i) Ratio of oxidised (di-sulphide) to reduced (sulfhydryl) protein thiols.
Protocol A: Oxidative stress in the quadriceps muscle, specifically the ratio of oxidised to
reduced protein thiols as measured by the novel 2-tag technique, was significantly
(P=0.01) higher in unexercised mdx compared to unexercised C57Bl/10 12 week old male
mice (Figure 9i). Protein thiol oxidation was also significantly elevated in mdx mice 0mins
(P=0.015) and 2 hours (P=0.04) after a single treadmill session compared to unexercised
mdx mice (Figure 9i). Protein thiol oxidation returned to unexercised level within 24hours.
Protocol B: Similarly, protein thiol oxidation was significantly increased (P=0.02) in mdx
mice after 4 weeks of treadmill exercise when sampled immediately (0mins) after exercise
(Figure 9ii) and returned to unexercised level within 24hours.
ii) Quantitation of Malondialdehyde (MDA).
Protocol A: Oxidative stress in the quadriceps muscle with respect to irreversible lipid
peroxidation, measured as concentration of MDA, showed no significant difference
between C57Bl/10 and mdx mice, and was unaffected by exercise even in mdx mice
(Figure 10). MDA levels were not measured for Protocol B.
Discussion
Treadmill exercise is widely used in pre-clinical experiments to increase the extent of
dystropathology in mdx mice, yet the cellular consequences of a single 30 minute
treadmill exercise session (Protocol A) have not been described previously. The present
20
study analysed the time-course of molecular and cellular changes after a single
standardised 30 minute treadmill exercise session (Protocol A). These data were
compared to data from age matched mdx mice subjected to 4 weeks of treadmill exercise
(Protocol B) a protocol currently widely used for pre-clinical drug screening in mdx mice
[11, 20, 22, 25].
Running ability of mdx mice:
Adding a short warm-up period for 8 minutes at a slower speed (8m/min) produced much
more consistent running by the mdx mice in both treadmill exercise protocols. With only a
single exercise session it is important that all mice complete the exercise protocol to
reduce variation. It is not recommended to remove the ‘non-running’ mice from the
sample group as these may represent mice with the most severe dystropathology; in
comparison to a full range of dystropathology being represented in unexercised control
mice. Preliminary experiments showed that adding a warm-up period significantly
increased the ability of mdx mice (from 45% up to 92%) to complete the 30 minute
treadmill exercise session.
The intermittent running pattern of mdx mice and reduced capacity for exercise
(compared to C57Bl/10 mice) on a voluntary exercise wheel is well documented [11, 35,
36]. With such voluntary running mdx mice can stop and start as they wish, yet they still
manage to run a considerable total amount e.g. up to 14km over 48hrs and up to 435km
over 8 weeks for untreated adult mdx mice [26-28]. In contrast, treadmill exercise requires
21
continuous running (for at least 30 minutes) and mdx mice can struggle with this type of
exercise (indicating fatigue) [23, 29, 30, 34]. Thus, during long-term studies with repeat
(often bi-weekly) treadmill sessions it is important to note the number of times that a
mouse stops running during each treadmill exercise session as this provides some insight
into running ability and thus muscle condition.
In the present study, 4 weeks of treadmill exercise training (Protocol B) significantly
improved the running ability of mdx mice (Figure 1). Despite exhibiting a significant
reduction in both absolute forelimb strength (kg) and change in normalised forelimb
strength (kg/g bw) compared to unexercised mdx mice (Figure 2), exercised mdx mice
show a small increase in normalised forelimb strength after 4 weeks of treadmill exercise
(Protocol B). This increase in normalised forelimb strength along with behavioural
adaptation to exercise training may account for the improvement in treadmill running
ability. However, it must be noted that during the treadmill exercise protocol mice were
run in groups of 3 or 4 and if one mouse fatigued during the 30minute protocol all mice on
the treadmill were rested and this may have impacted the results. Some studies document
an improvement in the voluntary wheel exercise ability (distance run) of mdx mice over
time, especially when exercise is started at a young age [Reviewed in [37]]. However, an
improvement in running capacity on a treadmill over-time has not been previously
reported for mdx mice.
22
Myofibre necrosis:
Increased myofibre necrosis after both treadmill exercise protocols is transient (Figure 4
and 5) and muscles must be sampled between 24 - 48 hours after exercise to visualise the
increase in this histological parameter. While dystrophic skeletal muscles appear fully
capable of regenerating between repeat exercise sessions (Figure 5) it must be noted that
grip strength is significantly reduced in mdx mice subjected to repeated bouts of treadmill
exercise compared with age-matched unexercised mdx mice. Skeletal muscle fibrosis was
not measured in this study although it is likely that, due to exercise induced cycles of
myofibre necrosis associated with inflammation (and regeneration), fibrosis is indeed
progressively increased after 4 weeks of treadmill exercise and may impact negatively on
forelimb grip strength.
Similar to histological results seen after voluntary wheel exercise, treadmill exercise
induces a large amount of damage in the quadriceps muscle [24, 26, 27]. Large variation in
the extent of myofibre necrosis is seen for most muscles; with the quadriceps muscle
showing the highest increase in necrosis after a single 30 minute exercise session
(Protocol A - Figure 4) and the quadriceps, triceps and diaphragm muscles all showing
increased necrosis after 4 weeks of treadmill exercise (Protocol B - Figure 5).
4 weeks of treadmill exercise (Protocol B) induced a more consistent increase in myofibre
necrosis in many muscles (excluding the gastrocnemius) compared to a single 30 minute
exercise session (Protocol A). These data emphasise that large groups of mdx mice (at
23
least 8 mice) are required for histological analysis, due to the notorious variation in mdx
mice [Reviewed in detail [11, 17]] and that the choice of muscle is critical to assess the
impact of exercise induced damage.
Blood serum creatine kinase level:
Serum CK activity is widely used as an indirect measure of muscle damage (sarcolemma
leakiness) and levels are consistently increased in mdx mice after exercise [22, 26, 28, 29,
38]. There is no absolute correlation between the extent of dystropathology in an
individual mdx mouse and CK activity [28], with many factors including stress and muscle
mass influencing CK activity [Reviewed in [11]]. CK is an enzyme and has a short circulating
half life of approximately 12hours [39], thus there is considerable interest in
understanding the kinetics of CK release from dystrophic muscle after treadmill exercise.
Serum CK was strikingly increased immediately (0mins) after a single 30 minute exercise
session (Protocol A) and returned to baseline within 24 hours post exercise (Figure 6). This
elevated level is similar to the transient elevation in mdx serum CK reported at 1 hour
after eccentric exercise (16º downhill, 10m/min for 5 min) [19] and to the increased (x10
fold) serum CK after 8 weeks of voluntary wheel exercise [26]. This immediate increase in
serum CK after exercise indicates that Protocol A was sufficiently strenuous to render
myofibres ‘leaky’ and allow the release of CK into circulation; however it was not
damaging enough to induce widespread myofibre necrosis (Figure 4) in many muscles with
the exception of the quadriceps. Serum CK activity is a measure of muscle damage
24
through-out the whole body, unlike histological analysis of myofibre necrosis which
exclusively examines sections of an individual muscle. In addition to the results presented,
some mice received Evans Blue Dye injections 24hours prior to the single 30 minutes
exercise session to enable histological quantitation of ‘leaky’ myofibres; however this did
not produce any consistent results (data not shown).
In contrast to a single exercise session (Protocol A), CK levels in mdx mice subjected to 4
weeks treadmill training (Protocol B) were significantly increased at 24hrs after exercise,
but not immediately after exercise (Figure 6). This suggests either a delayed or sustained
release of CK from leaky myofibres to account for the prolonged elevation of blood CK
activity. Many factors including training and type of exercise can affect the level and
duration of CK increase; for example in humans a single session of high intensity
resistance exercise immediately increases blood serum CK which peaks at 24 hours and
begins to decline within 48 hours [40]. Our result emphasises the importance of
documenting the timing of such events after different exercise regimes in order to
determine the optimal time for sampling (sacrifice and tissue collection) after exercise
when measuring specific parameters.
Gene expression of inflammatory cytokines:
IL-1β, IL-6 and TNF are 3 major pro-inflammatory cytokines; however it is also recognised
that IL-6 is a myokine, with many important anti-inflammatory and metabolic effects,
produced by and released from contracting myofibres in vivo [Reviewed in [41, 42]].
25
Accordingly, a significant increase in IL-6 mRNA was seen in the quadriceps muscle from
both C57Bl/10 and mdx mice after a single 30 minute treadmill session (Figure 7ii). The
increased IL-6 mRNA in C57Bl/10 mice after treadmill exercise does not coincide with any
inflammation (no change in the expression level of IL1β or TNF – Figure 7i & 7iii) or with
muscle damage (no increase in blood serum CK level or muscle necrosis) and further
demonstrates the capacity of normal muscle contraction to increase IL-6 production. The
pronounced increase in IL-6 mRNA in mdx mice after a single 30 minute exercise session
(Protocol A) is presumably caused by both a response to myofibre contraction and
exercise induced muscle damage. There was no significant increase in IL-6 mRNA in mdx
mice after 4 weeks of treadmill exercise (Protocol B); this may reflect an adaptation
(training) in response to treadmill exercise over time.
The level of TNF mRNA was significantly reduced after both treadmill exercise protocols in
mdx mice. While there is strong evidence that TNF plays a major role in the
dystropathology of mdx mice and blockade of TNF can reduce myofibre necrosis [26, 28,
38, 43], increased IL-6 after exercise can inhibit TNF [44-46] and this may explain the
transient reduction in TNF mRNA seen in response to exercise in the present study.
While there were transient changes in TNF, IL1β and IL-6 mRNA expression after exercise,
changes and bioavailability at the protein level were not measured. This is in part because
of issues with sensitivity of quantification [47]. Indeed, attempts were made to measure
TNF protein levels in blood serum using a standard ELISA (Invitrogen, USA), however
26
serum TNF levels for all mice were below the lowest standard (15.6ρg/ml) and therefore
results were uninformative (data not shown). Another important factor to consider is that
high levels of TNF (and other cytokine) protein are already present, yet sequestered,
within resident and invading inflammatory cells (e.g. mast cells, neutrophils and
macrophages) in dystrophic muscle [13, 27, 28, 48-51]. Protein quantification makes no
comment on the bioavailability or the re-distribution of these cytokines: for example
these proteins are rapidly released from activated mast cells and other inflammatory cells
(e.g. neutrophils and macrophages) that accumulate after myofibre damage. Thus it is
likely that, despite a reduction in TNF mRNA after treadmill exercise, localised bioavailable
TNF protein increases rapidly independent of gene transcription [52].
Oxidative stress:
A significant increase in oxidative stress in unexercised mdx compared to unexercised
C57Bl/10 quadriceps muscle was demonstrated by the increased ratio of oxidised to
reduced protein thiols (reversible oxidative modification) as measured by the novel 2-tag
technique (Armstrong et al, 2010 submitted [32]. This elevated oxidative stress in
dystrophic muscle supports earlier reports that measured oxidative stress in dystrophic
muscle (from both DMD patients and mdx mice) [16, 25, 53-56].
Our second measurement of oxidative stress, using a HPLC method to quantitate MDA
and irreversible peroxidation of membrane lipids, showed no significant difference in
oxidative stress between C57Bl/10 and mdx quadriceps muscle and MDA levels were not
27
increased in mdx muscle after 30 minutes treadmill exercise (Protocol A). While previous
studies have reported increased MDA in the hind limbs of young (<20days) mdx mice [56]
and the gastrocnemius muscle of 90 day old mdx mice [53], compared to age-matched
C57Bl/10 mice, there are numerous reasons for this discrepancy in MDA results, including
animal age and specific muscle examined. It is also important to note that many
commonly used methods for measuring MDA have specificity issues and only HPLC based
methods are recommended [Reviewed in [57]].
Despite no evidence of increased lipid peroxidation in adult mdx quadriceps muscle,
reversible changes in oxidative state were evident by the ratio of oxidised to reduced
protein thiols in mdx muscle. Importantly this 2-tag method identified rapid and
significant increases in protein thiol oxidation immediately (0 mins) and 2 hours after a
single 30 minute treadmill exercise session, emphasising the speed of such changes in vivo
and the sensitivity of this specific assay for oxidative stress.
There is an increasing need for standardised experiments involving mdx mice in order to
readily compare data between laboratory groups globally; this is the subject of recent
reviews [11, 17] that aim to establish a set of recommendations for pre-clinical mdx drug
trials. The present study provides strong support for a single 30 minute exercise session in
adult mdx mice as an appropriate fast protocol to conduct preliminary ‘proof-of-concept’
testing of potential therapeutic drugs to reduce the severity of myofibre necrosis
associated with muscular dystrophy. We have successfully conducted in vivo studies in
28
adult mdx mice examining the potential benefits of on N-acetylcysteine (NAC) using the 30
minute treadmill exercise protocol established in this manuscript (Terrill et al, submitted).
A single 30 minute exercise session (Protocol A) results in a similar level of muscle damage
(muscle necrosis, serum CK, oxidative stress) as 4 weeks of treadmill exercise (Protocol B)
and thus the single exercise session appears suitable as a high through-put screening test.
However a short term protocol does not allow for monitoring of running pattern or
changes in normalised grip strength over time. It is noted that potential therapeutic drugs
for DMD (identified in pre-clinical proof-of-concept short studies) should be tested
chronically to examine efficiency, toxicity and also possible negative side effects e.g on
heart function.
We conclude that a single 30minute treadmill exercise session is a suitable screening
protocol for assessing therapeutic interventions in adult mdx mice (proof-of-concept
studies), that serum CK level and myofibre necrosis and oxidative stress in the quadriceps
muscle are key endpoints which should be monitored when assessing the efficacy of drug
treatments in combination with treadmill exercise (summarised in Table 2), and
emphasise the importance of specific muscle and sampling time (sacrifice and tissue
collection). It is hoped that this simplified single exercise protocol will help accelerate preclinical drug trials in mdx mice and that further insight into the very early events that lead
to myofibre necrosis will identify more precise and better targets for drug interventions to
reduce the severity of the dystropathology.
29
Acknowledgements
The authors thank Greg Cozens and Griffin Grounds for excellent technical assistance.
Research funding from the Australian National Health and Medical Research Council (MG,
TS and PA) and Australian Postgraduate Award Scholarships (HR-C and JT) are gratefully
acknowledged.
30
Table 1. Summary of the animal groups and numbers used for both Protocol A (1x single 30minute
exercise session) and Protocol B (4 weeks of treadmill exercise).
Table 2) Summary of reliable indicators of muscle damage after treadmill exercise in dystrophic
mdx mice. * indicates an extremely high level of high biological variation (large standard error)
thus making the parameter unreliable as an indicator of muscle damage.
Figure 1. Running pattern of mdx mice over 4 weeks of treadmill exercise (8 sessions) compared
to mice subjected to a single exercise session. The average number of rests required to complete
each 30 minute exercise sessions is shown. Mice were 8 weeks old at the start of the Protocol B
and all mice were sampled when 12 weeks old (Protocol A & B). The number of rests is
significantly reduced (*) in 12 week old mdx mice after 4 weeks of treadmill exercise (Protocol B)
in comparison to both untrained 12 week old (Protocol A) and untrained 8 week old mdx mice.
P<0.05, N=24 mice in each session. Bars represent standard error.
Figure 2. Forelimb grip strength (kg) and normalised strength change (kg/g bw); a comparison of
unexercised mdx mice with mice subjected to 4 weeks of treadmill exercise (Protocol B).
Absolute forelimb strength is significantly decreased (*) in 12 week old treadmill exercised mice
and normalised change in forelimb strength is also decreased after 4 weeks of treadmill exercise.
Normalised change in forelimb strength was calculated by normalising absolute force to
bodyweight (kg force / g body weight) and by subtracting normalised forelimb strength of 12 week
old mice from normalised forelimb strength of 8 week old mice. * denotes significant difference
(P<0.05), in exercised compared to unexercised mdx mice. N= 32, 8, 24, 8, 24 mice respectively.
Normalised change in forelimb strength is increased by a scale factor of 100 for graphical purposes
only.
Figure 3. Biological variation in myofiber necrosis in the quadriceps muscle of 12 week old
unexercised male mdx mice. Each bar represents the unexercised control group from a separate
experiment conducted in our laboratory over a period of 12 months. When pooled together (n=60
muscles) the average amount of myofibre necrosis in the quadriceps muscle of unexercised 12
week old male mice is 6.12%. Bars represent standard deviation (to highlight range) and n = 4-8
muscles per group. A,B denotes significant differences, groups with different letters are
significantly different from each other (P<0.05).
Figure 4. Myofibre necrosis (% CSA) in the quadriceps, triceps and gastrocnemius muscles of 12
week old male mdx mice: a comparison of unexercised mice with mice subjected to a single 30
minute exercise session (Protocol A). Myofibre necrosis in the quadriceps muscle is significantly
increased in treadmill exercised mice when sampled both 24hrs (n=25) and 48hrs (n=43) after
exercise in comparison to unexercised mice (n=60). Bars represent standard error. N= 28, 19, 12
respectively for triceps and 11, 15, 20 respectively for gastrocnemius muscle. * denotes significant
difference (P<0.05), in exercised quadriceps compared to unexercised quadriceps.
Figure 5. Myofibre necrosis (% CSA) in the quadriceps, triceps, gastrocnemius, diaphragm and
tibialis anterior muscles of 12 week old male mdx mice: a comparison of unexercised mdx mice
with mdx mice sampled after 4 weeks of treadmill exercise (Protocol B). * denotes significant
increase in necrosis in exercised muscle in comparison to unexercised muscle (same muscle only).
# denotes significant decrease in necrosis in exercised muscle in comparison to unexercised
31
muscle (same muscle only). For unexercised mice n= 60, 28, 11, 8, 8 muscles respectively. N=8 for
all other groups. Bars represent standard error and significant differences were determined by P
<0.05.
Figure 6. Blood serum CK in 12 week old male mdx mice, comparing unexercised mice with mice
subjected to a single 30 minute treadmill session (Protocol A) and 4 weeks of treadmill exercise
(Protocol B). Serum CK is significantly increased after a single 30 minute exercise session (Protocol
A) when mdx mice are sampled 0mins and 2hrs post exercise in comparison to unexercised mdx
mice. Serum CK is significantly elevated in mdx mice subject to 4 weeks of treadmill exercise
(Protocol B) when sampled 24hrs post exercise in comparison to unexercised mdx mice. N=8 mice
per group and bars represent standard error. A,B,C denotes significant differences, groups with
different letters are significantly different from each other.
Figure 7. Gene expression (mRNA) changes in the quadriceps muscle of non-dystrophic C57Bl/10
mice and dystrophic mdx mice in response to a single 30 minute exercise session (Protocol A) for
(i) IL-1β, (ii) IL-6 and (iii) TNF. IL-1β mRNA is significantly increased in mdx mice at 2hrs post
exercise compared to unexercised mdx mice. IL-6 is significantly increased in both C57Bl/10 and
mdx mice immediately after treadmill exercise. TNF is significantly decreased after treadmill
exercise at all times in mdx mice. Bars represent standard error. N= 7-8 mice per group. A,B,C
denotes significant differences, groups with different letters are significantly different from each
other (P<0.05).
Figure 8. Gene expression (mRNA) changes in the quadriceps muscle of mdx mice in response to
4 weeks treadmill exercise (Protocol B) for (i) IL-1β, (ii) IL-6 and (iii) TNF. Both IL-1β and IL-6
mRNA are significantly decreased after 4 weeks of treadmill exercise when mdx mice are sampled
96hrs post exercise. TNF mRNA is significantly decreased at 0mins after exercise compared to
unexercised mdx mice. Bars represent standard error. N= 8 mice per group. A,B,C denotes
significant differences, groups with different letters are significantly different from each other
(P<0.05).
Figure 9. Ratio of oxidised to reduced protein thiols in the quadriceps muscle of non-dystrophic
C57Bl/10 and dystrophic mdx mice in response (i) a single 30 minute treadmill exercise session
(Protocol A) and (ii) 4 weeks of treadmill exercise (Protocol B). Oxidative stress is significantly
higher in 12 week old male mdx mice compared to C57Bl/10 mice. It is further elevated in mdx
mice at both 0mins and 2 hours after a single treadmill exercise session (Protocol A) and at 0 mins
after 4 weeks of treadmill exercise (Protocol B). A,B,C denotes significant differences, groups with
different letters are significantly different from each other (P<0.05).
Figure 10. Malondialdehyde (MDA) concentration in the quadriceps muscle of C57Bl/10 and mdx
mice in response to a single 30 minute treadmill exercise session (Protocol A). There is no
difference in the MDA level between non dystrophic C57Bl/10 and mdx muscle from both
unexercised mice and mice sampled immediately after exercise (0 mins). A,B denotes significant
differences, groups with different letters are significantly different from each other (P<0.05).
32
Figure 1.
Figure 2.
33
Figure 3.
Figure 4.
34
Figure 5.
Figure 6.
35
i
ii
iii
Figure 7.
36
i
ii
iii
Figure 8.
37
i
ii
Figure 9.
38
Figure 10.
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Terrill et al 2010.
Title.
N-acetylcysteine treatment of dystrophic mdx mice prevents exercise induced
myofibre necrosis
Authors.
Jessica R. Terrilla,b, Hannah G. Radley-Crabba, Miranda D. Groundsa, Peter G.
Arthurb.
a.
School of Anatomy and Human Biology, the University of Western Australia,
Crawley, Western Australia, 6009.
b.
School of Biomedical, Biomolecular and Chemical Sciences, the University of
Western Australia, Crawley, Western Australia, 6009.
Corresponding author: Jessica Terrill
Anatomy and Human Biology m309
University of Western Australia
35 Stirling Hwy, Hacket Drive entrance 2
Crawley, Western Australia, 6009
+61 (8) 6488 1498
[email protected]
Email addresses
Hannah G. Radley-Crabb: [email protected]
Miranda D. Grounds: [email protected]
Peter G. Arthur: [email protected]
1
Terrill et al 2010.
Abstract.
Duchenne Muscular Dystrophy (DMD) is a lethal muscle wasting disorder caused by
the absence of functional dystrophin protein in skeletal muscles. Myofibre necrosis is
the primary pathological consequence in DMD and oxidative stress is implicated in
dystropathology. Consequently, strategies to minimize oxidative stress have the
potential to be useful treatment options for DMD. In this study we examined the
potential beneficial effects of N-acetylcysteine (NAC) treatment (1% and 4% in
drinking water) on dystrophic mdx mice. NAC prevented treadmill exercise induced
myofibre necrosis and membrane damage. While NAC had no effect on
malondialdehyde level or protein carbonylation (two indicators of irreversible
oxidative damage), treatment with 4% NAC significantly decreased the oxidation of
glutathione and protein thiols in sedentary mdx mice.
Our data provide evidence
that the protective effects of NAC are multi-factorial, as NAC caused a decrease in
membrane permeability and enhanced protein thiol content. NAC is highly attractive
as a clinical treatment since it is readily available, inexpensive and a clinically proven
drug with few adverse side effects.
Key words.
Duchenne Muscular Dystrophy; mdx mouse; N-acetylcysteine; treadmill exercise;
oxidative stress; protein thiol oxidation.
2
Terrill et al 2010.
1. Introduction*
The muscular dystrophies are inherited muscle disorders characterised by
progressive muscle wasting and weakness. The most common is Duchenne
Muscular Dystrophy (DMD) a lethal, X-chromosome linked disease affecting about 1
in 3500-6000 boys worldwide (Reviewed in [1, 2]). DMD and the milder Becker
Muscular Dystrophy are caused by mutations in the dystrophin gene resulting in the
absence or decreased functional dystrophin protein. Skeletal and cardiac myofibres
lacking functional dystrophin have an increased susceptibility to sarcolemma
damage after muscle contraction which ultimately leads to myofibre necrosis and
regeneration [3, 4]. Repeat cycles of widespread myofibre necrosis lead to the
progressive loss of muscle mass and function in DMD, eventuating in premature
death often due to respiratory or cardiac failure (Reviewed in [2, 5]).
There is no definitive treatment for DMD and, despite many negative side-effects,
corticosteroids remain the standard pharmacological treatment to maintain muscle
mass and strength and help prolong life [2]. Numerous approaches to replace the
defective dystrophin gene have been investigated, however they are not yet clinically
*
Abbreviations
APF,
2-[6-(4-aminophenoxy)-3-oxo-3H-xanthen-9-yl]benzoic
acid;
CTAB,
hexadecyltrimethylammonium bromide; GSH, reduced glutathione; GSSG, oxidised
glutathione;
HPLC,
high
performance
malondialdehyde; NAC, N-acetylcysteine;
liquid
chromatography;
MDA,
SDS, sodium dodecyl sulphate; TBA,
thiobarbituric acid; TCA, trichloroacetic acid; TCEP, tris(2-carboxyethyl)phosphine.
3
Terrill et al 2010.
established [6]. Until genetic or molecular therapies are available, pharmacological
interventions may be useful as interim measures. Particularly attractive are
pharmacological agents where the toxicological information is already documented
for humans, as this should accelerate clinical adoption for DMD.
Myofibre necrosis is the primary pathological consequence in DMD, thus
pharmacological interventions to prevent necrosis are appealing. The mechanism by
which the absence of dystrophin leads to myofibre necrosis is not certain; but there
is evidence that a lack of dystrophin causes myofibre membrane fragility, which
leads to susceptibility to contraction induced injury and excess reactive oxygen
species production, possibly mediated by increased calcium and inflammation [7, 8].
Reactive oxygen species (ROS), such as hydroxyl radicals, can cause cellular
damage by directly damaging macromolecules such as proteins, membrane lipids
and DNA [9]. ROS, such as hydrogen peroxide, can also cause the reversible
oxidation of protein thiols which can affect the function of many intracellular proteins
including signal transduction proteins and transcription factors [10, 11].
For
example, ROS can mediate activation of the transcription factor nuclear factor kappa
B (NFĸB) [12] which itself regulates the expression of many genes, including those
involved in the inflammatory and stress response [13]. Whether it is by direct
damage or via reversible modification of protein thiols, ROS have the potential to be
major contributors to the pathology associated with muscular dystrophies.
Consequently, strategies to minimize the generation or actions of ROS have the
potential to be useful treatment options for DMD.
4
Terrill et al 2010.
A pharmacological agent that targets ROS is the orally available N-acetylcysteine
(NAC); NAC has antioxidant properties [14] and it has been used clinically to treat
various conditions, including acetaminophen overdose (Reviewed in [15]). NAC can
exert its antioxidant properties directly, as it contains a thiol which can directly
scavenge some types of ROS [14-16] and NAC is also a precursor of L-cysteine
which is required in the synthesis of the major intracellular antioxidant glutathione
[17, 18].
NAC has shown promise as an antioxidant to decrease contraction induced injury in
the mdx mouse model of DMD. Treatment of EDL muscles isolated from mdx mice
with 20 mM NAC improved force production and significantly decreased the impact of
eccentric contractions on myofibre membrane integrity [19]. Our objective was to
test whether NAC could prevent contraction induced injury in vivo. We establish that
NAC, when taken orally, decreased myofibre membrane damage and necrosis in
exercised mdx mice. Consistent with the actions of NAC as an antioxidant, there is
evidence of decreased oxidative stress in dystrophic muscle of mdx mice treated
with NAC.
2. Materials and Methods.
2.1 Animal procedures:
All experiments were carried out on 6 - 12 week old male dystrophic mdx
(C57Bl/10ScSnmdx/mdx) and non-dystrophic control C57Bl/10ScSn (C57) mice (the
parental strain for mdx). All mice were obtained from the Animal Resource Centre,
Murdoch, Western Australia. They were maintained at the University of Western
Australia on a 12-h light/dark cycle, under standard conditions, with free access to
5
Terrill et al 2010.
food and drinking water. All animal experiments were conducted in strict accordance
with the guidelines of the National Health and Medical Research Council Code of
practice for the care and use of animals for scientific purposes (2004), and the
Animal Welfare act of Western Australia (2002), and were approved by the Animal
Ethics committee at the University of Western Australia. For consistency, all
experiments (exercise and sampling) were started at 8am and completed by 11am
each day.
2.2 NAC treatment:
NAC (Sigma Aldrich) treatment was administered as either a 1% drinking water
solution for 6 weeks as previously described [19], or as a short term high dosage
regime of 4% drinking water solution for one week, where mice consumed about 2g
of NAC/kg per day) [20].
2.3 Exercise protocol:
In order to initiate contraction induced myofibre necrosis and myofibre membrane
damage, mdx mice were exercised for one single 30 minute exercise session on a
horizontal rodent treadmill (Columbus Instruments USA), according to a protocol
established by our laboratory group [21]. In brief, groups of 3 or 4 mdx mice were (1)
settled for 2 minutes on the stationary treadmill belt, (2) acclimatized with gentle
walking for 2 minutes at a speed of 4 meters/minute, followed immediately by (3) a
warm–up of 8 minutes at 8 meters/minute and then (4) the main exercise session for
30 minutes at 12 meters/minute. Mice were sampled either immediately after
exercise (0 minutes) or 24 hours after treadmill exercise.
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Terrill et al 2010.
2.4 Tissue collection and image acquisition:
All mice were sacrificed at 12 weeks of age by cervical dislocation while under
terminal anesthesia (2%v/v Attane isoflurane Bomac Australia). Various muscles
were collected (while mice were under terminal anesthesia) and immediately snap
frozen in liquid nitrogen for biochemical analysis, or prepared for histology. For
histology, quadriceps muscles were cut in half and mounted on cork board using
tragacanth gum (Sigma Aldrich G1128). Muscles were quenched in isopentane
cooled in liquid nitrogen and stored at -80oC until used for sectioning. Transverse
sections (5-8 μm) were cut through the mid-region of each muscle on a Leica
CM3050S cryostat, as per [21]. Slides were routinely stained with Haematoxylin and
Eosin (H&E). For morphological analysis, non-overlapping tiled images of transverse
muscle sections provided a picture of the entire muscle cross section. Images were
acquired using a Leica DM RBE microscope, a personal computer, a Hitachi
HVC2OM digital camera, Image Pro Plus 4.5.1 software and Vexta stage movement
software. Tiled images were taken at 10x magnification.
2.5 Histological image analysis:
Histological analysis of muscle necrosis was carried out on whole muscle cross
sections of the quadriceps muscle. Muscle morphology was drawn manually by the
researcher using Image Pro Plus 4.5.1 software. The area occupied by necrotic
myofibres (i.e myofibres with fragmented sarcoplasm and/or areas of inflammatory
cells) was measured as a percentage (area) of the whole muscle section. All section
analysis was done „blind‟. Histological analysis was completed as per the TREATNMD recommended standard protocol “Histological measurements of dystrophic
7
Terrill et al 2010.
muscle - M.1.2_007” http://www.treat-nmd.eu/research/preclinical/SOPs/. Due to the
level of variation in myofibre necrosis in 12 week old mdx mice, histological data for
untreated mdx mice consists of pooled data from previous experiments (n=60 for
unexercised mice and n= 25 for exercised mice).
2.6 Blood collection and Serum Creatine Kinase (CK) assay:
While mice were under terminal anaesthesia, whole blood (about 0.5 ml) was
collected via cardiac puncture using a 27.5 gauge tuberculin syringe (Sigma Aldrich),
into a 1.5 ml tube. Blood samples were immediately spun down in a refrigerated
centrifuged for 5 minutes (12000g), serum was removed and aliquoted. Blood serum
CK activity was determined in duplicate using the CK-NAC kit (Randox Laboratories)
and analyses kinetically using a BioTek Powerwave XS Spectrophotometer using the
KC4 (v 3.4) program. In order to confirm the sampling method and analytical method,
levels of CK were also assayed in the control (non-dystrophic) C57 mice. These
levels were very low (approximately 400 units/L) and comparable to previous reports
[22].
2.7 Carbonylated protein:
Oxidative damage to proteins in mdx gastrocnemius muscles was determined by
measuring the carbonyl content with 2,4-dinitrophenylhydrazine as previously
described [23, 24]. Frozen muscles were crushed under liquid nitrogen, before
protein was extracted with 20% TCA/acetone. The protein pellets were washed in
acetone and ethanol, precipitated, dried, re-suspended in 10 mM DNPH in 2 M HCl
and incubated for 30 min at room temperature in the dark. Proteins were washed
with ethyl acetate/ethanol (1:1) and dissolved in 6 M guanidine, and absorbance was
8
Terrill et al 2010.
measured at 370 nm. Protein concentration (mg/ml) was determined using the BioRad Bradford protein assay. Carbonyl concentrations are expressed as nmol of
carbonyl per mg protein.
2.8 Malondialdehyde levels:
Malondialdehyde (MDA) was measured by High performance liquid chromatography
(HPLC) with slight modifications to the method of Seljeskog (2006) [25]. Quadriceps
muscle was ground under liquid nitrogen before the addition of 100 µl per mg of 5%
perchloric acid. This was vortexed and incubated for 1 hour at 4°C. After
centrifugation, 150 µl of supernatant was removed and mixed with 150 µl of 40 mM
TBA. Samples and MDA standards were then incubated at 50°C for 60 minutes, 250
µl butanol was added and separation of the upper butanol layer was achieved with a
C18 column (5 µl, 4.6 x 150 mm, Dionex) using a Dionex Ultimate 3000 HPLC
system. MDA concentrations are expressed as nmol of MDA per mg tissue.
2.9 Glutathione assay:
The levels of reduced and oxidized glutathione were assayed using HPLC with
dansyl derivatives as fluorescent markers, based on Jones (1998) with slight
modifications [26]. Protein was extracted from ground quadriceps muscles by the
addition (1 ml solution per 100 mg tissue) of 10% perchloric acid containing 0.2 M
boric acid (Sigma Aldrich) and 20 µM γ-glutamylglutamate (internal standard).
Separation was achieved using a 3-Aminopropyl column (5µm, 100x2mm,
Phenomenex) using a Dionex Ultimate 3000 HPLC system. Concentrations of
reduced and oxidized glutathione were expressed as nmoles per mg tissue.
9
Terrill et al 2010.
2.10 Protein thiols:
Reduced and oxidized protein thiols were measured as described by Lui et al (2010)
with modifications. Frozen quadriceps muscle was crushed under liquid nitrogen,
before protein was extracted with 20% TCA/acetone.
Protein was solubilized in
0.5% sodium dodecyl sulphate, 0.5 M tris at pH 7.3 (SDS buffer) and protein thiols
were labeled with the fluorescent dye BODIPY FL-N-(2-aminoethyl) maleimide (FLM,
Invitrogen). Following removal of the unbound dye using ethanol, protein was resolubilized in SDS buffer, pH 7 and oxidized thiols were reduced with tris(2carboxyethyl)phosphine (TCEP, Sigma Aldrich) before the subsequent unlabeled
reduced thiols were labeled with a second fluorescent dye Texas Red C2-maleimide
(Invitrogen). The sample was washed in ethanol and resuspended in SDS buffer.
Samples were read using a fluorescent plate reader (Fluostar Optima) with
wavelengths set at excitation 485, emission 520 for FLM and excitation 595,
emission 610 for texas red.
A standard curve for each dye was created using
ovalbumin and results were expressed per mg of protein, quantified using Detergent
Compatible protein assay (BioRad).
2.11 Statistics:
Significant differences between groups were determined using one way ANOVA, and
all data are presented as mean ± standard error of the mean. Significance was set at
p < 0.05.
10
Terrill et al 2010.
3. Results.
3.1 Muscle damage (exercised mice):
We tested whether oral dosing with 1% NAC for 6 weeks could protect dystrophic
muscle from exercise induced myofibre membrane damage and necrosis in vivo. In
untreated mdx mice, a single 30 minute treadmill exercise session caused a
significant increase in necrosis in the quadriceps muscle of mdx mice (Fig. 1A). NAC
completely prevented this increase in exercise induced myofibre necrosis (Fig. 1A).
The levels of necrosis were comparable to the levels of necrosis measured in NAC
treated and untreated mice that had not been exercised (Fig. 1A).
We hypothesized that NAC would prevent myofibre necrosis by stabilizing dystrophic
sarcolemma membranes (as measured by serum creatine kinase level).
NAC
treatment did not affect the levels of CK measured in unexercised mice (Fig. 1B).
Following exercise, there was a rapid elevation in serum CK in untreated mdx mice,
indicative of membrane damage (Fig. 1B). However, while there was a trend for
decreased CK level, treatment with 1% NAC did not significantly attenuate the
increase in CK release after exercise (Fig. 1B).
We tested whether an increase in NAC dosage to 4% would be more successful in
preventing the exercise induced increase in plasma CK. After 1 week of treatment
with 4% NAC, the level of CK was not affected in unexercised mdx mice. However,
in mice treated with 4% NAC, there was about a 2.5 fold smaller increase in CK
following exercise compared to untreated exercised mdx mice (Fig. 2B). Similar to
the data for 1% NAC (Fig. 1A), the exercise-induced increase in myofibre necrosis in
the quadriceps muscle was completely prevented by 4% NAC treatment (Fig. 2A).
11
Terrill et al 2010.
3.2 Oxidative damage:
Since NAC has antioxidant properties, we examined if the protective effects of NAC
could be mediated by a decrease in oxidative stress. The protein carbonyl assay is
commonly used as an indicator of irreversible oxidative damage to proteins [27].
Treatment with either 1% or 4% NAC did not change the levels of protein
carbonylation in unexercised mdx gastrocnemius muscles (Fig. 3A). However, it was
unlikely that NAC protected mdx muscle via the prevention of irreversible oxidative
damage to proteins as protein carbonylation was not increased from pre-exercise
levels (4±0.4 nmol/mg protein, n=6) immediately after exercise (6.5±0.5 nmol/mg
protein, n=5) or 24 hours post-exercise (6.5±0.8 nmol/mg protein, n=7). MDA widely
used as a measure of irreversible oxidative damage to lipids [27]. Treatment with
either 1% or 4% NAC did not change the levels of MDA in unexercised mdx
quadriceps (Fig. 3B). However, it was unlikely that NAC protected mdx muscle via
the prevention of irreversible oxidative damage to lipids as MDA was not increased
from pre-exercise levels (0.38±0.06 nmol/mg tissue, n=9) immediately after exercise
(0.37±0.06 nmol/mg tissue, n=8) or 24 hours post-exercise (0.28±0.04 nmol/mg
tissue, n=9).
An increase in protein carbonyls or MDA following exercise would be indicative of
oxidative damage to proteins or membrane lipids
leading to membrane
destabilization or necrosis in dystrophic muscle. However, protein carbonylation was
not increased immediately after exercise (6.5±0.5 nmol/mg protein, n=5) or 24 hours
post-exercise (6.5±0.8 nmol/mg protein, n=7) from pre-exercise levels (4±0.4
nmol/mg protein, n=6).
Furthermore, MDA was not increased immediately after
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Terrill et al 2010.
exercise (0.37±0.06 nmol/mg tissue, n=8) or 24 hours post-exercise (0.28±0.04
nmol/mg tissue, n=8) from pre-exercise levels (0.38±0.06 nmol/mg tissue, n=9).
These data indicate that oxidative damage to proteins or membrane lipids was not a
likely cause of membrane destabilization or necrosis in dystrophic muscle following
exercise.
3.3 Thiol oxidation:
NAC can scavenge reactive oxygen species indirectly by enhancing glutathione
content [17].However, the protective effects of NAC were not likely mediated by an
increase in glutathione content as total glutathione content was not significantly
altered in the quadriceps muscle by 1% or 4% NAC treatment (Fig. 4A).
The
percentage of oxidized glutathione can also be used as an indicator of oxidative
stress [28]. Treating mdx mice with 1% NAC did not significantly decrease the
percentage of glutathione in the oxidized form (Fig. 4B). However, 4% NAC
significantly decreased the percentage of oxidized glutathione (Fig. 4B). These data
are consistent with NAC acting as a thiol antioxidant in dystrophic muscle.
As NAC is a thiol containing antioxidant we assessed whether NAC protected protein
thiol groups from oxidation. Treating mdx mice with 1% NAC did not change the total
protein thiol content of quadriceps mdx muscle, nor the percentage of protein thiols
oxidized (Fig. 4C and 4D). However, increasing NAC to 4% caused a significant
decrease in the extent of protein thiol oxidation (19 % untreated vs. 12% NAC
treated) (Fig. 4D). The decrease in the extent of protein thiol oxidation was not a
consequence of decreased oxidized protein thiol content; instead, it was a
13
Terrill et al 2010.
consequence of over a two fold increase in reduced protein thiol content from 40 to
86 nmol/mg protein.
An increase in oxidized glutathione or protein thiol oxidation following exercise would
implicate thiol oxidation as a potential cause of membrane destabilization or necrosis
in dystrophic muscle. For glutathione, the percentage of oxidized glutathione was not
increased immediately after exercise (13±3%, n=7) or 24 hours post-exercise
(10±2.8%, n=9) from pre-exercise levels (11.4±1.4%, n=9). However, for protein
thiols, there was a 30% increase in protein thiol oxidation immediately after exercise
(Fig 5B). As protein thiol state was affected by exercise, we examined the effect of
4% NAC treatment on protein thiol state following exercise in mdx quadriceps
muscle. Consistent with the observation in unexercised mice, total protein thiols were
increased in exercised mdx mice treated with 4% NAC. (Fig. 5A) In addition,
significantly more protein thiols were oxidized in muscles from 4% NAC treated mice
(20 nmol/mg protein) than untreated mice (13 nmol/mg protein) (Fig. 5B). Treatment
with 1% NAC had no effect on protein thiol groups after treadmill exercise (data not
shown).
Together these data indicate that the protective properties were a
consequence of NAC acting as a thiol antioxidant.
4. Discussion.
The effectiveness of NAC in preventing myofibre necrosis in dystrophic muscle has
not been previously reported in vivo. In this study we show that NAC prevented
exercise induced myofibre necrosis, using two different dosing regimens of 1% NAC
for 6 weeks and 4% NAC for 1 week. Prevention of exercised induced myofibre
necrosis is an important outcome, because repeated bouts of myofibre necrosis
14
Terrill et al 2010.
eventually lead to the replacement of myofibres with fatty and fibrous connective
tissue and thus a loss of function of dystrophic muscle [5, 29].
One possible mechanism by which NAC protects dystrophic muscle from exercise
induced myofibre necrosis is by stabilizing fragile dystrophic sarcolemma
membranes. Previous in vitro work supports this hypothesis, as uptake of Evans
Blue Dye into isolated myofibres following stretched induced contractions decreased
with exposure to NAC [19]. For the current in vivo experiments we used CK release
to assess myofibre membrane damage; mdx mice have elevated CK levels [30]
which are greatly exacerbated after treadmill exercise [22]. While there was a trend
for attenuation in CK level following exercise with 1% NAC treatment, this was not
statistically significant. This may be due to the high variation in CK levels following
exercise in mdx mice [31], resulting in poor sensitivity to the protective effects of
NAC (Type II error). However, when the dose of NAC was increased to 4%, there
was a significant attenuation of increased blood CK after treadmill exercise. These
findings are important as they indicate that NAC can rapidly (within one week)
protect dystrophic muscle from damage in vivo.
These data also imply that doses of NAC higher than 1% are more protective. This
will need to be further tested with consistent treatment protocols to determine the
optimal dose for in vivo treatment. Overall, our data and evidence from a previous in
vitro study [19] are consistent with the concept that NAC stabilizes fragile dystrophic
membranes, although how this occurs is not clear.
15
Terrill et al 2010.
The beneficial actions of NAC may primarily reflect its antioxidant properties since
oxidative stress has been identified as a possible cause of myofibre necrosis and
muscle wasting of dystrophic muscle [11, 32]. ROS can cause irreversible damage to
macromolecules such as membrane proteins and lipids, with extensive damage
potentially leading to membrane permeability. Such irreversible oxidation of proteins
can be assessed by measuring the protein carbonyl content of muscles. Increased
protein carbonyl content has been reported for both DMD biopsies and mdx muscles
[33, 34]. Oxidative damage to proteins can be caused by hydroxyl radicals, which
can be prevented by NAC [15, 16]. However, in the current study NAC treatment did
not decrease protein carbonyl content in mdx muscle. Oxidative damage to lipids
(lipid peroxidation) has also been reported in young mdx muscle [34, 35] and
irreversible damage to membrane lipids can be assessed by measuring
malondialdehyde (MDA), generated as a consequence of lipid peroxidation.
However, we did not observe a decrease in MDA level in mdx mice after either NAC
treatment. Overall, our data indicate that the protective effects of NAC in dystrophic
muscle do not seem to be a consequence of a decrease in irreversible oxidative
damage to protein and lipids.
Another mechanism proposed for the antioxidant properties of NAC is up-regulation
of the cellular antioxidant glutathione [17], since NAC is a precursor, via cysteine, for
the synthesis of glutathione [14, 36]. Glutathione, a non-protein containing thiol, is a
naturally occurring antioxidant present at high concentrations (mM) in tissues that
can react directly with ROS.
Of particular significance, glutathione is also an
essential co-factor involved in the removal of hydrogen peroxide by glutathione
peroxidase [9, 37, 38]. In this study, both 1% and 4% NAC treatment produced no
16
Terrill et al 2010.
increase in total glutathione in dystrophic myofibres. These observations imply that
decreased myofibre necrosis caused by NAC was not a consequence of increased
glutathione synthesis.
An effect of NAC treatment which has not been previously described was the
increase in reduced protein thiols in mdx muscle.
We propose the protective
properties of NAC are a consequence of this increase in reduced protein thiol
content. As reduced protein thiols can become preferentially oxidized as a result of
oxidative stress, protein thiols can be considered to be acting as antioxidants,
somewhat analogous to glutathione [36].
In NAC treated mice, the increase in
reduced protein thiol content in tissue would have enhanced the antioxidant capacity
of muscle protecting the muscle from increased oxidative stress. Consistent with
concept, there was considerably more oxidized protein thiols following exercise in
NAC treated mice than in untreated mice, suggesting the oxidation of the increased
protein thiol pool is preventing oxidation of other constituents of the dystrophic
muscle that leads to necrosis of the myofibres [36].
In summary, our novel in vivo data show that NAC protects dystrophic muscle from
exercise induced myofibre necrosis, the primary cause of severe muscle pathology
seen in DMD.
The exact mechanism responsible for these benefits of NAC is
unclear at this stage, mainly because there is still uncertainty regarding the precise
pathways leading to necrosis of dystrophic myofibres. Membrane permeability and
oxidative stress have been implicated in causing necrosis. Our in vivo data provide
evidence that the protective effect of NAC could be multifactorial, as NAC caused a
decrease in membrane damage and permeability and enhanced protein thiol
17
Terrill et al 2010.
content. NAC is highly attractive as a readily available, inexpensive and clinically
proven drug with few adverse side effects. Our data strongly support further
preclinical investigation into the potential dose of NAC to mitigate aspects of
dystrophic pathology.
Acknowledgements.
The authors thank Alex Armstrong and Dr Ahmed Elshafey for their expert
assistance in method development. Funding from the Australian National Health and
Medical Research Council (NHMRC) and Australian Postgraduate Research
Scholarships for JT and HR-C are also gratefully acknowledged.
Author Disclosure Statement.
The authors declare they have no conflicts of interest.
18
Terrill et al 2010.
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List of Figure Captions:
Figure 1. Effect of exercise on mdx mice treated with 1% NAC. (A) Myofibre
necrosis (% CSA) was assessed in the quadriceps muscles of untreated and 1%
NAC treated mdx mice prior to exercise (Unex mdx) and 24 hours after exercise
(24hrs after ex.). n= 60, 8, 25 and 6 respectively. (B) CK was measured in blood
serum from untreated and 1% NAC treated mdx mice prior to exercise (Unex mdx)
and
0
minutes after
exercise
(0mins after
ex.).
n= 13,
8,
8
and
8
respectively.*Significantly different between untreated and 1% NAC treated.
#
Significantly different between unexercised and exercised mice.
Figure 2. Effect of exercise on mdx mice treated with 4% NAC. (A) Myofibre
necrosis (% CSA) was assessed in the quadriceps muscles of untreated and 4%
NAC treated mdx mice prior to exercise (Unex mdx) and 24 hours after exercise
(24hrs after ex.). n= 60, 8, 25 and 6 respectively. (B) CK was measured in blood
serum from untreated and 4% NAC treated mdx mice prior to exercise (Unex mdx)
and 0 minutes after exercise (0mins after ex.). n= 13, 8, 8 and 8 respectively.
*Significantly different between untreated and 1% NAC treated.
#
Significantly
different between unexercised and exercised mice.
Figure 3. Oxidative damage in unexercised mdx mice treated with 1% and 4%
NAC. (A) Carbonyl content in gastrocnemius muscle of untreated and NAC treated
mdx mice. n= 6, 7 and 7 respectively. (B) Malondiadehyde content in quadriceps
muscle of untreated and NAC treated mdx mice. n= 9, 8 and 6 respectively.
24
Terrill et al 2010.
Figure 4. Thiols in unexercised mdx mice treated with 1% and 4% NAC. (A)
Total glutathione content and (B) percentage of oxidised glutathione in quadriceps
muscle of untreated and NAC treated mdx. n= 9, 7 and 7 respectively. (C) Total
protein thiol content and (D) percentage of oxidised protein thiol content quadriceps
muscle of untreated and NAC treated mdx mice. n= 9, 8 and 4 respectively.
*Significantly different from untreated.
Figure 5. Effect of exercise on protein thiols for mice treated with 4% NAC. (A)
Total protein thiol content and (B) percentage of oxidised protein thiol content of
untreated and NAC treated mdx quadriceps muscle prior to exercise (Unex mdx) and
0 minutes after exercise (0mins after ex.). *Significantly different between untreated
and NAC treated. #Significantly different between unexercised and exercised mice.
n= 9, 6, 4 and 7 respectively.
25
Terrill et al 2010.
Fig. 1
26
Terrill et al 2010.
Fig. 2
27
Terrill et al 2010.
Fig. 3
28
Terrill et al 2010.
Fig. 4
29
Terrill et al 2010.
Fig. 5
30
Radley-Crabb and Grounds, 2010. 1
The different impacts of a high fat diet on dystrophic mdx and control C57Bl/10 mice.
Hannah G Radley-Crabb* and Miranda D Grounds.
School of Anatomy and Human Biology, the University of Western Australia, Crawley,
Australia.
*Corresponding author: Hannah Radley-Crabb, (08) 6488 1498,
[email protected]
Key words:
Duchenne muscular dystrophy, mdx mouse, skeletal muscle, high fat diet, high protein
diet, body composition, dystropathology.
Radley-Crabb and Grounds, 2010. 2
Abstract:
The absence of functional dystrophin protein in patients with Duchenne muscular
dystrophy (DMD) and dystrophic mdx mice leads to fragile myofibre membranes and
cycles of myofibre necrosis and regeneration. Studies in both DMD patients and mdx
mice indicate that dystrophic muscles have altered metabolism and impaired energy
status and has led to the proposal that nutritional supplementation may reduce the
severity of dystropathology. This research compares the in vivo responses of
dystrophic mdx and normal control C57Bl/10 mice to a high protein (50%) or a high fat
(16%) diet. Consumption of a high protein diet had minimal effects on the body
composition or muscle morphology in both strains of mice. In contrast, striking
differences between the strains were seen in response to the high fat diet; which also
varied between mdx mice aged <24 weeks, and mdx mice aged 24 - 40 weeks.
C57Bl/10 mice demonstrated many negative side effects after consuming a high fat
diet including weight gain, significantly increased body fat and elevated inflammatory
cytokines. In contrast, mdx mice (< 24 weeks) remained lean with minimal fat
deposition and were resistant to changes in body composition after consuming the
high fat diet. These results support the proposal that dystrophic mdx mice have an
altered ‘energy status’ compared to normal C57Bl/10 mice because the mdx mice
appeared more capable of metabolising the high fat diet and avoiding fat deposition.
However, older mdx mice (24 - 40 weeks old), with increased energy intake, exhibited
some mild adverse effects of a high fat diet but to a far lesser extent than age-matched
C57Bl/10 mice. Benefits of the high fat diet on dystrophic muscles of young mice were
demonstrated by the significantly increased running ability (km) of voluntarily
exercised mdx mice and significantly reduced myofibre necrosis in 24 week old
sedentary mdx mice. These novel data clearly identify an ‘altered’ response to a high
fat diet in dystrophic mdx compared to normal C57Bl/10 mice. The high fat diet was
beneficial to young mdx mice and our data indicate that energy deficient mdx mice
may utilise excess dietary fat to improve muscle function and reduce muscle damage.
Radley-Crabb and Grounds, 2010. 3
Introduction:
Duchenne Muscular Dystrophy (DMD) is a lethal X-linked muscle wasting disease
resulting from defects in the myofibre subsarcolemmal protein dystrophin. DMD is
characterised by progressive muscle weakness and wasting with a limited life
expectancy of approximately 20 years in humans (Biggar, 2006; Bushby et al, 2010;
Emery, 2002; Sinnreich, 2010). The lack of functional dystrophin leads to myofibre
membrane fragility, repeated cycles of myofibre necrosis and regeneration, and the
eventual replacement of skeletal muscle by fatty and fibrous connective tissue.
Corticosteroids remain the standard pharmacological treatment to maintain muscle
mass and help prolong life, despite severe adverse side-effects including obesity,
Cushingoid symptoms, immune suppression, hypertension, behavioural changes and
cataracts (Angelini, 2007; Biggar et al, 2006; Bushby et al, 2010; Manzur et al, 2008).
Promising approaches to replace the defective dystrophin gene are not yet established
for clinical use although numerous potential treatment options have been thoroughly
investigated over the last 10 years (Cossu and Sampaolesi, 2007; Guglieri and Bushby,
2010; Manzur and Muntoni, 2009; Nagaraju and Willmann, 2009; Radley et al, 2007;
Smythe et al, 2001; Wells, 2008). The specific mechanism(s) leading to myofibre
necrosis are still unclear, yet there are strong associations with excessive
inflammation, increased intracellular calcium levels, elevated oxidative stress and
metabolic abnormality (Davidson and Truby, 2009; Evans et al, 2009; Even et al, 1994;
Kuznetsov et al, 1998; Radley et al, 2008; Tidball and Wehling-Henricks, 2007;
Whitehead et al, 2008).
Despite there being fundamental differences in growth parameters, body size and
muscle loading, that lead to very different disease severity between dystrophic mdx
mice (C57Bl/10ScSnmdx/mdx) and DMD patients; the mdx mouse is a widely used animal
model for pre-clinical DMD research [Reviewed in (Grounds, 2008; Grounds et al,
2008b; Vainzof et al, 2008; Willmann et al, 2009)]. Studies in both DMD patients and
mdx mice indicate that dystrophin defects may also lead to altered skeletal muscle
metabolism and an impaired energy status. Repeated cycles of myofibre necrosis and
regeneration, increased demand on the sarcoplasmic reticulum to regulate
Radley-Crabb and Grounds, 2010. 4
intracellular calcium, defective mitochondrial function, increases in both protein
synthesis and protein degradation rates, and disruption in nNOS signalling may all
contribute to an altered metabolic state (Dupont-Versteegden et al, 1994; Griggs and
Rennie, 1983; Han et al, 2006; Kuznetsov et al, 1998; Landisch et al, 2008; MacLennan
and Edwards, 1990; Passaquin et al, 2002; Whitehead et al, 2006; Zanardi et al, 2003).
In support of an altered metabolism in dystrophic muscle, 48 hours of fasting
significantly increased myofibre necrosis in muscles from the hind limb and lumbar
region of 6 month old mdx mice (yet no change in muscle morphology of control
C57BL/10 mice), suggesting a strong dependence on an adequate energy intake to
maintain dystrophic muscle structure (Helliwell et al, 1996).
Dietary interventions in the form of various amino acids (with different biochemical
effects) or other nutritional supplements have shown variable beneficial effects in both
mdx mice and DMD patients [Reviewed in (Bogdanovich, 2004; Davidson and Truby,
2009; Leighton, 2003; Pearlman and Fielding, 2006; Radley et al, 2007)]. Recently
completed clinical trials have examined the role of specific amino acid
supplementation (creatine and glutamine) in DMD patients and demonstrated some
beneficial effects (Escolar et al, 2005; Tarnopolsky et al, 2004). In mdx mice, a creatine
enriched diet (10% w/w in chow) fed to new-borns strongly reduced the onset of
muscle necrosis in the fast-twitch EDL muscle and also improved mitochondrial
respiration capacity (Passaquin et al, 2002). In addition, creatine, taurine and
glutamine treatments have shown beneficial effects in treadmill exercised adult mdx
mice (De Luca et al, 2003; Granchelli et al, 2000). A combined nutritional therapy
(creatine monohydrate, conjugated linoleic acid, α-lipoic acid, and β-hydroxy-βmethylbutyrate) administered, in addition to prednisolone, for 8 weeks increased
muscle strength and reduced the extent of dystropathology in 12 week old treadmill
exercised mdx mice (Payne et al, 2006). A combination of taurine (1g/(kg bw.day) orally) and prednisolone (1mg/(kg bw.day) - i.p. injection) treatment of treadmill
exercised mdx mice (4-8 weeks of age) also markedly improved forelimb grip strength,
compared to either taurine or prenisolone treatment alone (Cozzoli et al, 2010). In
addition, a high protein diet (50%) improved muscle morphology in dystrophic laminin
Radley-Crabb and Grounds, 2010. 5
deficient (129ReJ dy/dy) mice and caused a shift to a more ‘normal’ protein
metabolism (Zdanowicz et al, 1995). In many cases, the molecular/metabolic basis for
the variable benefits reported for the different interventions in dystrophic skeletal and
heart muscle remains to be determined. Additional work is required to determine the
optimal dietary intervention to reduce the severity of dystropatholgy for potential
application to DMD.
We were impressed by a preliminary study to test the effects of a high fat diet on the
dystrophic mdx heart (Andrew Hoey 2005, Queensland, data unpublished) that
reported a striking difference in bodyweight of dystrophic mdx and control mice fed a
high fat diet (~15% w/w) for 9 weeks (from 6–15 weeks of age). As expected, C57Bl/10
mice showed significantly greater body weight and % body fat; however, the body
weight of mdx mice was unaffected by the change in diet. The dystropathology of
skeletal muscles was not examined in this study, but these interesting results led us to
propose that a high fat diet may reduce the extent of dystropathology, and may be
metabolically beneficial to mdx mice due to their altered energy status.
The aims of the present study were to compare the effects of a high protein and high
fat diet on the body composition, muscle morphology and gene expression levels in
sedentary adult and voluntarily exercised C57Bl/10 and mdx mice. Body composition
was assessed for both strains by measuring total body weight, gastrocnemius muscle
weight, myofibre cross-sectional area and epididymal fat pad weight. Muscle
morphology and dystropathology (for mdx mice only) was assessed by histological
quantification of myofibre necrosis and adipocyte infiltration into the quadriceps
muscle, blood serum creatine kinase (CK) level and the distance run during voluntarily
exercise. Gene expression levels of 3 major inflammatory cytokines; Tumor necrosis
factor (TNF), Interleukin 1β (IL-1β) and Interleukin 6 (IL-6) were measured because of
their role in exacerbating myofibre necrosis in dystrophic muscle (Evans et al, 2009;
Grounds et al, 2008a; Radley et al, 2008; Tidball and Wehling-Henricks, 2005) and
because long-term consumption of a high fat diet and subsequent increase in visceral
Radley-Crabb and Grounds, 2010. 6
obesity is associated with low-grade chronic inflammation [Reviewed in (Pedersen,
2007; Stienstra et al, 2007; Wisse, 2004)]. Gene expression of the peroxisome
proliferator-activated receptors (PPARs), a group of nuclear receptor proteins
(transcription factors); PPAR alpha, PPAR delta/beta, PPAR gamma and PPAR gamma
coactivator 1 alpha (PGC-1α), were measured due to their pivotal role in metabolism,
inflammation, adiposity, and remodelling of skeletal myofibre type composition
[Reviewed in (Liang and Ward, 2006; Stienstra et al, 2007; Zierath and Hawley, 2004)].
Methods:
Animals. Experiments were carried out on male non-dystrophic (control) C57Bl/10ScSn
(hereafter referred to as C57) and dystrophic mdx (C57Bl/10ScSn mdx/mdx) mice; all mice
were obtained from the Animal Resource Centre, Murdoch, Western Australia. They
were maintained at the University of Western Australia on a 12-h light/dark cycle,
under standard conditions, with free access to food and drinking water. All animal
experiments were conducted in strict accordance with the guidelines of the National
Health and Medical Research Council Code of practice for the care and use of animals
for scientific purposes (2004), and the Animal Welfare act of Western Australia (2002),
and were approved by the Animal Ethics committee at the University of Western
Australia.
Experimental groups and custom diets. This study was conducted over three separate
experiments using the following groups of mice: Group 1) sedentary 8-24 week old C57
and mdx mice, Group 2) sedentary 24-40 week old C57 and mdx mice and Group 3)
voluntarily exercised 8-12 week old mdx mice. All mice were fed a standard (cereal
based) mouse chow (meat free, 5% fat, 19% protein) prior to the study. Sedentary
mice (groups 1 and 2) were fed a customised semi-purified diet (control, high fat or
high protein) for 16 weeks and voluntarily exercised mice were fed a customised semipurified diet for 4 weeks. N=8 mice for all groups. The three customised semi-purified
diets; Control (7% fat, 19% protein - AIN93G), High Fat (16% fat, 19% protein - SF 06040) and High Protein (7% fat, 50% protein – SF 00-252) were manufactured (all in
Radley-Crabb and Grounds, 2010. 7
pellet)
form
by
Specialty
Feeds
Glen
Forest
Western
Australia
www.specialtyfeeds.com.au (Table 1). Pilot trials were conducted prior to
commencement of the study to ensure that all diets were palatable. Throughout the
study food intake and body weight were continuously monitored. Food intake was
measured by removing and weighing all food from both the food compartment on the
cage lid and from inside the cage (e.g separated from the cage bedding). Energy intake
was calculated by adjusting food consumption (g) to metabolisable energy content of
each diet (Table 1): metabolisable energy content was calculated according to
guidelines
of
Food
and
Agriculture
Association
of
the
United
Nations,
http://www.researchgate.net/journal/0254-4725_FAO_food_and_nutrition_paper.
Voluntary exercise (8-12 week old mdx mice only). The low level of muscle damage in
dystrophic adult mdx mice can be elevated by voluntary exercise (Grounds et al,
2008b). Mice were caged individually with free access to a voluntary running wheel for
4 weeks (8-12 weeks of age). Exercise data were collected via a small magnet attached
to the mouse wheel, and a sensor from a bicycle pedometer attached to the back of
the cage. The pedometer records single wheel revolutions, allowing total distance (km)
run by an individual mouse to be determined, as per (Hodgetts et al, 2006; Radley et
al, 2008; Radley and Grounds, 2006). The mice run the most during the night since they
are normally nocturnal (Hara et al, 2002; Radley and Grounds, 2006). Mice were
monitored daily throughout the experiment for food consumption, distance run and
wheel function.
Tissue collection. Mice were sacrificed at either 12, 24 or 40 weeks of age by cervical
dislocation, while under terminal isoflurane (Bomac Australia) anaesthesia (2%v/v).
Total body weight, tibial length, epididymal fat pad weight and gastocnemius muscle
weight were recorded immediately. Blood serum was collected via cardiac puncture,
the gastrocnemius muscle was snap frozen in liquid nitrogen for molecular analysis and
the quadriceps and tibialis anterior (TA) muscles were collected for histology (paraffin
and frozen section preparation, respectively).
Radley-Crabb and Grounds, 2010. 8
Skeletal muscle histology. The quadriceps muscles were collected and immediately
fixed in 4% paraformaldehyde (Sigma P6148) for 48 hours. Muscles were then placed
into 70% ethanol, processed in a Shandon automatic tissue processor overnight, and
finally paraffin embedded for sectioning. Transverse sections (5μm) were cut through
the mid-region of each muscle on a Leica retractable microtome. Slides were stained
with Haematoxylin and Eosin (H&E) for morphological analysis of dystropathology (e.g
myofibre necrosis and adipocyte content) as per the TREAT-NMD recommended
standard protocol “Histological measurements of dystrophic muscle - M.1.2_007”
http://www.treat-nmd.eu/research/preclinical/SOPs/. The frozen TA muscles were
collected, mounted on a small cork block and stabilised (embedded) in tragacantch
gum (Sigma Aldrich G1128), quenched in isopentane cooled in liquid nitrogen and
stored at -80ºC until cut on a CM3050S Leicsa cryostat. TA sections were stained with
Sirius red, which readily distinguishes myofibres (yellow) from collagenous myofibre
membranes (red), to allow for easy and accurate quantification of myofibre crosssectional area (CSA).
Image capture and Histological image analysis. Non-overlapping tiled images of
transverse muscle sections provided a picture of the entire muscle cross section.
Images were acquired using a Leica DM RBE microscope, a personal computer, a
Hitachi HVC2OM digital camera, Image Pro Plus 4.5.1 software and Vexta stage
movement software. Tiled images were taken at 10x magnification. Histological
analysis was carried out on whole cross sections of the quadriceps muscle
(dystropathology and adipocyte content) and the frozen TA muscle (myofibre CSA).
Muscle morphology was drawn interactively by the researcher using Image Pro Plus
4.5.1 software.
Serum Creatine Kinase Assay. While under terminal anaesthesia blood from the mdx
mice was collected via cardiac puncture. Blood was refrigerated overnight, centrifuged
Radley-Crabb and Grounds, 2010. 9
for 3 min (1200 rpm) and serum removed. Blood serum creatine kinase (CK) analysis
was completed at the Murdoch Veterinary Hospital, Murdoch, WA.
Quantitation of Gene expression (relative to L-19). Consumption of a high fat diet
induced many changes in the body composition of both C57 and mdx mice, whereas a
high protein diet did not. Therefore gene expression was quantified only in mice which
consumed the control or high fat diet.
RNA was extracted from snap frozen
gastrocnemius muscles using Tri-reagent (Sigma T9424) and quantified using a Nano
Drop Spectrophotometer (ND 1000) and ND 1000 software version 3.5.2. The RNA was
DNAse treated using Promega RQ1 RNAse free DNAse (M610A), RQ1 RNAse free 10x
buffer (M198A) and RQ1 DNAse stop solution (M199A). RNA was reverse transcribed
into cDNA using Promega M-MLV Reverse Transcriptase (M3682), random primers
(C1181) and 10mM dNTPs (U1515) and the cDNA was purified using a Mo Biol Clean up
kit (12500-250). RT-PCRs were run on a Corbett 3000 (Corbett Research) using QIAGEN
quantifast SYBR green PCR mix (204054) and QIAGEN Quantitect Primer Assays for TNF
(QT00104006), IL-1β (QT01048355), IL-6 (QT00098875), PPAR alpha (QT00137984),
PPAR delta/beta (QT00166292), PPAR gamma (QT00100296), PGC-1α (QT00095578)
and standardised to an appropriate house-keeping gene; ribosomal protein L-19
(QT01779218) as per (Shavlakadze et al, 2010). mRNA expression was calculated and
standardised using Roto-gene 6.1 and Microsoft Excel software.
Statistical analysis. Analysis was completed using Microsoft excel and SPSS 16.0. All
variables were analysed by ANOVAs (to account for diet, strain, age and exercise) and
Least Significant Difference (LSD) post-hoc tests. All data are expressed as mean +/S.E.M throughout the manuscript.
Results:
Food consumption.
There were no significant differences in average daily food consumption (g/ g bw/ day)
across the 3 custom diets in both strains of 8-24 week old mice (Figure 1A). However
Radley-Crabb and Grounds, 2010. 10
there was a significant decrease in food consumption (g/ g bw/ day) with age in both
strains of mice and a significant decrease in food consumption in 24-40wk old C57
mice on the high fat diet compared to both the control and high protein diet (Figure
1A). Absolute energy intake (kj /day) was significantly decreased in both strains of 824 week old mice on the high protein diet (Figure 1B). There was a slight increase in
energy intake (kj /day) in both strains of mice on the high fat diet at 8-24 weeks of age
compared to mice on the control diet; however this difference was not significant. In
the older mice (24-40 wks) on the high fat diet there was a significant increase in
energy intake (kj /day) in both strains compared to mice on both the control and high
protein diet (Figure 1B). There were no differences in standardised energy intake (kj / g
bw /day) between 8-24 week old C57 or mdx mice on the control diet, however both
C57 and mdx mice on the high protein diet consumed significantly less standardised
energy compared to mice on both the control and high fat diet (Figure 1C). There was a
significant decrease in energy intake in both strains of mice with age, and 24-40 week
old mdx mice on the high fat diet consumed significantly (P<0.02) more energy (kj / g
bw /day) than all other groups of mice (Figure 1C).
Body Composition.
The linear growth of both adult C57 and mdx mice, as reflected by tibial bone length,
was not affected by diet (data not shown).
Group 1) 8 - 24 week old sedentary mice subjected to 3 diets. 8 week old dystrophic
mdx mice (weight at beginning of the study) fed a standard (cereal based) mouse chow
were significantly heavier (P=0.02) than age matched C57 mice (C57 21.6g +/- 0.23 vs.
mdx 23.4g +/- 0.72). This significant difference in body weight was maintained
throughout the study until 24 weeks of age (C57 30.6g +/- 0.74 vs. mdx 35.5g +/- 0.98)
(Figure 2A). The high protein diet had no significant effect on the bodyweight of either
strain of mice (Figure 2A). After consuming the high fat (HF) diet for 16 weeks there
was a significant (P=0.04) increase in the bodyweight of 24 week old C57 mice
compared to C57 mice on the control (C) diet (HF 34.54g +/- 1.2 vs. C 30.63g +/- 0.73). In
contrast, consumption of the high fat diet caused no significant change in the
bodyweight of mdx mice (HF 34.97g +/- 0.53 vs. C 35.54g +/- 0.98) (Figure 2A). The
largest gain in body weight (body weight at 24wks minus body weight at 8wks) was
Radley-Crabb and Grounds, 2010. 11
seen in C57 mice after consuming the high fat diet. The average group percentage
changes in body weight for C57 mice were 130% after control diet, 140% after high
protein diet and 159% after high fat diet. The average group percentage changes in
body weight for mdx mice were 130% after control diet, 132% after high protein diet
and 137% after high fat diet.
Obesity is the direct result of an imbalance between energy intake and energy
expenditure, with excess energy being primarily stored in adipose tissue in the form of
triglycerides [Reviewed in (Stienstra et al, 2007)]. Epididymal fat pad weight increases
in male C57Bl/6J mice after consumption of a high fat diet (Turpin et al, 2009) and was
measured in this study as an indicator of obesity. After consuming the control diet, the
standardised epididymal fat pad weight (g fat /g bw) was significantly (P<0.001)
heavier in C57 compared to mdx mice at 24 weeks of age (C57 0.019g/g bw +/- 0.009
vs. mdx 0.0066g/g bw +/- 0.002) (Figure 2B). The high protein diet had no significant
impact on epididymal fat pat weight in either strain of mice, in contrast with the
striking effects of the high fat diet. After consuming the high fat diet (from 8-24
weeks), the standardised epididymal fat pad was significantly (P=0.015) increased in
C57 mice (HF 0.028g/g bw +/- 0.002 vs. C 0.019g/g bw +/- 0.009) (Figure 2B) and, at
sacrifice, large amounts of adipose tissue were conspicuous around the sternum,
kidneys and hip joints. In striking contrast, 24 week old mdx mice were very lean and
had very small epididymal fat pads (Figure 2B); upon opening the abdominal cavity the
epididymal fat pads often could not be seen until the testes were pulled up into the
abdominal cavity and consumption of a high fat did not increase epididymal fat pad
weight in 24 week old mdx mice.
Standardised gastrocnemius muscle weight (g muscle /g bw) was significantly (P=0.04)
heavier in mdx mice fed a control diet compared to C57 mice at 24 weeks of age (C57
0.0050g/g bw +/- 0.0002 vs. mdx 0.0056g/g bw +/- 0.0001) (Figure 2C). Absolute
gastrocnemius muscle weight was also significantly (P<0.05) heavier in mdx mice at
this age (C57 0.152g +/- 0.002 vs. mdx 0.190g +/- 0.004) (data not shown). Neither the
high protein nor high fat diet caused any changes in standardised or absolute
gastrocnemius muscle weight in either strain of mice.
Radley-Crabb and Grounds, 2010. 12
Group 2) 24 - 40 week old sedentary mice subjected to 3 diets. There was no
significant difference in the body weight of 40 week old C57 mice on the control diet
compared to mdx mice (C57 37.3g +/- 2.2 vs. mdx 34.2g +/- 0.76) (Figure 2A), nor did the
high protein diet have any significant effect of the body weight of either strain of mice
(Figure 2A). However, after consuming the high fat diet for 16 weeks the body weight
of 40 week old C57 mice was significantly (P<0.01) increased (approximately 10g)
compared to C57 mice on the control diet (HF 47.3g +/- 1.5 vs. C 37.3 +/- 2.2g).
Consumption of the high fat diet also significantly (P=0.04) increased the body weight
of the older mdx mice (HF 37.2g +/- 1.1 vs. C 34.2g +/- 0.76) (Figure 2A). The largest
increase in body weight (body weight at 40wks minus body weight at 24wks) was seen
in C57 mice on the high fat diet.
The (average group percentage) change in
bodyweight for C57 mice was 116% after control diet, 102% after high protein diet and
141% after high fat diet. The change in body weights for mdx mice were 103% after
control diet, 94% after high protein diet and 110% after high fat diet.
On the control diet the standardised epididymal fat pad weight (g/g bw) was
significantly (P<0.01) increased in 40 week old C57 mice compared to mdx mice (C57
0.027g/g bw +/- 0.005 vs. mdx 0.007g/g bw +/- 0.0003) (Figure 2B). The high protein diet
had no impact on the epididymal fat pad weight in either strain of the older mice. On
the high fat diet, there was a slight increase (but not significant – presumably due to
the large increase in total bodyweight) in standardised epididymal fat pad weight (g
fat/ g bw) of C57 mice (HF 0.032g/g bw +/- 0.002 vs. C 0.027g/g bw +/- 0.005) (Figure
2B). The absolute epididymal fat pad weight (g fat) was significantly (P=0.03) increased
(HF 1.42g +/- 0.06 vs. C 1.16g +/- 0.2) (data not shown) and observations made during
tissue collection showed very large fat pads and pronounced adipose tissue around the
sternum, kidneys, intestines and hip joints. 40 week old mdx mice were still very lean
with no change in the standardised epididymal fat pad weight between 24 and 40
weeks for mdx mice fed a control diet (Figure 2B). The high fat diet significantly
(P=0.035) increased epididymal fat pad weight (HF 0.012g/g bw +/-0.002 vs. C 0.007g/g
bw +/-0.0003) in 40 week old mdx mice (Figure 2B), to a much lesser extent than C57
mice, whereas this effect was not seen in the younger 8-24 week old mdx mice.
Radley-Crabb and Grounds, 2010. 13
Standardised gastrocnemius muscle weight was significantly (P=0.045) heavier in mdx
mice fed a control diet compared to C57 at 40 weeks of age (C57 0.0048g/g bw +/0.0002 vs. mdx 0.0059g/g bw +/- 0.0001) (Figure 2C). Absolute gastrocnemius muscle
weight was also significantly (P<0.05) heavier in mdx mice at this age (C57 0.178g +/0.003 vs. mdx 0.198g +/- 0.005) (data not shown). There was no change in standardised
gastrocnemius muscle weight in mdx mice with age (e.g 24wk old mdx vs 40wk old
mdx), however there was a significant (P<0.05) increase in absolute gastrocnemius
muscle weight with age in C57 mice (24wk 0.152g/ +/- 0.002 vs. 40wk 0.178g +/- 0.003)
(data not shown). The high protein diet had no effect on the muscle weights of either
strain. The high fat diet caused no change in absolute gastrocnemius muscle weight in
either strain of mice (data not shown), thus due to the large increase in body weight in
C57 mice after consuming a high fat diet there was a strikingly significant (P=0.01)
reduction in standardised gastrocnemius muscle weight in C57 mice (HF 0.003g/g bw
+
/- 0.0006 vs. C 0.0048g/g bw +/- 0.0002) (Figure 2C), yet had no change on the
standardised gastrocnemius muscle weight in mdx mice.
Myofibre Size (control diet only).
There was no difference in myofibre size (CSA) in the tibialis anterior (TA) muscle from
12 week old C57 and mdx mice on the control diet (Figure 3). Between 12 -24 weeks of
age, dystrophic myofibres in the mdx TA continue to grow (hypertrophy) and were
significantly larger than C57 myofibres at 24 weeks (P=0.02) (Figure 3). Dystrophic
myofibres do not continue to infinitely hypertrophy and myofibre size in 40 week old
mdx mice was significantly reduced (P=0.025) compared to 24 weeks and there was no
difference in myofibre size between 12 and 40 week old mdx mice. There was no
change in myofibre size between 12, 24 and 40 week old C57 mice.
Muscle morphology and dystropathology.
8 – 24 week old sedentary mice. Myofibre necrosis (% cross sectional area) was
significantly (P=0.04) decreased in the quadriceps muscle of 24 week old mdx mice fed
the high fat diet, compared to the control diet (HF 1.49% +/-1.7 vs. C 3.3% +/-0.4)
Radley-Crabb and Grounds, 2010. 14
(Figure 4A); however the high protein diet had no effect on myofibre necrosis
(compared to control diet). Myofibre necrosis is not a feature of control C57 mice and
therefore it was not measured. There were no significant differences in adipocyte
content of the quadriceps muscle in 24 week old C57 and mdx mice after consuming
any of the 3 diets.
24 – 40 week old sedentary mice. There was no change in the level of myofibre
necrosis in the quadriceps muscle between 24 and 40 week old mdx mice (Figure 4A)
and no effect of the high protein nor high fat diet on myofibre necrosis in 40 week old
mdx mice (Figure 4A). The high protein diet had no effect on the adipocyte content of
the quadriceps muscle in either strain of 40 week old mice. Adipocyte content of the
quadriceps was significantly (P=0.03) increased with age in C57 mice (24wk 0.06% +/0.014 vs. 40wk 1.16% +/- 0.035) fed a control diet (Figure 4B) and was significantly
(P<0.001) increased further in C57 mice fed a high fat diet (HF 2.94% +/- 0.72 vs. C
1.16% +/- 0.35) (Figure 4B). The high fat diet also significantly increased adipocyte
content between 24 and 40 week old mdx mice (Figure 4B).
Blood serum CK levels are always lower (approximately 10 fold) in C57 mice compared
to mdx mice (Grounds et al, 2008b; Radley-Crabb et al, 2010; Spurney et al, 2009) and
were not measured for C57 mice in this study. For the mdx mice there was no change
in serum CK level between 24 and 40 weeks (24wk 3171U/L +/- 460 vs. 40wk 2791U/L
+
/- 422) and no change in CK level after consuming either a high fat or high protein diet
for both 24 and 40 week old mdx mice (data not shown).
Gene expression.
8 – 24 week old sedentary mice. mRNA levels of TNF and IL1β were significantly
elevated (approximately 2 fold) in the gastrocnemius muscle from 24 week old mdx
compared to C57 mice (Figure 5A&B). Consumption of the high fat diet significantly
increased the mRNA levels of all 3 inflammatory cytokines (TNF, IL1β and IL-6) in C57
mice (Figure 5A-C), however there was no change in 24 week old mdx mice.
Radley-Crabb and Grounds, 2010. 15
Levels of mRNA for PPAR alpha, PPAR delta/beta and PGC-1α alpha (but not PPAR
gamma) were significantly decreased (approximately 2 fold) in the gastrocnemius
muscle from 24 week old mdx compared to C57 mice (Figure 6A,C,D). Expression of the
3 PPARs was unchanged in either strain by consumption of the high fat diet. However,
PGC-1α was significantly reduced (approximately 2 fold) in 24 week old C57 mice fed
the high fat compared to the control diet.
24 – 40 week old sedentary mice. mRNA levels of TNF were significantly elevated
(approximately 2 fold) with age (24wk C57 vs. 40wk C57) in C57 mice (Figure 5A). The
high fat diet significantly increased (approximately 2x fold) TNF and IL-6 mRNA in 40
week old C57 mice and IL-6 mRNA (approximately 3x fold) in mdx mice (Figure 5A & C).
mRNA levels of PGC-1α were significantly decreased (approximately 2x fold) with age
(24wk C57 vs. 40wk C57) in C57 mice (Figure 5D) and PPAR alpha and PPAR delta/beta
mRNAs were significantly decreased (approximately 2x fold) in the gastrocnemius
muscle of 40 week old mdx mice compared to C57 mice (Figure 5A & C). The high fat
diet caused no change in the mRNA level of the 3 PPARs or PGC-1α in either strain.
Group 3) 8-12 week old voluntarily exercised mice.
Dystrophic mdx mice were voluntarily exercised for 4 weeks to increase the low level
of dystropathology in skeletal muscles (Radley et al, 2008; Radley and Grounds, 2006).
Increasing the extent of dystropathology allowed for further evaluation of the
beneficial effects of a high protein or high fat diet in mdx mice.
Food consumption.
There was no significant difference in average food consumption (g/ g bw /day) across
the 3 custom diets in both sedentary and exercised mdx mice (Figure 7A); however,
food consumption of all 3 custom diets (g /g bw /day) was significantly (P<0.04)
increased (approximately 30%) by voluntary exercise (Figure 7A). Standardised energy
intake (kj /g bw/day) was unchanged in sedentary 8-12 week old mdx mice across the
3 diets (Figure 7B); however energy intake was significantly increased in exercised mdx
mice for all three diets. Absolute energy intake (kj/day) showed the same pattern as
standardised energy intake (data not shown). Voluntarily exercised mdx mice on the
Radley-Crabb and Grounds, 2010. 16
high fat diet had a higher standardised energy intake (kj /g bw /day) than exercised
mdx mice on both the control and high protein diet (C Ex 1.91 kj/g bw/day +/- 0.08 vs.
HF Ex 2.28 kj/g bw/day +/- 0.08) (Figure 7B). Absolute energy intake (kj /day) was also
significantly increased in exercised mice on the high fat diet (C Ex 50.4 kj +/- 4.2 vs. HF
Ex 63.7 kj +/- 7.6) (data not shown).
Body composition.
Voluntary wheel exercise for 4 weeks caused no change in the body weight of 12 week
old mdx mice (sed 29.1g +/- 2.8 vs. ex 27.5g +/- 1.9), neither did consumption of a high
protein or high fat diet (Figure 8A). Exercising 8-12 week old mdx mice significantly
(P=0.03) reduced the standardised epididymal fat pad weight (g fat/ g bw), compared
to sedentary mdx mice on a control diet (sed 0.0081g +/- 0.0005 vs. ex 0.0051g +/0.0002) (Figure 8B). Neither the high protein nor the high fat diet had any significant
effect on the epididymal fat pad weight in sedentary 12wk old mdx mice; however the
high fat diet significantly (P=0.045) increased epidiymal fat pad weight in exercised
mdx mice (HF 0.0064g +/- 0.0006 vs. C 0.0051 +/- 0.0002) (Figure 7B). No significant
change in standardised or absolute gastrocnemius muscle weight was seen after
voluntary exercise or diet change (Figure 8C).
Dystropathology.
Exercise was completely voluntary and distance run is an indirect indicator of muscle
function and exercise capacity. Mdx mice fed the high fat diet ran significantly
(P=0.003) further during 4 weeks of exercise (approximately 50% further) compared to
mdx mice on the control diet (HF 221.8km +/- 13.2 vs. C 152.4km +/- 12.1) (Figure 9A)
and there was no effect of a high protein diet compared with control diet.
Voluntary exercise significantly (P=0.02) increased (approximately 2x fold) myofibre
necrosis (% CSA) in the quadriceps muscle from 12 week old mdx mice (sed mdx 3.68%
+
/- 0.49 vs. ex mdx 8.75% +/- 0.91). The high fat diet for 4 weeks had no effect on
myofibre necrosis in either voluntarily exercised or sedentary mdx mice. The high
protein diet had no effect on sedentary mice, but significantly (P=0.04) increased
Radley-Crabb and Grounds, 2010. 17
myofibre necrosis in exercised mdx mice (HP 12.51% +/- 2.00 vs. C 8.75 +/- 0.91) (Figure
9B).
Blood serum CK level was significantly (P=0.04) higher (approximately 3x fold) in
voluntarily exercised compared to sedentary 12 week old mdx mice (sed mdx 4697U/L
+
/- 1898 vs. ex mdx 12913U/L +/- 4537). The CK levels in mdx mice were not affected by
a high protein or high fat diet (data not shown).
Exercise did not change the adipocyte content of the quadriceps muscle from 12 week
old mdx mice (sed mdx 0.54% +/- 0.07 vs. ex mdx 0.43% +/- 0.07), nor was there any
change in adipocyte content after consuming a high protein or high fat diet in
sedentary or exercised mice (data not shown).
Gene expression.
Voluntary exercise (for 4 weeks) and/or a high fat diet caused no significant change in
the mRNA levels of TNF, IL1β or IL-6 in 12 week old mdx mice (data not shown).
Consumption of the high fat diet had no significant affect on mRNA levels of the 3
PPARs or PGC-1α in sedentary 12 week old mdx mice; however the combination of
voluntary exercise and a high fat diet significantly increased (approximately 3x fold)
the mRNA levels of PPAR alpha, PPAR gamma, PPAR delta/beta and PGC-1α (Figure
10A-D).
Discussion:
Sedentary mdx and C57 mice.
To our knowledge, the response of adult mdx mice to a high fat diet has not been
previously described; furthermore, while it is widely documented that C57Bl/6J mice
are highly susceptible to diet induced obesity (obesity-prone) (Alexander et al, 2006),
the effect of a high fat diet on C57Bl/10 mice does not seem to have been reported. It
is commonly accepted that obesity is the direct result of an imbalance between energy
intake and energy expenditure, that long-term consumption of a high fat diet increases
deposition of adipose tissue and eventually leads to obesity in both sedentary
laboratory rodents and humans and that obesity is associated with numerous changes
Radley-Crabb and Grounds, 2010. 18
in cell signalling that are believed to underlie much of the morbidity associated with
the condition [Reviewed in (Buettner et al, 2007; Pedersen, 2010; Stienstra et al,
2007)].
Dystrophic mdx mice were significantly heavier than control C57 mice at both 8 and 24
weeks of age (Figure 2A), primarily due to a well described increase in lean muscle
mass (Anderson et al, 1987; Connolly et al, 2001; Coulton et al, 1988; Mokhtarian et al,
1996; Peter and Crosbie, 2006; Shavlakadze et al, 2010; Spurney et al, 2009). In
contrast, 40 week old mdx mice were the same weight as C57 mice; this equalising in
bodyweight occurred at approximately 32 weeks of age (data not shown) and was
most likely due to an increase in body fat in C57 mice (Figure 2B). The body weight of
mdx mice did not increase between 24 and 40 weeks of age, nor was there a change in
standardised epididymal fat pad nor standardised gastrocnemius muscle weights
(Figure 2A), supporting reports of stabilisation of the dystrophic phenotype in adult
mdx mice (Grounds et al, 2008b; Shavlakadze et al, 2010). Both 24 and 40 week old
mdx were very lean, with significantly smaller epididymal fat pads, larger
gastrocnemius muscles and larger myofibres CSA compared to C57 mice (Figures 2B,
2C, 3).
The muscular body composition of mdx mice (<40 weeks) is in stark contrast to DMD
patients who can move between the spectra of over-nutrition (and obesity) to undernutrition within their shortened lifespan, and lose muscle mass at a rate of 4% per year
as adults (Davidson and Truby, 2009; Griffiths and Edwards, 1988; Leighton, 2003). It is
important to consider that the striking loss of muscle mass in DMD patients occurs
during the growth phase of the boys (approximately 20 years). In mice however, the
damaging main growth phase is exceedingly short by comparison (approximately 6
weeks) (Grounds, 2008; Grounds et al, 2008b) with a reduction in myofibre necrosis
occurring after growth has ceased in mdx mice.
There was significant myofibre hypertrophy in 24 week old mdx mice (compared to all
C57 and 12 week old mdx mice); however, dystrophic myofibres do not continue to
hypertrophy indefinitely, and by 40 weeks of age myofibre size returned to the same
Radley-Crabb and Grounds, 2010. 19
size as 12 week old mdx mice (Figure 3). We propose that the reduced myofibre size in
40 week old mdx mice was due to myofibre splitting or branching. Shavlakadze et al
(2010) reported no difference in myofibre size in the quadriceps muscle between 12
and 52 weeks due to significant myofibre splitting in 52 week old mdx mice
(Shavlakadze et al, 2010). Our study examined myofibre size at 24 weeks and identified
significant myofibre hypertrophy in mdx mice at this age, so it seems that 24 weeks is
as a suitable time point to visualise myofibre hypertrophy in the TA muscle before
myofibre splitting occurs (by 40 weeks onwards).
The body composition of dystrophic mdx mice changed between 12 and 24 weeks of
age, with significant increases in body weight, absolute and standardised
gastrocnemius muscle weight and individual myofibre CSA (µm2) (Figures 2A, 3 & 8A).
The 8-24 week old mdx mice had similar food consumption and energy intake levels
across the 3 diets compared to age matched C57 mice (Figure 1A-C) with both strains
on the high fat diet showing a slight increase in energy intake (kj/day) compared to the
control diet. Yet, there was a large difference in the way the two strains of mice
responded to the high fat diet. The resistance of dystrophic mice (<24 weeks of age) to
the high fat diet (i.e. they did not deposit significant body fat) may be due to excess
dietary fat (or increased energy intake) assisting with muscle growth and hypertrophy
of individual myofibres (Figure 3). Between 24 and 40 weeks of age, there was no
change in bodyweight (Figure 2A) or gastrocnemius muscle weight (absolute or
standardised) and myofibre CSA was significantly reduced (Figure 3) for mdx mice on a
control diet. The older mdx mice (24-40 weeks) were susceptible to the negative
effects of a high fat diet, showing significant increases in total bodyweight and
standardised fat pad weight (Figures 2A&B). This altered trend is presumably due to
the large increase in energy intake (Figure 1B&C) without the corresponding high
energy expenditure associated with ongoing myogenesis/regeneration, due to a
progressive reduction of myofibre necrosis in old mdx mice; e.g. 6% myofibre necrosis
in 12 week old mdx mice (Radley-Crabb et al, submitted) reducing to approximately 2%
by 40 weeks of age.
Radley-Crabb and Grounds, 2010. 20
In contrast to mdx mice, after consumption of a high fat diet (for 16 weeks) C57 male
mice at both 24 and 40 weeks exhibit significantly increased bodyweight primarily due
to a gain in body fat (Figure 2A&B) and significant deposition of adipocytes in the
quadriceps muscle of 40 week old mice (Figure 4B). The increase in daily energy intake
(kj/day) in 8-24 week old C57 mice on the high fat diet is only minor compared to mice
on the control diet (Figure 1B), but when accumulated chronically it was enough to
cause large changes in body composition; whereas the increase in energy intake
(kj/day) in 24-40 week old C57 mice on the high fat diet is significantly increased
compared to mice on the control diet (Figure 1B).
In contrast to the high fat diet (16% fat, 19.4% protein), a high protein diet (50%
protein, 7% fat) had little impact on the body composition of 8 – 40 week old C57 and
mdx mice. Initially these two diets were designed to be isocaloric based on digestible
energy content (HP – 18.2 MJ/Kg vs. HF – 18.1 MJ/Kg) (Table 1). Major differences in
body composition were seen in response to the high fat and high protein diets and it
became apparent that metabolisable energy content was a more appropriate way to
assess energy intake of the mice. There are large differences in the metabolisable
energy content of the high fat and high protein diet (HP – 14.09 MJ/Kg vs. HF – 16.96
MJ/Kg) due to the incomplete in vivo oxidation of dietary proteins which may explain
the major differences in energy intake and thus body composition in response to the
two diets. A high protein diet or supplementation with certain amino acids (such as
leucine) can stimulate skeletal muscle protein synthesis in both young and adult
animals, but there is a threshold of protein requirement for maximum growth and
once reached, additional protein intake does not further stimulate protein synthesis in
skeletal muscle [Reviewed in (Shavlakadze and Grounds, 2006)]. Therefore, it is
possible that in well nourished sedentary mice (with adequate protein intake), the high
protein diet had no effect on muscle mass. While it is possible that a beneficial
response to high protein consumption might be seen in combination with resistance
exercise, such demanding exercise is usually not recommended for dystrophic boys or
animal models of DMD due to likely additional damage to dystrophic muscles.
Radley-Crabb and Grounds, 2010. 21
Our initial hypothesis was that a high protein and/or high fat diet would help to
maintain myofibre integrity and thus reduce dystropathology in mdx mice.
Consumption of either diet for 16 weeks did not reduce levels of serum CK (an
indicator of myofibre leakiness), however, the high fat diet did significantly reduce
(approximately 2.5 fold) myofibre necrosis in the quadriceps muscle of 24 week old
mdx mice. It is possible that a high fat diet (and increased energy intake) assists mdx
mice metabolically and thus helps them to maintain skeletal muscle structure and
mass, or that increased lipid coming from the high fat diet stabilises fragile membrane
lipids and prevents myofibre necrosis.
The lack of any benefits with the high protein diet contrasts with studies in mdx mice
that show mildly reduced dystropathology after dietary supplementation of creatine,
taurine and glutamine (both alone and in combination with prednisolone) (Cozzoli et
al, 2010; De Luca et al, 2003; Granchelli et al, 2000; Passaquin et al, 2002; Radley et al,
2007). Taurine and glutamine are not substrates for protein synthesis, but may
improve dystropathology via regulation of calcium homeostasis (Conte Camerino et al,
2004) or by inhibiting whole body protein degradation (Mok et al, 2006). Amino acid
supplementation is also widely used amongst DMD patients despite there being no
conclusive supporting evidence in the literature (Bushby et al, 2010; Davidson and
Truby, 2009; Radley et al, 2007). It is important to consider possible counteracting
effects when combined interventions are administered. Our data from mdx mice on
the high protein diet (which contains increased concentrations of the essential amino
acids such as leucine and phenylalanine which are used in protein synthesis, but no
increase in metabolisable energy - Table 1) do not support such an intervention and
elevated myofibre necrosis was observed in voluntarily exercised mdx mice.
A prolonged positive imbalance between energy intake and expenditure resulting in
obesity, is associated with chronic inflammation and may be a potential mechanism by
which obesity leads to insulin resistance (in both humans and mice) [Reviewed in
(Arkan et al, 2005; Stienstra et al, 2007; Wisse, 2004)]. Expression of 3 major proinflammatory cytokines, TNF, IL1β and IL6 was up-regulated in the gastrocnemius
Radley-Crabb and Grounds, 2010. 22
muscle of both 24 and 40 week old C57 mice after consumption of the high fat diet
(Figure 5A-C). These increases in gene expression were not seen in mdx mice (Figures
5A-C) which demonstrates, in addition to the lack of changes in body composition, the
resistance of mdx mice (particularly <24 weeks) to a high fat diet. Inflammation plays a
major role in myofibre necrosis and regeneration in skeletal muscle, thus the
inflammatory state can be an in direct indicator of the extent of dystropathology
(Grounds et al, 2008a; Radley et al, 2008). Consistent with the lack of significant
adiposity, there were no major changes in cytokine gene expression in mdx mice after
consuming a high fat diet.
The high fat diet significantly reduced PGC-1α gene expression in 24 week old C57 mice
but, apart from this, there were no other significant changes in PPAR expression
caused by a high fat diet in either strain of mice. Compared with C57 mice, the levels of
PGC-1α, PPAR alpha and PPAR delta/beta were all much lower (approximately half) in
mdx mice on a control diet; again endorsing altered metabolic processes in dystrophic
mdx mice.
Due to the multiple systemic effects that the 3 PPARs and PGC-1α have on whole body
metabolism (e.g. lipid metabolism) (Ehrenborg and Krook, 2009; Liang and Ward, 2006;
Stienstra et al, 2007) we assume that decreased gene expression of the 3 PPARs and
PGC-1α in dystrophic skeletal muscle would have a wide range of effects. However the
precise consequence(s) in dystrophic skeletal muscle, to our knowledge, are not
documented. Decreased PGC-1α expression in skeletal muscles of C57Bl/6J mice
decreases exercise capacity and fatigue resistance [Reviewed in (Liang and Ward,
2006)] and low levels of PGC-1α mRNA in mdx mice on a control diet, correspond with
decreased exercise capacity and increased fatigue that are common features of
dystrophic muscle (Grounds et al, 2008b; Hara et al, 2002; Piers et al, 2010). It has also
been shown that several gene programmes linked to PGC-1α are dysregulated in
dystrophic muscle (e.g mitochondrial function, calcium handling and ROS) and
introduction of a muscle specific PGC-1α transgene into mdx mice improved muscle
function and structure, possibly via up-regulation of utrophin (Davies and Khurana,
2007; Handschin et al, 2007). Overall there seems to be an association with decreased
Radley-Crabb and Grounds, 2010. 23
PGC-1α expression having detrimental effects in both dystrophic and normal muscle.
Although the combined consequences of generally PGC-1α, PPAR alpha and PPAR
delta/beta in dystrophic (compared with normal) muscle are not clear, they are all
elevated in mdx muscles in response to voluntary exercise combined with a high fat
diet (see below) with beneficial effects.
Voluntarily exercised mice.
In order to further examine the potential benefits of a high fat or high protein diet on
dystrophic muscle, 8-12 week old mdx mice were voluntarily exercised, to exacerbate
the dystropathology (Hodgetts et al, 2006; Radley et al, 2008; Radley and Grounds,
2006). The 12 week old mdx mice were very lean, yet there was a significant further
reduction in epididymal fat pad weight after 4 weeks of voluntary exercise (Figure 8B).
It appears that mdx mice were able to maintain body weight and muscle mass (Figures
8A & C) over 4 weeks of voluntary exercise by increasing food consumption and energy
intake (Figure 7A&B).
Despite a high fat diet reducing myofibre necrosis in sedentary 24 week old mdx mice
(Figure 4), this diet had no significant benefit on myofibre necrosis or serum CK levels
in voluntarily exercised mdx mice. Energy intake was significantly increased in
voluntarily exercised mdx on the high fat diet compared to mdx mice on the control
diet (Figure 7B) and this may have enabled mdx mice to run significantly more
(approximately 50%); and yet this increased exercise did not result in increased muscle
damage (e.g. myofibre necrosis or serum CK level). It appears that a high fat diet is
assisting energy deficient mdx mice metabolically, providing them with additional
energy to run further while also maintaining myofibre integrity.
A striking increase in the mRNA levels of PPAR alpha, PPAR gamma, PPAR delta/beta
and PGC-1α was seen in the gastrocnemius muscle of exercised mdx mice on the high
fat diet.
Exercise is known to increase PGC-1α expression (due to increased
neuromuscular input and elevated levels of MEF2 and CREB expression with exercise)
(Liang and Ward, 2006) and increases in PGC-1α and PPAR delta/beta are both
beneficial to mdx dystropatholgy in vivo (Handschin et al, 2007; Miura et al, 2009).
Radley-Crabb and Grounds, 2010. 24
Therefore it is proposed that the increased capacity for exercise in dystrophic mdx
mice fed a high fat diet may be modulated, in part, via up regulation of the 3 PPARs
and PGC-1α.
Conclusion:
This research clearly identifies an ‘altered’ response to a high fat diet in dystrophic
mdx mice compared to C57 controls; this response was pronounced in young mdx
mice <24 weeks old, but diminished with age (by 40 weeks). Energy deficient mdx mice
appear to be utilising excess dietary fat to reduce the severity of dystropathology and
increase voluntary exercise ability. The high protein diet had no significant effects on
the body composition of either strain of mice and no benefit on mdx dystropathology
(this may be due in part to the fact that while it was isocaloric for digestible energy it
was lower in metbolisable energy content compared with the high fat diet). While it is
appreciated that in corticosteroid treated inactive DMD patients, a high fat diet may
negatively contribute to obesity and thus impact on disease severity (especially since
corticosteroid usage is known to contribute to obesity and Cushingoid symptoms), our
new data highlight the potential benefits of a high fat diet on dystrophic muscle. It is
not yet clear if the extent to which this benefit of a high fat diet is due largely to
increased energy (kj) per se, relative to specific benefits of increased dietary fat on
dystrophic muscles. Clearly careful management is required to achieve a fine balance
between the increased metabolic demands of dystrophic muscle, disease treatment
and the potential negative effects of a high fat diet.
Acknowledgements:
The authors gratefully acknowledge the expert assistance of Dr Marta Fiorotto of
Baylor College, Houston, USA and her valuable comments regarding the manuscript.
The authors also thank Mr Greg Cozens (School of Anatomy & Human Biology, UWA)
for excellent technical assistance, Dr Andrew Hoey (University of Southern
Queensland) for consultation regarding the preliminary studies and Mr Warren Potts
(Specialty Feeds, WA) for design and manufacture of the 3 semi-pure diets. Research
funding from the Australian National Health and Medical Research Council (MG) and
Australian Postgraduate Award Scholarships (HR-C) are gratefully acknowledged.
Radley-Crabb and Grounds, 2010. 25
Table captions:
Table 1. Diet specifications for the 3 semi-pure diets used in this study: control
(AIN93g), high protein (SF00-252) and high fat (SF06-40). Energy intake was calculated
based on the metabolisable energy content of each diet. All diets were manufactured
by Specialty Feeds, Glen Forrest, WA.
Figure captions:
Figure 1. Average daily food consumption (g/ g bw/ day) (A), absolute energy intake
(kj/ day) (B) and standardised energy intake (kj /g bw /day) (C) for C57 and mdx
mice; a comparison of sedentary mice on a control diet, high fat diet or high protein
diet between 8-24 weeks of age and 24-40 weeks of age. Bars represent standard
error. N= 8 for all groups. A,B,C denotes significant differences, groups with different
letters are significantly different from each other (P<0.05).
Figure 2. Body composition of sedentary 24 week and 40 week old C57 and mdx
mice; a comparison of sedentary mice on a control diet, high fat diet or high protein
diet. A) Total body weight. B) Standardised epidiymal fat pad weight (g fat/g bw). C)
Standardised gastrocnemius muscle weight (g muscle /g bw). Bars represent standard
error. N= 8 for all groups. A,B,C,D denotes significant differences, groups with different
letters are significantly different from each other (P<0.05).
Figure 3. Myofibre cross-sectional area in 12, 24 and 40 week old sedentary C57 and
mdx mice. Bars represent standard error. N= 6 mice for all groups (at least 500
myofibres measured per mouse). A,B denotes significant differences, groups with
different letters are significantly different from each other (P<0.05).
Figure 4. Myofibre necrosis in sedentary mdx mice and adipocyte content of both
C57 and mdx mice aged 24 and 40 weeks; a comparison of sedentary mice on a
control diet with mice on either a high fat or high protein diet. A) Myofibre Necrosis in
Radley-Crabb and Grounds, 2010. 26
the quadriceps muscle (mdx only). B) Adipocyte content in the quadriceps muscle of
both strains. Bars represent standard error. N= 8 for all groups. A,B,C denotes
significant differences, groups with different letters are significantly different from
each other (P<0.05).
Figure 5. Gene expression (mRNA) changes in the gastrocnemius muscle of 24 and 40
week old C57 and mdx mice; a comparison of sedentary mice on a control diet with
mice on a high fat diet. A) Tumour Necrosis Factor (TNF). B) Interleukin 1β (IL-1β). C)
Interleukin 6 (IL-6). N= 8 mice per group. A,B,C denotes significant differences, groups
with different letters are significantly different from each other (P<0.05).
Figure 6. Gene expression (mRNA) changes in the gastrocnemius muscle of 24 and 40
week old C57 and mdx mice; a comparison of sedentary mice on a control diet with
mice on a high fat diet. A) Peroxisome proliferator-activated receptor (PPAR) alpha. B)
PPAR gamma. C) PPAR delta/beta. D) Peroxisome proliferator-activated receptor
gamma coactivator 1 alpha (PGC1-α). N= 8 mice per group. A,B denotes significant
differences, groups with different letters are significantly different from each other
(P<0.05).
Figure 7. Average daily food consumption (g/ g bw/ day) (A) and energy intake (kj /g
bw /day) (B) for mdx mice. Data for dietary consumption between 8-12 weeks of age
is shown for both sedentary and voluntarily exercise mdx mice on either a control diet,
high fat diet or a high protein diet. Bars represent standard error. N= 8 for all groups.
A,B,C denotes significant differences, groups with different letters are significantly
different from each other (P<0.05).
Figure 8. Body composition of 12 week old sedentary and voluntarily exercised mdx
mice; a comparison of mice on a control diet with a high fat diet or high protein diet.
A) Total body weight. B) Standardised epidiymal fat pad weight (g fat /g bw). C)
Standardised gastrocnemius muscle weight (g muscle /g bw). Bars represent standard
Radley-Crabb and Grounds, 2010. 27
error. N= 8 for all groups. A,B denotes significant differences, groups with different
letters are significantly different from each other (P<0.05).
Figure 9. Total distance run (km) over 4 weeks of voluntary exercise and myofibre
necrosis in sedentary and exercised mdx 12 week old mice; a comparison of mice on a
control diet, high fat diet or high protein diet. A) Total distance run (km) over 4 weeks
of voluntary exercise (mdx only). B) Myofibre necrosis in the quadriceps muscle (mdx
only). Bars represent standard error. N= 8 for all groups. A,B,C denotes significant
differences, groups with different letters are significantly different from each other
(P<0.05).
Figure 10. Gene expression (mRNA) changes in the gastrocnemius muscle of 12 week
old sedentary and voluntarily exercised mdx mice; a comparison of sedentary mice on
a control diet with mice on a high fat diet. A) PPAR alpha. B) PPAR gamma. C) PPAR
delta/beta. D) PGC1-α. N= 8 mice per group. A,B denotes significant differences,
groups with different letters are significantly different from each other (P<0.05).
Radley-Crabb and Grounds, 2010. 28
DIET SPECIFICATIONS
AIN93G - Control
SF00-252 - High protein
SF06-040 - High Fat
Calculated Nutritional parameters
Calculated Nutritional parameters
Calculated Nutritional parameters
Total Protein
19.40%
Total Protein
52%
Total Protein
19.40%
Total Fat
7%
Total Fat
7%
Total Fat
16%
Total Carbohydrate
57%
Total Carbohydrate
27%
Total Carbohydrate
50%
Crude Fibre
4.70%
Crude Fibre
4.70%
Crude Fibre
4.70%
Acid Detergent Fibre
4.70%
Acid Detergent Fibre
4.70%
Acid Detergent Fibre
4.70%
% digestible energy from lipid
16.00%
% digestible energy from lipid
16.00%
% digestible energy from lipid
32.00%
% digestible energy from protein
21.00%
% digestible energy from protein
55.00%
% digestible energy from protein
19.00%
Digestible Energy (MJ/Kg)
16.1
Digestible Energy (MJ/Kg)
18.2
Digestible Energy (MJ/Kg)
18.1
% metabolisable energy from lipid
17.79%
% metabolisable energy from lipid
18.70%
% metabolisable energy from lipid
35.80%
% metabolisable energy from protein 16.30%
% metabolisable energy from protein 46.10%
% metabolisable energy from protein 14.20%
Metabolisable energy (MJ/Kg)
Metabolisable energy (MJ/Kg)
Metabolisable energy (MJ/Kg)
14.82
Major ingredient
14.09
Major ingredients
16.96
Major ingredients
Casein (Acid)
200g/Kg
Casein (Acid)
540g/Kg
Casein (Acid)
200g/Kg
DL Methionine
3.0g/Kg
DL Methionine
1.4g/Kg
DL Methionine
3.0g/Kg
Sucrose
100g/Kg
Sucrose
100g/Kg
Sucrose
100g/Kg
Wheat Starch
404g/Kg
Wheat Starch
66g/Kg
Wheat Starch
313g/Kg
Dextrinised Starch
132g/Kg
Dextrinised Starch
132g/Kg
Dextrinised Starch
132g/Kg
Cellulose
50g/Kg
Cellulose
50g/Kg
Cellulose
50g/Kg
Canola Oil
70g/Kg
Canola Oil
70g/Kg
Canola Oil
124g/Kg
Cocoa Butter
-
Cocoa Butter
-
Cocoa Butter
37g/kg
Calculated Amino Acids
Calculated Amino Acids
Calculated Amino Acids
Valine
1.30%
Valine
3.40%
Valine
1.30%
Leucine
1.80%
Leucine
4.90%
Leucine
1.80%
Isoleucine
0.90%
Isoleucine
2.40%
Isoleucine
0.90%
Threonine
0.80%
Threonine
2.10%
Threonine
0.80%
Methionine
0.80%
Methionine
1.60%
Methionine
0.80%
Cystine
0.06%
Cystine
0.15%
Cystine
0.05%
Lysine
1.50%
Lysine
4.00%
Lysine
1.50%
Phenylalanine
1.00%
Phenylalanine
2.70%
Phenylalanine
1.00%
Tyrosine
1.00%
Tyrosine
2.80%
Tyrosine
1.00%
Tryptophan
0.30%
Tryptophan
0.70%
Tryptophan
0.30%
Calculated Fat Composition
Calculated Fat Composition
Calculated Fat Composition
Myristic Acid 14:0
trace
Myristic Acid 14:0
trace
Myristic Acid 14:0
0.02%
Palmitic Acid 16:0
0.40%
Palmitic Acid 16:0
0.30%
Palmitic Acid 16:0
1.50%
Stearic Acid 18:0
0.10%
Stearic Acid 18:0
0.10%
Stearic Acid 18:0
1.60%
Palmitoleic Acid 16:1
trace
Palmitoleic Acid 16:1
trace
Palmitoleic Acid 16:1
0.04%
Oleic Acid 18:1
4.20%
Oleic Acid 18:1
3.90%
Oleic Acid 18:1
8.10%
Gadoleic Acid 20:1
0.10%
Gadoleic Acid 20:1
trace
Gadoleic Acid 20:1
0.10%
Linoleic Acid 18:2 n6
1.30%
Linoleic Acid 18:2 n6
1.50%
Linoleic Acid 18:2 n6
2.80%
a Linolenic Acid 18:3 n3
0.70%
a Linolenic Acid 18:3 n3
1%
a Linolenic Acid 18:3 n3
1.70%
Total n6 fats
1.30%
Total n6 fats
1.51%
Total n6 fats
2.79%
Total n3 fats
0.78%
Total n3 fats
0.98%
Total n3 fats
1.75%
Total Saturated fats
0.50%
Total Saturated fats
0.50%
Total Saturated fats
3.26%
Total monounsaturated fats
3.98%
Total monounsaturated fats
3.98%
Total monounsaturated fats
8.27%
Total polyunsaturated fats
2.50%
Total polyunsaturated fats
2.50%
Total polyunsaturated fats
4.55%
Table 1.
Radley-Crabb and Grounds, 2010. 29
A
0.12
Food consumption ( g/ g bw /day)
A A A
A AB AB
B C B
B BC B
Control
0.08
High Fat
0.04
High Protein
0.00
C57 8-24wks
mdx 8-24wks
C57 24-40wks
mdx 24-40wks
B
60.00
A AC B
A A B
A C AB
A C AB
Energy intake (kj /day)
50.00
40.00
Control
30.00
High Fat
20.00
High Protein
10.00
0.00
C57 8-24wks
mdx 8-24wks
C57 24-40wks
A A B
A A B
B B B
mdx 24-40wks
B C
C
B
Energy intake (kj / g bw /day)
2.00
Control
1.50
High Fat
1.00
High Protein
0.50
0.00
C57 8-24wks
Figure 1.
mdx 8-24wks
C57 24-40wks
mdx 24-40wks
Radley-Crabb and Grounds, 2010. 30
A
55
Final Bodyweight (g)
50
A B A
BD B B
BD C B
B D AB
Control
45
40
High Fat
35
High Protein
30
25
20
C57 24 wks
mdx 24 wks
C57 40wks
mdx 40 wks
0.04
Std Epididymal fat pad (g/g bw)
A B AD
C C C
AB B AB
B
C D C
0.03
Control
High Fat
0.02
High Protein
0.01
0.00
C57 24 wks
mdx 24 wks
C57 40wks
mdx 40 wks
0.008
C
Std gastroc weight (g/g bw)
A A A
B B B
A C A
B B B
0.006
Control
High Fat
0.004
High Protein
0.002
0.000
C57 24 wks
Figure 2.
mdx 24 wks
C57 40wks
mdx 40 wks
Radley-Crabb and Grounds, 2010. 31
6000
A
A
A
B
A
A
Myofibre CSA (um)
5000
4000
C57
3000
mdx
2000
1000
0
12wks
Figure 3.
24wks
40wks
Radley-Crabb and Grounds, 2010. 32
Necrosis in quadricpes (% CSA)
4
A
B
AB
AB
B
A
B
3
Control
2
High Fat
High Protein
1
0
24wks mdx
Adipocytes in quadriceps (% CSA)
4.0
A A AB
A A AB
40wks mdx
B C AB
B
AB B AB
3.0
Control
High Fat
2.0
High Protein
1.0
0.0
C57Bl/10 24 wks
Figure 4.
mdx 24 wks
C57Bl/10 40wks
mdx 40 wks
Radley-Crabb and Grounds, 2010. 33
A) TNF
TNF gene expression /L19 (gastroc)
0.0035
0.003
A
B
B
B
B
C
BC
BC
0.0025
Control
0.002
High Fat
0.0015
0.001
0.0005
0
C57 24wks
IL1B gene expression / L19 (gastroc)
0.014
mdx 24wks
C57 40wks
mdx 40wks
B) IL1-β
A
B
B
B
AB B
B
B
0.012
0.01
Control
0.008
High fat
0.006
0.004
0.002
0
C57 24wks
mdx 24wks
C57 40wks
mdx 40wks
C) IL-6
IL-6 gene expression /L19 (gastroc)
0.008
A
B
A
AB
A
B
A
B
0.006
Control
0.004
High Fat
0.002
0
C57 24wks
Figure 5.
mdx 24wks
C57 40wks
mdx 40wks
Radley-Crabb and Grounds, 2010. 34
PPAR alpha gene expression / L19
0.08
A) alpha
A
A
B
B
A
A
B
B
0.06
Control
0.04
High Fat
0.02
0
C57 24wks
mdx 24wks
C57 40wks
mdx 40wks
PPAR gamma gene expression /L19
0.00035
B) gamma
A
0.0003
A
A
A
AB B
A
AB
0.00025
Control
0.0002
High Fat
0.00015
0.0001
5E-05
0
C57 24wks
PPAR delta/beta gene expression /L19
3.5
A
A
mdx 24wks
B
B
C57 40wks
A
A
mdx 40wks
B
B
C)
delta/beta
3
2.5
Control
2
High Fat
1.5
1
0.5
0
C57 24wks
mdx 24wks
C57 40wks
mdx 40wks
PGC-1 alpha gene expression /L19
0.015
D) PGC-1α
A
B
B
B
B
B
B
B
0.01
Control
High Fat
0.005
0
C57 24wks
Figure 6.
mdx 24wks
C57 40wks
mdx 40wks
Radley-Crabb and Grounds, 2010. 35
Food consumption (g /g bw /day)
0.2
A
A
A
B
B
A
B
0.15
Control
High Fat
0.1
High Protein
0.05
0
Sedentary mdx
Energy intake (kj /g bw /day)
3
A
A
A
Exercised mdx
B
C
B
B
Control
2
High Fat
High Protein
1
0
Sedentary mdx
Figure 7.
Exercised mdx
Radley-Crabb and Grounds, 2010. 36
Final bodyweight (g) 12 wk old mdx
35
A
30
Control
25
20
High Fat
15
High Protein
10
5
0
Sedentary mdx
Exercised mdx
Std Epididymal fat pad (g/g bw)
0.014
0.012
A
A
AB
B
A
B
AB
0.01
Control
0.008
High Fat
0.006
High Protein
0.004
0.002
0
Sedentary mdx
Exercised mdx
C
Std gastroc weight (g/g bw)
0.009
0.008
0.007
Control
0.006
0.005
High Fat
0.004
0.003
High Protein
0.002
0.001
0.000
Sedentary mdx
Figure 8.
Exercised mdx
Radley-Crabb and Grounds, 2010. 37
Total distance run (km) over 4 weeks.
300
A
A
B
A
Control
200
High Fat
High Protein
100
0
8-12 wks old mdx
16
A
A
A
B
B
B
C
Necrosis in quadriceps (% CSA)
14
12
Control
10
High Fat
8
6
High Protein
4
2
0
Sedentary mdx
Figure 9.
Exercised mdx
Radley-Crabb and Grounds, 2010. 38
PPAR alpha gene expression /L19
0.06
A) alpha
A
A
A
B
0.05
0.04
Control
0.03
High Fat
0.02
0.01
0
Sedentary mdx
Exercised mdx
B) gamma
PPAR gamma gene expression /L19
0.0007
A
0.0006
A
A
B
0.0005
Control
0.0004
High Fat
0.0003
0.0002
0.0001
0
Sedentary mdx
Exercised mdx
C) delta/beta
PPAR delta/beta gene expression /L19
8
A
7
A
A
B
6
5
Control
4
High Fat
3
2
1
0
Sedentary mdx
Exercised mdx
PGC-1α gene expression /L19
0.007
0.006
D) PGC-1α
A
A
A
B
0.005
Control
0.004
High Fat
0.003
0.002
0.001
0
Sedentary mdx
Figure 10.
Exercised mdx
Radley-Crabb and Grounds, 2010. 39
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Radley-Crabb et al, 2010. 1
A comparison of metabolism and protein synthesis rates in young and adult
dystrophic mdx and control C57Bl/10 mice.
Hannah G. Radley-Crabb1, Miranda D. Grounds1, Marta L. Fiorotto2.
1. School of Anatomy and Human Biology, the University of Western Australia, Perth,
Australia.
2. USDA/ARS Children’s Nutrition Research Center, Department of Pediatrics, Baylor
College of Medicine, Houston, Texas.
Key words:
Mdx mice, body composition, energy expenditure, protein turnover.
Radley-Crabb et al, 2010. 2
Abstract:
The mdx mouse is a widely used animal model for pre-clinical Duchenne Muscular
Dystrophy (DMD) research. Impaired function or absence of the skeletal muscle
subsarcolemmal protein dystrophin leaves dystrophic myofibres susceptible to damage
during contraction, yet the specific mechanism by which the absence of dystrophin
leads to myofibre necrosis is still unclear. Dystrophic skeletal muscle is also subject to
many metabolic abnormalities that can greatly impact on the maintenance of muscle
mass and function. This research thoroughly investigated the metabolic differences
between young (4 week old) and adult (14 week old) dystrophic mdx and control
(normal) C57Bl/10 mice specifically measuring; energy expenditure, activity level, body
composition and protein synthesis rates. Striking differences were observed in the
metabolic processes between both young and adult dystrophic and control mice.
Young mdx mice havd a ‘stunted’ growth; they were significantly lighter with reduced
fat free mass (lean muscle mass). Young mdx mice also had a significantly increased
protein synthesis rate (in both whole muscle samples and isolated myofibrillar
fraction) and increased metabolic rate (Heat kcals/24hr/kg ffm). The acute onset of
myofibre necrosis and regeneration at 3-4 weeks of age appears to have a significant
effect on many parameters related to muscle mass and function, and metabolic rate in
young mdx mice. There is significant ‘catch-up’ growth in mdx mice between 4 and 14
weeks of age, with 14 week old adult mdx mice being heavier, with significantly
increase muscle mass and bone length than C57Bl/10 mice. The adult mdx mice were
very lean, with significantly increased fat free mass (lean muscle mass) and increased
protein synthesis rates (in both whole muscle samples and isolated myofibrillar
fraction). These findings have many implications for understanding the basic pathology
of DMD especially with respect to susceptibility to myofibre damage of both growing
and mature muscle and impact on potential therapeutic interventions to protect
dystrophic muscle from damage.
Radley-Crabb et al, 2010. 3
Introduction:
Duchenne Muscular Dystrophy (DMD) is an X-linked lethal muscle wasting disorder
affecting approximately 1/3500-6000 male births (1, 2). Patients with DMD exhibit a
progressive loss of muscle mass and function with premature death frequently
occurring due to respiratory and/or cardiac complications (1, 3). There is currently no
cure for DMD and corticosteroids are still the most widely used pharmacological
treatment for patients with the disease (4, 5), however numerous interventions to
correct the gene defect (and replace dystrophin), or other drugs to ameliorate the
severity of dystropathology have been identified over the last 10 years [Reviewed in
(6-9, 10)].
Impaired function or absence of the skeletal muscle subsarcolemmal protein
dystrophin leaves dystrophic myofibres susceptible to damage during mechanical
contraction (11, 12) and this initial membrane damage progresses to myofibre
necrosis. While successful muscle regeneration occurs initially, over time regeneration
is impaired and myofibres are permanently replaced by fatty/fibrotic connective tissue
(2, 13). The primary defect in the dystrophin gene was identified in 1987, although the
specific mechanism(s) leading to myofibre necrosis are still unclear. There are strong
associations with excessive inflammation, increased intracellular calcium levels,
oxidative stress and metabolic abnormality, yet the precise importance of these
aspects requires in vivo elucidation (14-20).
Boys with DMD can move between the spectra of over-nutrition to under-nutrition
within their shortened lifespan. Young DMD patients are often over-weight or obese
(due to a combination of inactivity, steroid therapies and metabolic abnormalities), yet
patients are also often underweight and experience malnutrition in later life [Reviewed
in (20, 21)]. Despite these observations, there is limited literature available on the
precise nutritional, metabolic and energy requirements for DMD patients.
Radley-Crabb et al, 2010. 4
Skeletal muscle is a metabolically active tissue and therefore loss of muscle mass in
DMD patients should lead to a decline in resting energy expenditure. However, DMD is
considered to be a hyper-metabolic condition [Reviewed in (20, 21)] and resting
energy expenditure, when adjusted to muscle mass (fat free mass), is increased in
DMD patients aged 6-13 years (kJ/kg FFM) compared to healthy controls (22, 23).
Similarly, Okada et al (1992) reported an increase in basal metabolic rate (kcal/kg/day)
in DMD patients aged 11-29 years (24).
The confounding effects of different proportions of undamaged mature myofibres,
actively growing myotubes and myofibres (regenerating muscle), and fatty/fibrotic
connective tissue throughout all muscles of individual boys at different ages and stages
of the disease, makes fundamental differences in the metabolism of dystrophic muscle
difficult to evaluate. The problem is further complicated by gross changes in body
composition (obesity vs. malnutrition) and corticosteroid therapies. Thus, an
experimental model, such as the mdx mouse is useful to address the issue of metabolic
status of dystrophic muscle because it enables more invasive measurements to be
performed that can isolate the numerous variables that contribute to the alterations in
whole body measurements.
The mdx mouse (C57BL/10ScSnmdx/mdx) is an animal model for DMD, and has been
extensively described and widely used in pre-clinical experiments to investigate DMD
[Reviewed in (25-28)]. The severity of dystropathology varies throughout the life of
mdx mice. There is a spontaneous onset of acute myofibre necrosis and subsequent
regeneration in limb and paraspinal muscles around 3 weeks of age (29, 30), and
skeletal muscle damage (necrosis and regeneration) peaks around 4 weeks. Muscle
damage then decreases significantly to stabilise by 8-12 weeks of age to a relatively
low level of damage where approximately 6% of the quadriceps muscle is actively
necrotic (14, 26, 31, 32).
Radley-Crabb et al, 2010. 5
The extent and progression of dystropathology is far more pronounced in humans
compared to the relatively small and short-lived mdx mouse (33). The striking
difference in the severity of pathology between mdx mice and DMD patients is widely
acknowledged and may reflect the size, lifespan and especially the duration of the
postnatal growth phase (e.g approximately 6 weeks in mice vs 20 years in humans)
when muscle may be especially susceptible to damage (33). Continuous cycles of
myofibre necrosis and regeneration, regenerating myofibres in various stages of
growth, increased demand on the sarcoplasmic reticulum to regulate intracellular
calcium, defective mitochondrial function and disruption in nNOS signalling may all
contribute to an altered metabolic state in dystrophic muscle (17, 23, 34-39).
Changes in protein metabolism have been reported in various mdx muscles both in
vivo and ex vivo (40-44). The rates of protein synthesis and protein degradation are
elevated in the gastrocnemius muscle of mdx mice (various ages, sex unspecified) and
it is suggested that this process is mediated by an increase in ribosome concentration
in mdx muscles (42). Interestingly, in mdx liver where there is no dystropathology, the
rate of protein synthesis is similar to control C57BL/10 mice, indicating that altered
protein metabolism in mdx mice is restricted to skeletal muscles (42). The extent to
which this reflects the increased protein demand of growing myotubes and myofibres
compared with mature myofibres is unclear in dystrophic muscle.
Adult mdx mice maintain skeletal muscle mass and experience significant skeletal
muscle hypertrophy (27, 45-49), indicating that protein synthesis must outweigh
protein degradation in adult mdx mice. Skeletal muscle hypertrophy is also
pronounced in dystrophic cat models of muscular dystrophy (HFMD) (50, 51). Skeletal
muscle hypertrophy is also a feature of young DMD boys (~ 5 years of age) where
hypertrophic calf muscles are a diagnostic feature (1, 51). This is in striking contrast to
older (6-18 years), wheel chair bound, DMD patients that lose lean muscle mass at an
approximate rate of 4% per year (52). Skeletal muscle hypertrophy in adult mdx mice
may be linked to an increase in AKT signalling (45), since AKT is a key downstream
Radley-Crabb et al, 2010. 6
molecule of IGF-1 signalling that results in increased protein synthesis and hypertrophy
of growing muscles such as that found during the regenerative process in mdx mice
(48, 53). There are also reports of increased circulating growth hormone in 8-10 month
old female mdx (54) and increased circulating IGF-1 levels in 8-10 week old (sex
unspecified) mdx mice (55).
It was hypothesised by Dupont-Vesteegden et al (1994) that continuous cycles of
myofibre necrosis and subsequent regeneration, increased levels of intracellular
calcium and elevated protein synthesis in dystrophic mdx muscles could lead to an
overall increase in whole body metabolic rate. However, they reported no change in
the metabolic rate [kcal/(kg bodyweight0.75. d)] of one year old adult mdx mice (mixed
sex). In contrast, there was a decrease in the whole body metabolic rate [kcal/(kg
bodyweight0.75.d)] of 4-6 week old (mixed sex) mdx mice, and it was proposed that this
was a consequence of reduced physical activity and reduced food consumption (g food
/g body weight) (34). Mokhtarian et al (1996) also reported no difference in total
energy expenditure, basal energy expenditure and spontaneous activity between mdx
and control C57Bl/10 mice (sex unspecified) at 6-12 months of age (49). Clearly, mouse
age, extent of dystropathology and the presence of regenerating (growing) muscle,
may have a major impact on metabolic demand and energy usage.
The purpose of this longitudinal study (over approximately 10 days) was to measure
food intake, energy expenditure, activity level, body composition and protein synthesis
rate (for both whole muscles and the myofibrillar fraction) of dystrophic mdx and
control C57BL/10 mice at 2 different ages; 1) Young mice aged 3-4 weeks (shortly after
the acute onset of muscle necrosis in mdx mice when muscle damage regeneration is
at a peak). 2) Adult mice aged 13-14 weeks (when adult mdx mice exhibit a lower level
of muscle damage and dystropathology).
Radley-Crabb et al, 2010. 7
It was hypothesised that:
A) There would be no difference in absolute daily food intake (g) between strains.
However, due to their higher fat free mass (lean muscle), mdx mice will consume less
food (g), when adjusted to fat free mass. B) Continuous cycles of muscle necrosis and
regeneration in both young and adult mdx mice will promote an increase in whole
body metabolic rate (relative to control mice) and that this difference will be amplified
in young mdx mice when muscle necrosis and regeneration are most pronounced. C)
As hypertrophy is a feature of adult mdx muscle, PIXImus body composition scans will
show a higher total % lean muscle mass and a lower total % fat mass in adult mdx mice
compared to age-matched control mice. However, there will be no marked differences
in body composition between young strains of mice.
Materials and Methods:
Animals
All animal experiments and analysis were carried out at The Children’s Nutrition
Research Center (CNRC) at Baylor College of Medicine, Houston USA. Experiments
were conducted in strict accordance with the U.S. National Research Council’s Guide
for the Care and Use of Laboratory Animals and all procedures were approved by the
Baylor Medical College Animal Care and Use Committee. The young 3 - 4 week old
male mice (mdx and C57Bl/10) were bred at the CNRC from breeders originally from
Jackson Laboratories. Litters were standardised to 7 pups per dam on the day of birth
and mice were changed to a wire bottom cage on the day after weaning (22 days of
age). Only male mice were studied. Adult 13-14 week old male mice (mdx and
C57BL/10) were ordered from Jackson’s laboratory (USA) and changed to a wire
bottom cage on the day after arrival. Mice were left to stabilize for a minimum of 6
days at CNRC prior to experimentation (3 days stabilization was allowed before food
consumption was measured). A typical experimental timeline for both young (4wks)
and adult (14wks) mdx and C57Bl/10 mice is outlined in Table 1. An additional set of 8
(4 C57 and 4 mdx) 14-wk-old mice were used only for calorimetry and activity
Radley-Crabb et al, 2010. 8
measurement, as the complete set of the activity measurements was not obtained on
the first set of mice.
Food consumption. All mice were maintained on a standard diet based on AIN93G
containing 19% protein (Research Diets, New Brunswick, NJ). Daily food intake (g) was
measured before and during measurements of energy expenditure. Food intakes
determined before the energy expenditure measurements were estimated from the
change in the weight of food in the food hoppers corrected for losses. Measurements
made during the energy expenditure determinations utilized the automated in cage
feeding assembly of the CLAMS system (Columbus Instruments)
Calorimetry. Mice were placed individually into calorimetry chambers for 72 hrs and
energy expenditure was measured using a Columbus Instruments (Columbus OH)
CLAMS Oxymax System. A known flow of air was passed through the chambers and the
O2 and CO2 gas fractions were monitored at both inlet and outlet ports. The gas
fraction and flow measurements were used to calculate VO2 and VCO2, from which the
Respiratory Exchange Ratio and heat production (kcal/hr) were calculated. At the same
time spontaneous activity level (both horizontal and vertical movements) was
monitored from the interruption of infra-red beams projected across the cage.
Whole Body Composition Scans. Total body lean muscle and fat mass were measured
by dual energy X-ray absorptiometry (DXA) using a PIXImus (General Electric, USA).
Whilst under isoflurane anaesthesia mice were aligned on a PIXImus measuring tray
and then inserted into the machine for measurements. All mice were measured in
duplicate and then returned to their cage to recover from the anaesthetic, a minimum
of 2 days elapsed before protein turnover measurements were performed. Correction
factors derived specifically for the CNRC PIXImus were applied to the fat and lean mass
data to adjust for the inherent errors in fat and lean mass obtained using all PIXImus
instruments (http://www.bcm.edu/cnrc/intranet/MMRU/PIXImus%20corrections.xls).
Radley-Crabb et al, 2010. 9
Isotope infusion and tissue collection to measure rate of protein synthesis. Food was
removed from all mice 7 hours prior to isotope infusion. A flooding dose of L-4-[3H]phenylalanine (American Radiolabeled Chemicals, Inc., St. Louis, MO) at 20mL/kg BW,
1.5mmol phenylalanine/kg and 250µCi/mouse was given through the tail vein, 15
minutes after injection mice were sacrificed (by decapitation) and trunk blood was
immediately collected and acidified to 0.2M Perchloric acid (PCA). The supernatant
was frozen and reserved for estimation of the specific radioactivity of blood free
phenylalanine pool. The hindlimbs were immediately detached, wrapped in foil and
chilled on ice. A range of muscles including; gastrocnemius and plantaris complex
(referred to hereafter as the gastrocnemius), quadriceps, soleus, tibialis anterior,
diaphragm and heart, were then dissected on ice and frozen in liquid nitrogen. The
weights of all muscles were recorded and muscles were stored at -80ºC.
Laboratory methods: These are based on Fiorotto et al (2000) and Welle et al (1993)
with some modifications (56, 57).
Total Protein, and Myofibrillar Proteins isolation: Frozen muscles from each mouse
were powdered, weighed and 50mg was homogenized in a low salt homogenising
buffer (50 mM potassium phosphate, 0.25 M sucrose, 1% Triton X-100 pH 7). For each
muscle, an aliquot of the homogenate was retained for measurement of total protein;
a second aliquot was acidified to 0.2M PCA and centrifuged; the supernatant
containing the muscle free amino acid precursor pool was collected and neutralized as
described below. The PCA-insoluble precipitate was washed, and after assaying for
total RNA, was processed for the measurement of L-[4-3H] phenylalanine incorporated
into all muscle proteins. The remainder of the low salt/sucrose buffer homogenate
was left to incubate with agitation at 4°C for 60 minutes; the proteins that were
insoluble after low speed centrifugation (1,500 g at 4ºC for 10min) contained the
myofibrillar proteins (56, 57). After several washes of the pellet in the low salt/sucrose
buffer followed by ice-cold water, the purified myofibrils were acidified to 0.2M PCA.
The acid insoluble myofibrillar and total protein precipitates were separated from the
supernatant by centrifugation at 10,000 g at 2°C for 30 min, and the pellets were
Radley-Crabb et al, 2010. 10
washed three times in 0.2M PCA. The supernatant from the total protein fraction and
the pellets from all protein fractions were processed for determination of L-[4-3H]
phenylalanine specific radioactivity.
Quantification of L-[4-3H] phenylalanine specific radioactivity in the protein bound
and tissue free amino acid precursor pools: The acid-insoluble pellets were first
hydrolysed for 24 hrs in 6M HCL (ultrapure) at 110oC under N2 gas. After evaporation
of the HCl in a Savant Speedvac concentrator, the remaining precipitate was subject to
several washes in water and finally resuspended in mQ water for HPLC analysis. The
PCA-treated blood and muscle free pool supernatants samples were neutralized on ice
with 4M KOH, evaporated to dryness and resuspended in 1M acetic acid. The amino
acids were purified over a Dowex anion exchange column (AG-50w-x8, 100 - 200 mesh,
Biorad). The amino acids were eluted in 3M NH4OH evaporated to dryness in the
Savant Speedvac concentrator and finally dissolve in mQ water. All samples were
filtered using a 0.45µm syringe filter, supernatant dried down and each sample was redissolved in 100ul of mQ water. Phenylalanine in the total and myofibrillar protein
hydrolysates, muscle homogenate and blood supernatants were isolated by anion
exchange HPLC (AminoPac1 Analytical column; Dionex, Sunnyvale, CA). Amino acids
were post-column derivatized with o-phthalaldehyde reagent and detected with an online fluorimeter. The fraction of the eluant that included the phenylalanine peak was
collected, and the associated radioactivity was measured in a liquid scintillation
counter (Packard Tricarb, Perkin Elmer, Waltham, MA). The phenylalanine
concentration was determined by comparing the peak areas of the samples with that
of a known standard (Pierce Labs.)
Calculations of in vivo fractional protein synthesis rates: The fractional rate of
protein synthesis (FSR), i.e., the percentage of the protein mass synthesized in a day
was calculated as follows: FSR = (SB/ SA )  (1440/t) 100
Where SB is the specific radioactivity of the protein-bound labelled amino acid, SA is the
specific radioactivity of the precursor pool and t is the the labelling time in minutes
(58). It has been demonstrated that the specific radioactivity of the tissue free
phenylalanine after a flooding dose of phenylalanine is in equilibrium with the
Radley-Crabb et al, 2010. 11
aminoacyl-tRNA specific radioactivity; hence, the tissue free phenylalanine reflects the
specific radioactivity of the tissue precursor pool (59). Blood and tissue free
phenylalanine specific radioactivity values were compared to verify equilibration.
Protein quantification: Total protein concentration of all muscle samples was
determined using a standard BCA protein assay (60) after first solubilising the aliquot
of total muscle homogenate in 0.1 M NaOH for 1 hr at 37 oC.
Total RNA measurement: Because the RNA content of a tissue is dominated by
ribosomal RNA, total RNA was measured quantitatively on all muscle samples and
provides an estimate of ribosomal abundance. Total RNA was quantified in the total
protein PCA-insoluble precipitate using a modified Schmidt-Thannhauser procedure as
described by Munro and Fleck (1966) (61).
Statistical analysis:
Statistical analysis was completed by using Microsoft excel and SPSS software. Values
throughout the manuscript are expressed as mean +/- standard error of measurement
(SEM). Significant difference between 2 groups was determined by conducting
student’s T-test. To establish the extent to which differences in the variables related
to energy balance might be accounted for by differences in body weight or fat free
mass (FFM), ANOVA was performed, with strain as the main independent effect and
body weight or FFM as covariates. Results are presented as least square means +/- the
standard error.
Results - Young mice:
Body composition: The young dystrophic mdx mice weighed significantly less
(approximately 25%) than age-matched C57Bl/10 mice (16.5 g +/- 0.3 vs. 20.7 g +/- 0.4).
This difference in body weight is due to a significant and proportional reduction in both
fat free mass (FFM) (e.g, lean muscle tissue) and fat mass as measured by PIXImus
(Table 2). Bone mineral content (BMC) was also significantly reduced in dystrophic
mdx mice. The reduction in BMC was attributable in part to a reduction in total bone
area (5.56 +/- 0.10 cm2 vs. 6.55 +/- 0.06 cm2, P< 0.001), but also to a reduction in bone
Radley-Crabb et al, 2010. 12
mineralization as indicated by the bone mineral density (Table 2). In addition to the
PIXImus body scans, individual muscle weights (g) and bone lengths (mm) recorded at
time of animal sacrifice were significantly lower for the gastrocnemius and tibialis
anterior muscles and there was also a significantly shorter length of femur and tibia
bone in young mdx mice (Table 2). After adjusting for these differences in bone length,
only the gastrocnemius muscle remained significantly smaller in the mdx mice
(P<0.05). Absolute heart and diaphragm weights were similar between young mdx and
C57Bl/10 mice, however when adjusted to FFM (an indicator of the body’s active
metabolic mass) there was a significant increase in both heart and diaphragm weights
of young mdx mice (Table 2).
Calorimetry and activity level:
As the same diet was consumed by both strains at all ages, differences in nutrient
intakes are entirely explained by differences in daily food intake. Energy intake (kcal/d)
was significantly lower in young mdx mice compared to C57Bl/10 mice (10.3 +/- 0.5 vs
11.3+/- 0.3) (Table 3). However, when adjusted to individual bodyweight and/or fat free
mass (FFM), young mdx mice consume significantly more energy than C57Bl/10 mice
(Table 3). Respiratory exchange ratio (RER – ratio of carbon dioxide production /
oxygen production) was similar between strains suggesting both strains are using the
same type of fuel over a 24hr period. Total energy expenditure (heat production kcal/d) was unchanged between strains; however, when adjusted for difference in
FFM, energy expenditure was significantly increased in young mdx mice (Table 3). Daily
energy balance, calculated as the difference between energy intake and energy
expenditure was significantly less positive in mdx mice. After adjusting for differences
in body weight or FFM, the values remained numerically smaller but were no longer
significant. Total daily activity was reduced by approximately 35% in young mdx mice
(Table 3), especially at night (during their active phase). The young mdx mice
performed significantly less vertical movements (standing on back legs) during the day
(819 +/- 71 vs 1683 +/- 294) and during night (3812+/- 625 vs 17093 +/- 2389) compared
Radley-Crabb et al, 2010. 13
to young C57Bl/10 mice. The mdx mice also performed significantly less horizontal
movement during the night (33486 +/- 1887 vs 46797 +/- 3490) (Table 3).
Protein synthesis: We initially compared fractional synthesis rates (FSR) for total and
myofibrillar proteins in the quadriceps, gastrocnemius and diaphragm muscles in a
subset of mice. The values were quite similar for the gastrocnemius and quadriceps,
and thus, for the full set of mice we performed measurements only on the
gastrocnemius. Additionally, we evaluated the response of the diaphragm to establish
if the differences in the pathophysiology of this muscle in the mdx mouse are
manifested in protein synthesis rates. FSR of both total and myofibrillar proteins were
greatly increased in both gastrocnemius and diaphragm muscles of young mdx mice
(Table 4). Total proteins FSRs were more than 2 fold higher in young mdx mice than in
controls, and there was no difference between muscles. The myofibrillar protein FSR
was approximately 2.5 fold higher in the gastrocnemius muscle of the young mdx mice,
whereas for the diaphragm the increase was only approximately 2 fold; there was no
difference between the FSR of myofibrillar proteins in diaphragm and gastrocnemius
muscles of the control mice (strain X muscle, P=0.02) (Table 4). Absolute rates of
synthesis for total proteins (mg/d) reflected the differences between strains in FSR
(mdx>C57, P<0.001). Absolute rates of myofibrillar protein synthesis could not be
calculated as the isolation of myofibrillar proteins was not quantitative.
Although the FSR values were similar for the two muscles in both strains of mice, this
was achieved by slightly different mechanisms. The increase in gastrocnemius FSR in
the young mdx mice was due, in part, to an 80% increase in ribosome abundance
(RNA/TP – mg/g) together with a 40% increase in their translational efficiency (KRNA –
mg/g) (Table 4). The ribosomal abundance in the diaphragm was higher overall than in
gastrocnemius muscle for both strains of mice, and the difference in ribosomal
abundance between mdx and C57Bl/10 in the diaphragm was approximately 30%. Yet,
the overall translational efficiency of the diaphragm ribosomes was significantly lower
than for the gastrocnemius ribosomes in mdx mice and the difference between
Radley-Crabb et al, 2010. 14
C57Bl/10 and mdx diaphragm ribosomes was significantly greater (approximately 60%)
than for the gastrocnemius muscle.
Despite the 80% increase in total gastrocnemius muscle protein synthesis rate (mg/d)
of the mdx mice, total protein content (mg) of the gastrocnemius muscle was
decreased by approximately 4mg or 40% in young mdx mice (Table 4), and this can
only be explained by a substantial increase in protein degradation. Myofibrillar FSR is
increased in both the gastrocnemius and diaphragm muscle in mdx mice; however the
difference between strains is much greater in the gastrocnemius.
The strain difference in total muscle protein synthesis rate (mg/d) was greater in the
diaphragm (increased in mdx approximately 100%) than the gastrocnemius muscle
(increased in mdx approximately 85%), yet there was no change in total muscle protein
(mg) in the diaphragm muscle between strains (Table 4). Thus, protein degradation
rate in the diaphragm must be substantially higher in the mdx mice compared to
C57Bl/10, but probably not to the same extent as in the gastrocnemius muscle, as the
mdx diaphragm is able to maintain protein mass (mg). As the strain difference in
myofibrillar FSR in the diaphragm was not as great as in the gastrocnemius muscle, one
interpretation is that the replacement of nonmyofibrillar proteins makes a greater
contribution in the diaphragm of mdx mice either because the protein composition is
different or because the contractile proteins are less impacted by dystrophy than in
the gastrocnemius muscle.
Results - Adult mice:
Body composition: There was no difference in body weight of adult mdx mice and agematched C57BL/10 mice (Table 5). PIXImus body composition scans revealed that mdx
mice had significantly more FFM (approximately 15%); this increase largely reflects
increased lean skeletal muscle but also includes some other tissues such as gut and
liver. Mdx mice also had a huge reduction (approximately 50%) in fat mass which,
together with the increase in FFM, was manifested as a 45% reduction in percentage
Radley-Crabb et al, 2010. 15
body fat. In addition to the PIXImus body scans, individual muscle weights and bone
lengths (recorded at time of animal sacrifice) showed significantly heavier
gastrocnemius, quadriceps, soleus, tibialis anterior, and diaphragm muscles in mdx
mice (Table 5) and also significantly longer femur and tibia bone length (Table 5).
Although there was no difference in total bone mineral content (g), given the larger
bone area, bone mineral density (g/cm2) was significantly lower in the mdx mice.
Absolute heart weight (g) was similar between adult mdx and C57Bl/10 mice; however
when adjusted for FFM, there was a significant decrease in the heart weight of mdx
mice (g/ kg FFM) (Table 5).
Calorimetry and activity level:
There were no differences in energy intake (kcal/d), energy expenditure (heat
production – kcal/d) or energy balance (both absolute or when adjusted for body
weight or FFM) between strains (Table 6). RER was similar between strains suggesting
both were using the same fuel over a 24hr period. Total activity level was significantly
lower in the adult mdx mice, and was attributable primarily to a reduction in vertical
movements (stand on back legs) at night (during their active phase) compared to adult
C57Bl/10 mice (Table 6).
Protein synthesis:
Fractional synthesis rates (FSR) of both total and myofibrillar proteins in both the
gastrocnemius and diaphragm muscles exhibited the normal developmental decline in
both strains of mice. FSRs of the gastrocnemius muscle in adult mdx mice were
significantly higher than age-matched C57Bl/10 mice; total protein FSR was
approximately 2.5x fold higher and myofibrillar protein FSR was approximately 2x fold
higher in adult mdx mice (Table 7); the age-related decline in myofibrillar FSR was
significantly greater for the mdx mice than controls. Total protein synthesis (mg/day)
and total muscle protein content (mg) in the gastrocnemius muscle were also
significantly increased in adult mdx mice: total protein synthesis rate was
approximately 3x fold higher in adult mdx and total protein content of the
Radley-Crabb et al, 2010. 16
gastrocnemius muscle was increased by approximately 4 mg or 20% (Table 7). These
increases in FSR were due to the combination of an increase in ribosome abundance
(40% greater) and ribosomal translational efficiency (70% greater) in adult mdx mice
(Tables 7).
A quantitative dissection of the diaphragm in the adult mice was achieved for only 4
mice of each strain. All other parameters evaluated did not differ between this sub
group of mice and the total group. Thus, the average trends over time and differences
between strains and muscles are likely to be valid reflections for the diaphragm
muscle. The developmental decline in FSR for the diaphragm was less marked than for
the gastrocnemius muscle proteins, and was more pronounced in the mdx. Total
protein FSR was approximately 60% higher and myofibrillar protein FSR was
approximately 50% higher in the diaphragm of adult mdx compared to C57Bl/10 mice
(Table 7). Total daily protein synthesis rates (mg/d) and total diaphragm protein mass
(mg) were significantly greater in mdx than C57Bl/10 mice; this difference in protein
mass was more pronounced in the diaphragm than the gastrocnemius muscle. In
contrast to the gastrocnemius muscle, the difference in ribosomal abundance between
strains (45% greater in mdx) made a greater contribution than differences in ribosomal
efficiency (20% greater in mdx).
Percentage change from 4-14 weeks of age:
Despite being significantly (approximately 4g) lighter at 4 weeks of age, mdx mice were
similar in weight to the C57Bl/10 mice at 14 weeks of age, representing a 176%
increase in bodyweight over 10 weeks, compared to a 142% increase for C57Bl/10
mice. Over this 10 week period, mdx mice showed a large increase in FFM
(approximately 12.5 g) and deposited very little fat mass (approximately 0.2 g); this
was in stark contrast to C57Bl/10 mice that gained approximately 5.9 g of FFM and
deposit 2.8 g of fat over the 10 weeks (Table 8). The percentage growth of all
individual muscles (except heart) over this time was also much higher in mdx
compared to C57Bl/10 mice; for example, there was a 240% increase in weight in the
Radley-Crabb et al, 2010. 17
mdx gastrocnemius compared to only a 152% increase in the C57Bl/10 gastrocnemius
(Table 8). The lack of fat deposition suggests that mdx mice, in contrast to the
C57Bl/10 controls, were not consuming energy in excess of their needs during this
time, and that a greater portion of dietary protein was partitioned toward protein
deposition rather than being oxidized. In addition to rapid muscle growth, bone length
also caught up and overtook the controls during these 10 weeks.
Discussion:
Young mice:
Young mdx mice (aged 3-4 weeks) were significantly smaller (by approximately 25%)
than age-matched C57Bl/10 mice due to reductions in both fat free mass (lean muscle
tissue), fat mass, and individual muscle weights, with bone length and bone mineral
content also significantly reduced in young mdx mice (Table 2). Since bone strength
(and thus mineral content) is determined by the dynamic load from skeletal muscles
this smaller bone size may directly result from the reduced muscle mass. In addition,
15 day old dystrophic mdx mice are significantly lighter than aged matched C57Bl/10
mice (Mdx 7.53g +/- 0.18 vs. C57 8.24g +/- 0.16) (Radley-Crabb 2010 unpublished) and it
is also known that the dystropathology of mdx mice begins in early development (62).
These combined data indicate that growth of young mdx mice is decreased (stunted)
during development, and this occurs well before the onset of severe pathology
(around 3 weeks of age).
Multiple factors could contribute to the reduced growth of young mdx mice. The rate
of protein synthesis (both total protein and myofibrillar proteins) is more than doubled
(2.5 fold higher) in young (growing) mdx mice compared to C57Bl/10 control mice
(Table 4) and protein synthesis is a very energetically expensive process [Reviewed in
(63)] that, if not supported by adequate increases in food intake, may occur at the
expense of overall growth. After accounting for the differences in size, young mdx mice
(26-28 days) have an increased metabolic rate which supports the increased energy
demands of a high protein synthesis rate and the increased energy demands of
Radley-Crabb et al, 2010. 18
myofibre regeneration and myofibre growth due to the acute onset of dystropatholgoy
at this time. However, this result disagrees with a previous study that measured
metabolic rate in 4-6 week old mdx mice (34). Nonetheless, it should be noted, that
our data indicating greater energy expenditure are supported by the significantly
increased heart weight, even after adjusting for differences in FFM, and cardiac
hypertrophy is a classic functional response to an increase in the oxygen demand of
body tissues.
It appears that the increased energy needs of young mdx mice were met partly by a
relative increase in food intake (after taking into account differences in body size) and
partly by expending less energy by being less active, especially at night (Table 3). This
possibly reflects the acute onset of pathology around this age; perhaps this may be an
attempt to conserve energy at this time of high muscle damage or the mdx mice may
feel ‘unwell’. It must also be considered that mdx mothers (who are energy deficient
themselves) may not produce sufficient milk for their pups.
It seems likely that the cycles of myofibre necrosis and regeneration (which have an
acute onset around 3 weeks of age and peak around 28 days in mdx mice) are
responsible (entirely or in part) for the increase in protein fractional synthesis rate in
young mdx mice (e.g. due to the acute inflammation, proliferation of myoblasts and
rapidly growing myotubes in the regenerating muscle). The large difference between
protein synthesis rates in whole mdx muscle (Total protein FSR) and the isolated
sarcomeric myofibrillar fraction (myofibrillar protein FSR) is probably due to the many
inflammatory cells present in necrotic/regenerating dystrophic muscles at this young
age (14, 26), since the inflammatory cells inherently have higher protein synthesis
rates than myofibrillar proteins.
It remains unknown if ‘intact’ dystrophic skeletal myofibres have an inherent increase
in protein synthesis rate or if the elevated protein synthesis is a result of continuous
cycles of myofibre necrosis and regeneration. However, the delayed growth of young
Radley-Crabb et al, 2010. 19
mdx mice,that must occur in the pre-necrotic neonatal phase strongly supports the
proposal that there are inherent differences in the metabolic rate of ‘intact’ dystrophic
myofibres. This issue might be addressed, in part, by similar studies conducted in prenecrotic mdx mice, i.e. <20day old, that have either been cross-fostered onto normal
non-dystrophic mothers, to avoid complications of reduced milk quality or quantity by
mdx mothers, or born to heterozygous mdx mothers to mimic the human situation.
Adult mice:
Adult male mdx mice (14 wks) attained the same weight as C57Bl/10 mice due to a
significant increase in fat free mass (lean muscle tissue) and significant hypertrophy of
many individual muscles (Table 5). Adult mdx mice are very lean and have very little
fat mass (approximately 50% less than C57Bl/10 mice) and a significantly reduced
percentage body fat (Table 5). The same differences in body composition are seen in
40 week old mdx and C57Bl/10 mice (Radley-Crabb et al, 2010 unpublished). The body
composition of adult mdx mice is in stark contrast to DMD patients who may
experience obesity (due to a combination of inactivity and steroid therapies) and
extreme loss of muscle mass and function as the disease progresses during the growth
phase.
The increased protein synthesis rate and increased ribosomal efficiency in adult mdx
mice (Table 7) facilitates skeletal muscle hypertrophy. Previous studies also report an
increased protein synthesis rate (approximately 2x fold) in the gastrocnemius, soleus
and EDL muscles from adult mdx mice (40, 42, 44). The present data show that the
myofibrillar protein synthesis rate of mdx mice is about twice as high as C57Bl/10 mice
in both adult and young mice. This suggests that the increased myofibrillar protein
synthesis rate in mdx mice may reflect inherent differences in dystrophic myofibres (at
all ages) rather than merely the consequence of necrosis (with associated
inflammation) and regeneration since myofibre necrosis is far less pronounce in adult
compared with young mdx mice. It should be noted that all the measurements in this
study were performed with the mice in the fasted condition, and it is difficult to infer if
Radley-Crabb et al, 2010. 20
these differences are maintained, or are greater or smaller in the postprandial
condition.
Because ribosomal abundance is increased, it is likely that in the
postprandial condition FSR values would remain higher in the mdx muscle than in the
controls, even if translational efficiency were to increase more in the control muscles.
The ability of the mdx muscle to sustain higher translational efficiency even in the
fasted state is unusual, as translational efficiency is highly dependent on the activation
of the insulin and mTOR signalling pathways (64), and these are usually inactive in the
fasted state. However, these pathways can be activated by stimulation of the type
IGF1-R, and both IGF-I and IGF-II expression are relatively high in the early stages of
muscle regeneration in young mdx mouse muscles (65, 66).
Overall energy expenditure was not significantly different between adult mdx and C57
mice even after adjusting for the differences in FFM. There also was no difference in
energy intake, so that overall there was no difference in energy balance at this age.
The adult mdx mice, however, were significantly less active than controls (Table 6)
primarily because they reduced vertical movements (rear-up on back legs) at night.
This result agrees with previous studies which document significantly reduced general
activity levels in mdx mice (27, 67). The reduction in energy expenditure for activity
likely compensated for the greater energy demands of maintaining their higher
metabolic activity. This could be demonstrated by performing measurements of
resting energy expenditure when there would be no contribution from activity and the
thermal effect of food on total energy expenditure.
It appears, therefore, that adult mdx mice focus their energy on the maintenance of
skeletal muscle mass and growth of new myofibres (that leads to hypertrophy) and
because they do not increase food intake, they do not have have any excess energy to
deposit as adipose tissue. In support of the importance of this vulnerable metabolic
/energy balance, 48 hours fasting causes a significant increase in myofibre necrosis in
muscles from the hind limb and lumbar region of 6 month old mdx mice, yet no change
Radley-Crabb et al, 2010. 21
in the muscle morphology of control C57BL/10 mice, suggesting a strong dependence
on energy intake to maintain dystrophic muscle structure (44).
These findings have many implications for understanding the basic pathology of DMD
especially with respect to susceptibility to myofibre damage of both growing and
mature muscle and the design and impact of potential nutritional therapeutic
interventions to protect dystrophic muscle from damage.
Acknowledgements:
This collaborative research was made possible by the following funding bodies:
Hannah Radley-Crabb (Australian postgraduate award, UWA PSA & UWA GRST), and
Miranda Grounds (NHMRC & MHRIF). The authors thank the following people at the
CNRC for technical assistance – Firoz Vohra, Deborah Karson, Horacio Sosa, and Sona
Buddhiraja.
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Table Captions:
Table 1) Example experimental Timeline for both young (4 wks) and adult (14 wks)
mdx and C57Bl/10 mice. *Actual sacrifice age of young mice was between 31-32 days
of age and for adult mice was between 90 – 100 days of age.
Table 2) Body composition parameters, individual muscle weights and bone lengths
in young (4 wks) male mice. n=12 for mdx and n=11 for C57Bl/10 mice for body
composition parameters and n=8 for individual muscle weights and bone lengths.
Significant differences are shaded in grey.
Table 3) Food intake, energy expenditure and activity levels in young (4 wks) male
mice. n=8 for mdx and n=8 for C57Bl/10 mice. Significant differences are shaded in
grey. § Values are least square means adjusted for differences in body weight. ¶ Values
are least square means adjusted for differences in FFM. * P=0.01 for strain x bw,
P=0.003 for bw; P=0.04 for strain x FFM; P=0.007 for FFM.
Table 4) Protein synthesis measures in gastrocnemius and diaphragm muscles of
young (4 wks) male mice. n=8 for both mdx C57Bl/10 mice. Significant differences are
shaded in grey.
Table 5) Body composition parameters, individual muscle weights and bone lengths
in adult (14 wks) male mice. n=16-20 for mdx and n=15 for C57Bl/10 mice for body
composition parameters and n=8 for individual muscle weights and bone lengths; *
n=4 per strain for diaphragm. Significant differences are shaded in grey.
Table 6) Food intake, energy expenditure and activity levels in adult (12-14-wk-old)
male mice. n=16-20 for mdx and n=15 for C57Bl/10 mice. Significant differences are
shaded in grey. * for all activity measurements: n=10 for mdx, n=8 for C57Bl/10. §
Values are least square means adjusted for differences in body weight. ¶ Values are
least square means adjusted for differences in FFM.
Table 7) Protein synthesis measures in gastrocnemius and diaphragm muscles of
adult (14 wks) male mice. n=8 for both mdx and C57Bl/10 mice. *indicates n=4 per
strain.
Table 8) Percentage increase in body composition parameters (comparison of group
averages shown in Table 2 & 5), individual muscle weights and bone lengths in mdx
and C57Bl/10 mice over 10 weeks (4-14 weeks of age). Calculated as:
(14wks/4wks)x100. n=16-20 for mdx and n=15 for C57Bl/10 mice for body composition
parameters and n=8 for individual muscle weights and bone lengths. Important
differences are highlighted in grey.
Radley-Crabb et al, 2010. 29
1) Young mdx and C57Bl/10 male mice
(4wks)
Age (Days) Activity/Analysis
1
21
Standardise litters to 7 pups
Wean & separate
22 – 25
Stabilize in wire bottom cage
(food consumption measured)
Calorimetry
Calorimetry
Calorimetry & PIXImus
Stabilize in wire bottom cage
(food consumption measured)
Protein turnover and sampling
26
27
28
29
31*
2) Adult mdx and C57Bl/10 male mice(14
wks)
Age
Activity /Analysis
(Days)
80
Arrive at CNRC
81 - 85
Stabilize in wire bottom cage
(food consumption measured)
86
Calorimetry
87
88
89
Calorimetry
Calorimetry & PIXImus
Stabilize in wire bottom cages
(food consumption measured)
92 *
Protein turnover and sampling
Bone
length
Muscle weight
Body composition
Table 1.
Strain/Parameter
Weight (g)
Bone Mineral Content (g)
Bone Mineral Density (g/cm2)
Fat free mass (g)
Fat tissue (g)
Body fat (%)
Gastrocnemius (g)
Quadriceps (g)
Soleus (g)
Tibialis anterior (g)
Diaphragm (g)
Diaphragm (g/kg ffm)
Heart (g)
Heart (g / kg ffm)
Femur (mm)
mdx
16.5
0. 18
0.032
14.56
1.96
11.8
0.085
0.122
0.007
0.030
0.054
3.71
0.089
6.08
12.66
Tibia (mm)
15.54
Table 2.
0.057
C57Bl/10
20.7
0.23
0.034
18.39
2.30
11.1
0.10
0.127
0.008
0.032
0.052
2.87
0.086
4.71
13.03
0.029
15.99
+/- SE
0.3
0.004
0.001
0.28
0.16
0.9
0.003
0.003
0.0002
0.0006
0.003
0.19
0.005
0.325
+/- SE
P for Strain effects
0.40
0.079
P<0.001
P<0.001
P<0.005
P<0.001
P<0.05
NS (P=0.63)
P<0.001
NS (P=0.1)
NS (P=0.07)
P=0.03
NS (P=0.7)
P=0.002
NS (P=0.7)
P=0.002
P<0.001
0.049
P<0.001
0.01
0.0004
0.31
0.09
0.3
0.002
0.002
0.0001
0.001
0.002
0.11
0.003
0.16
Radley-Crabb et al, 2010. 30
Strain/Parameter
mdx
+/-SE
Energy intake (kcal/d)
Energy intake (kcal/d/bw) §
Energy intake (kcal/d/ffm) ¶
Heat (kcal/d)
Heat (kcal/d/bw) §
Heat (kcal/d/ffm) ¶
RER (24 hr avg)
Energy balance (kcal/d)
Energy balance (kcal/d/bw) §
Energy balance (kcal/d/ffm) ¶
Total Activity (total cts/d)
Horizontal Activity (cts/d)
Verical Activity (cts/d)
Horizontal Activity – Day (cts/12hr)
Verical Activity – Day (cts/12hr)
Total Activity – Day (cts/12hr)
Horizontal Activity – Night (cts/12hr)
Verical Activity - Night (cts/12hr)
Total Activity – Night (cts/12hr)
10.3
12.5
12.0
9.16
10.03
10.20
0.907
1.11
1.03
1.12
52315
47685
4630
14199
819
15017
33486
3812
37298
0.49
0.6
0.5
0.33
0.34
0.31
0.01
0.20
0.39
21.9
2429
1854
629
617
71
671
1887
625
2416
C57Bl/10 +/-SE
0.3
11.3
0.4
11.1
0.5
11.0
0.12
9.59
0.34
8.71
0.31
8.55
0.003
0.916
0.15
1.71
0.39
1.79
9.4
1.70
5525
80167
3827
61393
2425
18776
921
14596
294
1683
1173
16290
3490
46797
2389
17093
5306
63889
P for Strain effects
P= 0.01
P= 0.01*
P= 0.05*
NS (P=0.18)
NS (P=0.057)
P=0.015
NS (P=0.42)
P=0.04
NS (P=0.32)
NS (P=0.44)
P<0.001
P=0.006
P<0.001
NS (P=0.73)
P=0.01
NS (P=0.36)
P=0.005
P<0.001
P<0.001
Table 3.
Strain/Parameter
Gastrocnemius
Total Protein FSR (% / d)
Myofibrillar proteins FSR (% / d)
Total protein synthesis (mg/d)
Ribosome abundance (RNA/TP - mg/g)
Ribosome efficiency (KRNA - mg/g)
Total muscle protein (mg)
Diaphragm
Total Protein FSR (% / d)
Myofibrillar proteins FSR (% / d)
Total protein synthesis (mg/d)
Ribosome abundance (RNA/TP - mg/g)
Ribosome efficiency (KRNA - mg/g)
Total muscle protein (mg)
Table 4.
mdx
sterr +/-
C57Bl/10
sterr +/-
P for Strain effects
31.13
24.67
3.46
18.28
17.16
10.93
2.22
12.31
10.27
1.86
10.24
12.16
15.07
0.57
P<0.001
P<0.001
P<0.001
P<0.001
P<0.001
P<0.001
28.23
20.16
2.12
25.01
11.35
7.35
1.81
13.91
11.55
1.05
19.62
7.13
7.49
0.90
1.54
0.20
0.95
1.23
0.42
1.44
0.24
1.30
0.58
0.37
0.59
0.11
0.46
0.70
0.33
0.82
0.09
1.19
0.31
.39
P<0.001
P<0.001
P<0.01
P<0.01
P<0.01
P<0.05
Bone
length
Muscle weights
Body composition
Radley-Crabb et al, 2010. 31
Strain/Parameter
Weight (g)
Bone Mineral Content (g)
Bone Mineral Density (g/cm2)
Fat free mass (g)
Fat tissue (g)
Body fat (%)
Gastrocnemius (g)
Quadriceps (g)
Soleus (g)
Tibialis anterior (g)
Diaphragm (g)*
Diaphragm (g/kg FFM)*
Heart (g)
Heart (g / kg ffm)
Femur (mm)
mdx
29.18
0.47
0.049
27.01
2.17
7.39
0.203
0.317
0.0127
0.071
0.113
3.96
0.119
4.32
16.06
sterr +/-
sterr +/-
P for Strain effects
0.80
0.06
C57Bl/10
29.37
0.47
0.051
24.28
5.10
16.89
0.152
0.212
0.010
0.044
0.077
3.41
0.113
4.85
15.65
0.10
NS (P=0.81)
NS (P=0.56)
P<0.01
P<0.001
P<0.001
P<0.001
P<0.001
P<0.001
P<0.001
P<0.001
P<0.001
P<0.05
NS (P=0.26)
P=0.01
P=0.02
Tibia (mm)
18.15
0.06
17.91
0.08
P=0.03
0.39
0.006
0.0003
0.33
0.13
0.38
0.007
0.016
0.001
0.004
0.005
0.20
0.005
0.17
0.01
0.001
0.43
0.56
1.49
0.002
0.005
0.0003
0.0004
0.001
0.05
0.002
0.04
Table 5.
Strain/Parameter
mdx
+/-SE
Energy intake (kcal/d)
Energy intake (kcal/ d) §
Energy intake (kcal/d) ¶
Heat (kcal/d)
Heat (kcal/d) §
Heat (kcal/d) ¶
Energy balance (kcal/d)
Energy balance (kcal/d) §
Energy balance (kcal/d) ¶
RER
Horizontal Activity (cts/d)*
Vertical Activity (cts/d)
Total Activity (total cts/d)
Horizontal Activity – Day (cts/12hr)
Vertical Activity – Day (cts/12hr)
Total Activity – Day (cts/12hr)
Horizontal Activity – Night (cts/12hr)
Vertical Activity - Night (cts/12hr)
Total Activity – Night (cts/12hr)
11.08
12.8
12.3
11.08
11.08
10.84
1.75
1.76
1.47
0.927
54432
10571
62002
17539
2405
20943
38263
7679
45939
0.49
Table 6.
0.5
0.5
0.21
0.19
0.19
0.38
0.35
0.37
0.01
3196
1266
3518
1008
296
1418
2869
1066
3737
C57Bl/10 +/-SE
0.3
10.74
0.5
12.3
0.5
12.9
0.16
10.74
0.22
10.73
0.22
11.05
0.34
1.58
0.38
1.57
0.37
1.91
0.02
0.946
4938
59416
4869
24761
9207
84176
846
14611
764
2986
1335
17597
4076
46059
3480
20195
7231
66254
P for Strain effects
NS (P=0.241)
NS
NS
NS (P=0.24)
NS
NS
NS
NS
NS
NS (P=0.33)
NS
P<0.01
P<0.05
P<0.05
NS
NS (P=0.11)
NS (P=0.13)
P<0.01
P<0.02
Radley-Crabb et al, 2010. 32
Strain/Parameter
mdx
sterr +/-
C57Bl/10
sterr +/-
P for Strain effects
9.75
6.74
2.86
7.58
12.82
29.0
0.66
3.82
3.12
0.94
5.31
7.10
24.8
0.44
P<0.001
P=0.001
P<0.001
P<0.001
P<0.001
P=0.02
15.81
10.65
2.60
18.64
9.00
15.53
0.57
Gastrocnemius
Total protein FSR (% / d)
Myofibrillar proteins FSR (% / d)
Total protein synthesis (mg/d)
Ribosome content (RNA/TP - mg/g)
Ribosome efficiency (KRNA - mg/g)
Total muscle protein (mg)
Diaphragm
Total protein FSR (% / d)
Myofibrillar proteins FSR (% / d)
Total protein synthesis (mg/d) *
Ribosome content (RNA/TP – mg/g)*
Ribosome efficiency (KRNA - mg/g)*
Total muscle protein (mg)*
0.76
0.28
0.29
0.59
1.34
0.60
0.10
0.59
0.24
0.53
Table 7.
Strain/Parameter
Weight (g)
Bone Mineral Content (%)
Fat free mass (g)
Fat mass (g)
% body fat
Gastrocnemius (g)
Quadriceps (g)
Soleus (g)
Tibialis anterior (g)
Diaphragm (g)
Heart (g)
Femur (mm)
Tibia (mm)
Table 8.
mdx
176
261
185
C57Bl/10
110
221
152
62
240
259
180
236
209
133
127
117
142
204
132
152
166
125
137
148
131
120
112
9.66
6.94
1.04
12.92
7.27
11.2
0.40
0.097
0.25
0.65
0.7
0.65
0.47
0.07
0.89
0.68
0.2
P<0.001
P=0.001
P<0.001
P=0.002
P=0.05
P<0.001
ARTICLE IN PRESS
Available online at www.sciencedirect.com
Neuromuscular Disorders xxx (2010) xxx–xxx
www.elsevier.com/locate/nmd
Blockade of TNF in vivo using cV1q antibody reduces
contractile dysfunction of skeletal muscle in response to
eccentric exercise in dystrophic mdx and normal mice
A.T. Piers a,b, T. Lavin a, H.G. Radley-Crabb b, A.J. Bakker a, M.D. Grounds b,
G.J. Pinniger a,⇑
a
School of Biomedical, Biomolecular and Chemical Sciences, The University of Western Australia, Crawley, WA 6009, Australia
b
School of Anatomy and Human Biology, The University of Western Australia, Crawley, WA 6009, Australia
Received 14 May 2010; received in revised form 22 September 2010; accepted 23 September 2010
Abstract
This study evaluated the contribution of the pro-inflammatory cytokine, tumour necrosis factor (TNF) to the severity of exerciseinduced muscle damage and subsequent myofibre necrosis in mdx mice. Adult mdx and non-dystrophic C57 mice were treated with
the mouse-specific TNF antibody cV1q before undergoing a damaging eccentric contraction protocol performed in vivo on a custom built
mouse dynamometer. Muscle damage was quantified by (i) contractile dysfunction (initial torque deficit) immediately after the protocol,
(ii) subsequent myofibre necrosis 48 h later. Blockade of TNF using cV1q significantly reduced contractile dysfunction in mdx and C57
mice compared with mice injected with the negative control antibody (cVaM) and un-treated mice. Furthermore, cV1q treatment significantly reduced myofibre necrosis in mdx mice. This in vivo evidence that cV1q reduces the TNF-mediated adverse response to exerciseinduced muscle damage supports the use of targeted anti-TNF treatments to reduce the severity of the functional deficit and dystropathology in DMD.
Ó 2010 Elsevier B.V. All rights reserved.
Keywords: Inflammation; Muscle damage; Isokinetic dynamometer; Duchenne muscular dystrophy; mdx mouse
1. Introduction
Dystrophin is part of the dystrophin–glycoprotein complex that links intra-cellular F-actin to the extra-cellular
matrix [1] and provides mechanical protection for the sarcolemma during muscular contraction [2,3]. In patients
with Duchenne muscular dystrophy (DMD) and in the
mdx mouse and golden retriever (GRMD) dog models of
DMD, the absence of dystrophin disrupts normal force
transmission and reduces the stability of the sarcolemma
[4]. As a result, dystrophic skeletal muscles are more
susceptible to exercise-induced muscle damage (EIMD),
⇑ Corresponding author. Tel.: +61 8 6488 3380; fax: +61 8 6488 1025.
E-mail address: [email protected] (G.J. Pinniger).
particularly during lengthening (eccentric) muscle actions
[5,6]. The high susceptibility to muscle damage and
repeated cycles of myofibre necrosis, especially throughout
the growth phase [7], ultimately results in the failure of
myofibres to regenerate and the replacement of myofibres
with fat or fibrous connective tissue. It is likely that an elevated inflammatory response contributes to the progressive
loss of skeletal muscle mass and function seen in the dystrophic condition [8]. While the pro-inflammatory cytokine
tumour necrosis factor (TNF) has been associated with
chronic muscle wasting conditions such as cachexia [9]
and sarcopenia [10], it has also been implicated in the acute
inflammatory response that exacerbates muscle damage
and contractile dysfunction in mdx mice [11–14].
EIMD is characterized by an initial decrease in the force
producing capacity of the muscle (contractile dysfunction)
0960-8966/$ - see front matter Ó 2010 Elsevier B.V. All rights reserved.
doi:10.1016/j.nmd.2010.09.013
Please cite this article in press as: Piers AT et al., Blockade of TNF in vivo using cV1q antibody reduces contractile dysfunction of skeletal muscle in
response to eccentric exercise in dystrophic mdx and normal mice, Neuromuscul Disord (2010), doi:10.1016/j.nmd.2010.09.013
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immediately after the exercise and a subsequent secondary
decline in force producing capacity that develops over the
following days. Although the initial force deficit has been
attributed to the impairment of excitation–contraction
coupling [15] and/or disruption of the contractile filaments
[16], this muscle damage also coincides with an inflammatory response including the rapid release of TNF from
the damaged muscle and resident mast cells [17,18]. TNF
attracts neutrophils and further inflammatory cells (that
also produce TNF) to the injured site thereby increasing
the inflammatory cascade. When myofibre injury occurs,
inflammation and associated cytokines are essential for
coordinating the removal of damaged tissue and for formation of new skeletal muscle [19]. However, in situ experiments examining EIMD in mice have shown that the
accumulation of neutrophils and macrophages after injury
contributes to further muscle damage [20,21] and, along
with ex vivo studies on cultured myotubes [22], it has been
suggested that this effect may be mediated by increased
production of reactive oxygen species (ROS). In the mdx
mouse, an excessive state of inflammation can exacerbate
myofibre necrosis [19,23]. In normal (non-dystrophic) muscle, experimentally elevated TNF (in vivo for 7 days) results
in the accumulation of inflammatory cells without necrosis
[24]: however, elevated TNF (ex vivo) has rapid adverse
effects on contractile function of normal myofibres with
early loss of force due (at least in part) to associated oxidative stress [25]. Therefore, the immediate acute increases in
TNF may contribute to the initial muscle weakness following EIMD, whereas a prolonged inflammatory response
may lead to the myofibre necrosis and progressive muscle
wasting that is characteristic of the dystrophic pathology.
Despite some equivocal reports [26], numerous studies
indicate that TNF levels are elevated in mdx mice and
human DMD patients [27–30]. Elevated local TNF, when
combined with the increased susceptibility to EIMD, contributes to myofibre necrosis and, ultimately, the fibrosis
inherent in DMD, GRMD and mdx muscle [23]. In vivo
experiments on mdx mice have demonstrated that blockade
of TNF [11,12,14], or depletion of host neutrophils [13],
reduces the severity of the dystrophic pathology. The two
drugs used to block TNF activity, RemicadeÒ (antibody
to human TNF, also known as Infliximab) and EnbrelÒ
(soluble receptor to TNF, also known as Etanercept), are
widely used clinically to treat inflammatory disorders such
as arthritis and Crohn’s disease [31]. To increase the efficacy of TNF blockade in mice and to avoid potential problems of immune response to the human constant domain
sequences of infliximab, a rat monoclonal antibody specific
for mouse TNF [32] was modified to generate a chimerical
monoclonal antibody with rat variable and mouse IgG2a,
kappa constant domains (cV1q; [11,33]).
This study used a custom built mouse dynamometer [34]
to quantify precisely the initial damage of dystrophic muscle and the impact of cV1q mediated TNF blockade on the
initial contractile dysfunction and subsequent myofibre
necrosis in response to a bout of EIMD performed in situ.
2. Materials and methods
2.1. Animals
All experiments were performed on 10–13 week old
adult male dystrophic C57BL/10ScSnmdx/mdx and control
C57Bl/10ScSn mice, hereafter referred to as mdx and
C57, respectively. Adult mdx mice were used in these experiments since the level of myofibre necrosis and regeneration
has stabilised to a relatively low level at this age [35,36].
The mice, obtained from a specific pathogen free colony
at the Animal Resource Centre, (Murdoch, Western Australia), were housed in cages, supplied with food and water
without restriction, and maintained in a 12 h light/dark airconditioned (20–25 °C) environment. All animal procedures were approved by the Animal Ethics and Experimentation Committee of the University of Western Australia in
accordance with the guidelines of the National Health and
Medical Research Council of Australia.
2.2. Experimental outline
Experiments were performed using a custom built
mouse dynamometer [34] to induce controlled and consistent eccentric EIMD in vivo and to assess the contractile
parameters of muscles from mdx and C57 mice. Initial contractile dysfunction was quantified by comparing the isometric torque production before and after the EIMD
protocol (initial damage), while histological techniques
were used to quantify subsequent (secondary) myofibre
necrosis. The effect of cV1q mediated blockade of TNF
on initial and secondary muscle damage was compared
with un-treated and sham injected (cVaM) control mice.
2.3. Experimental procedures
Mice were anaesthetised by inhalation of a gaseous mixture of isoflurane (isoflurane, 0.4 L/min N2O, 0.4 L/min
O2) which was maintained for the duration of the experiment (approximately 1 h) via a flow-through facemask over
the mouse’s head. Anaesthetised mice were connected to
the dynamometer for the measurement of contractile function and to induce EIMD. The dynamometer protocol performed in this experiment is described in detail by Hamer
et al. [37] and explained briefly below.
2.4. Dynamometer protocol for eccentric exercise-induced
muscle damage
Before performing the EIMD protocol, optimal stimulation parameters were determined based on the torque–volt
and torque–frequency relationships for isometric contractions (200 ms train) recorded at a neutral ankle angle (i.e.
the foot positioned perpendicular to the tibia). The optimal
stimulation voltage from all experiments ranged from 1 to
10 V and the optimal stimulation frequency ranged from
125 to 300 Hz. There were no significant differences in these
Please cite this article in press as: Piers AT et al., Blockade of TNF in vivo using cV1q antibody reduces contractile dysfunction of skeletal muscle in
response to eccentric exercise in dystrophic mdx and normal mice, Neuromuscul Disord (2010), doi:10.1016/j.nmd.2010.09.013
ARTICLE IN PRESS
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parameters between mouse strain or treatment group, nor
did the EIMD protocol significantly affect these parameters.
These stimulation parameters were then used when recording the torque–angle relationship and when performing the
EIMD protocol consisting of 20 eccentric contractions.
After a 10 min recovery period following EIMD, the initial
procedures of torque–volt, torque–frequency and torque–
angle were repeated for assessment of initial contractile dysfunction. Specific details of these procedures are as follows.
2.5. Torque–angle relationship
A torque–angle relationship for the anterior crural muscles was determined from isometric contractions elicited at
5° increments between 15° dorsiflexion and 55° plantar
flexion. The foot was passively rotated to 15° dorsiflexion
and a 200 ms train of pulses delivered to the common peroneal nerve to elicit an isometric contraction. Immediately
after stimulation had ceased the ankle was rotated 5°
towards plantar flexion and rested at this angle for 30 s
prior to recording of isometric torque at the new joint
angle. The procedure was repeated at 5° increments up to
55° plantar flexion, with the isometric torque being
recorded at each position.
Each isometric contraction was analysed for the mean
peak torque during the final 80 ms of activation. The peak
torque was plotted against joint angle and a Gaussian fit
was applied using Curve Expert v1.36. The Gaussian model
fitted was of the form:
y¼a
ðxbÞ2
2c2
The coefficients of this fit are: a = the amplitude of the
Gaussian fitted curve, equivalent to the peak joint torque;
b = the angle at which peak torque was generated and c
is the width at half the peak.
2.6. Exercise-induced muscle damage
After the torque–angle relationship was established, an
exercise protocol of 20 eccentric contractions was per-
3
formed between ankle angles of 15° and 55° plantar flexion
with a 30 s rest interval between each trial. The anterior
crural muscles were stimulated initially at an ankle angle
of 15° plantar flexion to produce an isometric contraction
(Fig. 1A). After 100 ms of stimulation the ankle was
rapidly rotated to 55° plantar flexion (Fig. 1B) at an
angular velocity of 1000 °s1, thus producing an eccentric
contraction in the anterior crural muscles. Immediately
following the eccentric contraction, the position of the foot
was passively returned to the starting position at 10 °s1
under servomotor control. After a 10 min rest period, the
torque–volt, torque–frequency and torque–angle procedures were repeated to determine the decrease in the peak
joint torque resulting from the EIMD protocol (Fig. 2).
At completion of the final procedure the incision was
sutured and the mouse was allowed to recover. All mice
recovered quickly from anaesthesia, were fully mobile
within minutes of regaining consciousness and displayed
no signs of abnormal limb movement.
2.7. Evans Blue Dye injections
Intra-peritoneal (IP) injections of a 1% sterile solution
(w/v) of Evans Blued Dye (EBD) in phosphate–buffered
saline (PBS, pH 7.5) at a volume of 1% of body weight were
administered 24 h before the EIMD protocol. Prior to
injection, the EBD solution was sterilised by passage
through a MillexÒ-GP 0.22 mm filter and stored at 4 °C.
After the injection, animals were returned to their cage
and allowed food and water ad libitum.
2.8. Histological assessment of myofibre necrosis and
sarcolemmal damage
Mice were sacrificed by cervical dislocation while under
terminal anaesthesia at 48 h after the EIMD protocol.
Body weight was recorded prior to sacrifice. The TA muscles of the test limb (subjected to EIMD) and the nontested (contralateral) limb were excised, sliced transversely
through the mid-belly of the muscle, mounted vertically in
Fig. 1. Mouse dynamometer apparatus with the ankle of the right hind limb at (A) 15° plantar flexion and (B) 55° plantar flexion during the EIMD
protocol. The knee was clamped and the foot attached to a foot plate with the ankle aligned with the axis of the torque transducer. The anterior crural
muscles were stimulated to contract via hook electrodes connected to the common peroneal nerve via a small incision in the skin and muscle fascia near the
head of the fibular. During the EIMD protocol the anterior crural muscles were stimulated to contract at an initial ankle angle of 15° PF and rapidly
rotated at 1000 °s1 to 55° PF during the contraction.
Please cite this article in press as: Piers AT et al., Blockade of TNF in vivo using cV1q antibody reduces contractile dysfunction of skeletal muscle in
response to eccentric exercise in dystrophic mdx and normal mice, Neuromuscul Disord (2010), doi:10.1016/j.nmd.2010.09.013
ARTICLE IN PRESS
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Fig. 2. The joint torque–angle relationships before (solid symbols) and after (open symbols) the dynamometer EIMD protocol in mdx and C57 mice. Data
are presented as the mean (±SE) normalised joint torque at ankle joint angles from 15° (dorsiflexion) to 55° (plantar flexion) for (A) untreated C57 mice,
(B) C57 mice treated with cV1q, (C) untreated mdx mice, and (D) mdx mice treated with cV1q. Vertical dashed line indicates the neutral ankle angle (0°,
when the foot is perpendicular to the leg).
tragacanth gum on cork blocks and snap frozen in isopentane, cooled by liquid nitrogen, and stored at 80 °C.
Alternate frozen sections (8 lm) were cut for EBD and
Haemotoxylin and Eosin (H&E) slides on a Leica
(CM3050) cryostat at 21 °C.
The H&E stained muscle sections were observed with
bright-field light microscopy, non-overlapping tiled images
of transverse muscle sections were taken at 10 magnification to produce an overall image of the entire muscle crosssection. Images were acquired using a Leica DM RBE
microscope, Stage Pro movement software and a Q Imaging
Micropublisher 3.3 RTV digital camera. Muscle morphology was analysed from the tiled images using Image Pro
Plus 6.2 software. Areas of myofibre necrosis were identified
by the presence of fragmented sarcoplasm and the infiltration of inflammatory cells. Myofibre necrosis was calculated
as a percentage of the whole muscle cross-sectional area.
The unstained EBD sections were viewed by red autofluorescence microscopy (Fluoro filter N.2.1: Green) at
10 magnification and tiled the same as for H&E. Sections
with less than 20 EBD positive fibres (less than 1% of total
area) were not tiled. EBD positive fibres were measured
using Image Pro Plus 6.2 and expressed as a percentage
of total muscle cross-sectional area.
2.9. Use of cV1q antibody to block TNF
This study investigated the effectiveness of blocking
TNF using the mouse specific anti-TNF antibody cV1q
(Centocor USA), which is a rat/mouse chimeric, specific
for murine TNF [11]. A negative control solution of cVaM
was also administered, which is an isotope matched control
antibody for cV1q. These reagents were generously provided to us by Centocor (USA). In the initial experiments
(cV1q-1w) both mdx and C57 mice received two intra-peritoneal injections of either cV1q or cVaM made up in PBS
at 1 week and 1 h prior to the EIMD at a concentration of
20 lg/g body weight. In additional experiments (cV1q-4 h),
mdx mice received a single injection of cV1q (20 lg/g body
weight) at 4 h prior to EIMD (dosages based on [11]).
2.10. Data analysis
All data are presented as mean ± SE, unless stated
otherwise. Unpaired student t-tests were performed for
direct comparison of untreated C57 and mdx mice. Multiple comparisons between all groups and between tested and
un-tested (contralateral) limb muscles were made using univariate analysis of variance (ANOVA). A least significant
difference (LSD) post-hoc t-test was used to identify differences between groups. Significance was set at P 6 0.05 for
all groups.
As a quantitative analysis of the benefit of cV1q treatment, a ‘recovery score’ for the correction of the mdx
defect was calculated as ([cV1q-untreated mdx]/[C57untreated mdx]) 100 where a score of 100% indicates that
the parameter in treated mdx mice was equal to that of
control C57 mice, and 0% indicates that no gain was
Please cite this article in press as: Piers AT et al., Blockade of TNF in vivo using cV1q antibody reduces contractile dysfunction of skeletal muscle in
response to eccentric exercise in dystrophic mdx and normal mice, Neuromuscul Disord (2010), doi:10.1016/j.nmd.2010.09.013
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5
obtained compared with untreated mdx mice [38]. Recovery scores were generated for the initial torque deficit and
subsequent myofibre necrosis for mdx mice from the
cV1q-1w and cV1q-4h protocols.
3. Results
The number of animals in each group, along with mean
age and body weight, are presented in Table 1. When the
body weights of mice in each treatment were grouped by
mouse strain, the mean body weight for mdx mice
(29.3 ± 0.53 g) was significantly higher than for C57 mice
(24.9 ± 0.38 g, P < 0.05). Due to the significant difference
in body weights between mdx and C57 mice, peak isometric
torque was normalised to body weight and presented in
units Nmm/kg.
3.1. Muscle contractile properties of untreated mdx and C57
mice
3.1.1. Peak torque production and optimal joint angle
Peak isometric torque normalised to body weight
(Fig. 3A) was significantly lower in mdx mice compared
to C57 mice (P = 0.029). The muscles of mdx mice were
25% weaker than the non-dystrophic C57 mice, which
is consistent with the significantly lower specific force in
TA recorded by Dellorusso et al. [6], from in situ experiments in mdx and C57 mice. Furthermore, the optimal
angle at which the peak torque was recorded was significantly greater for mdx mice compared to C57 mice
(Fig. 3B). This indicates a shift in maximal force production to longer muscle lengths which may reflect an increase
in series compliance within the muscle [39] due to the
underlying dystropathology in mdx mice.
3.1.2. Susceptibility to EIMD
The effects of the EIMD protocol on torque–angle relationships for C57 and mdx mice are presented in Fig. 2A
and C, respectively. Initial contractile dysfunction was
quantified physiologically as the percentage decrease in
peak isometric torque after the EIMD protocol (Fig. 3C).
Both mdx and C57 mice experienced a marked decrease
in mean peak joint torque after the 20 eccentric contractions. However, the deficit in mean peak joint torque
resulting from the EIMD protocol was twofold greater in
Table 1
Age and body weight of mdx and C57 mouse groups. Data are presented
as mean ± SE
mdx untreated
mdx cVaM
mdx cV1q (1 week)
mdx cV1q (4 h)
C57 untreated
C57 cVaM
C57 cV1q (1 week)
n
Age (weeks)
Body weight (g)
7
4
7
5
4
4
5
12.0 ± 0.3
12.2 ± 0.2
11.7 ± 0.7
11.3 ± 0.8
11.0 ± 0.6
11.7 ± 0.7
10.2 ± 0.2
30.2 ± 1.1
29.0 ± 0.7
29.2 ± 2.1
29.2 ± 0.9
25.4 ± 0.9
24.9 ± 0.5
24.4 ± 0.6
Fig. 3. Mean (±SE) for (A) peak isometric torque production in mdx and
C57 mice before the dynamometer EIMD protocol. (B) Optimal joint
angle, at which the peak torque was recorded (data presented as ankle
angle in degrees plantar flexion). (C) The torque deficit (contractile
dysfunction) in mdx and C57 mice after the EIMD protocol. (D) Muscle
necrosis in TA muscles from the dynamometer tested and control (contralateral) limbs of mdx mice sampled 48 h after the EIMD protocol.
*significantly different compared to mdx mice (P < 0.05); # significantly
different to test limb (P < 0.05).
mdx compared to C57 mice (P = 0.02). The significantly
greater torque deficit in mdx compared to C57 mice confirms that dystrophic muscles are more susceptible to
EIMD resulting from controlled eccentric contractions,
as performed here in vivo with the mouse dynamometer.
This is consistent with previous EIMD studies in mdx mice
using isolated EDL muscles [40] and also for in vivo dynamometer studies in GRMD dogs [5].
3.1.3. Histological assessment of myofibre necrosis and
sarcolemmal damage
Myofibre necrosis was not evident in any C57 muscles,
in accordance with previous observations in our laboratory
following voluntary wheel running or horizontal treadmill
running (Radley-Crabb, unpublished data). Therefore, to
obtain a reliable measure of the extent of myofibre necrosis
resulting from EIMD in mdx mice, comparisons were made
between the TA muscles of the test limb and the un-tested
limb (contralateral control). The mean area of necrosis
increased about fourfold in tested mdx muscle compared
to un-tested (contralateral) muscle (P = 0.002) (Fig. 3D).
Evidence of necrosis in the un-tested control leg gives an
Please cite this article in press as: Piers AT et al., Blockade of TNF in vivo using cV1q antibody reduces contractile dysfunction of skeletal muscle in
response to eccentric exercise in dystrophic mdx and normal mice, Neuromuscul Disord (2010), doi:10.1016/j.nmd.2010.09.013
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indication of the severity of the background dystropathology in both legs prior to the EIMD protocol, although it is
recognised that there can be biological variation even
between both legs of an individual mdx mouse [35].
As for the analysis of myofibre necrosis, there was no
evidence of EBD positive fibres in any of the tested muscle
from C57 mice. For the TA muscles of mdx mice, however,
EBD positive fibres accounted for 5.6 ± 1.3% of total muscle cross-sectional area (Fig. 8A).These results indicate that
the muscles of the mdx mice are weaker (lower peak torque) and more susceptible to EIMD than non-dystrophic
C57 mice. The absence of myofibre necrosis and EBD positively stained fibres in any muscles from the C57 mice
reflects the capacity for normal (non-dystrophic) mice to
tolerate EIMD.
3.2. The effect of cV1q treatment on mdx and C57 muscles
after EIMD
3.2.1. The effect of cV1q on peak torque and optimal joint
angle
Peak isometric torque prior to EIMD normalised to
body weight for all mdx and C57 groups is presented in
Fig. 4A. There was a significant main effect of mouse strain
on peak torque (F1,28 = 10.6, P < 0.001) (mdx versus C57)
and post-hoc analysis revealed that the mdx mice were significantly weaker than the non-dystrophic C57 mice
(P < 0.01). There was no significant main effect of treatment (F2,28 = 8.6, P = 0.08) on the peak torque. The joint
angle at which the peak torque was recorded (optimal
angle) for all mdx and C57 groups is presented in
Fig. 4B. The optimal angle for untreated mdx and C57
mice that was reported in Fig. 3B is also included here
for comparison with the treatment groups. There was no
significant main effect of treatment (F2,28 = 1.09,
P = 0.35) on the optimal joint angle.
3.2.2. Effect of cV1q treatment on susceptibility to EIMD
The torque–angle relationships recorded before and
after the EIMD protocol in cV1q-treated C57 and mdx
mice are presented in Fig. 2B and D, respectively. The initial contractile dysfunction as determined by torque deficit
after EIMD is presented in Fig. 5 for all mdx and C57
groups. There was a significant main effect of mouse strain
(F1,28 = 43.3, P < 0.001) (mdx versus C57) on the torque
deficit after EIMD. Post-hoc analysis revealed that the
C57 mice experienced significantly less torque deficit than
the mdx mice (P < 0.001). Furthermore, there was a significant main effect of treatment (F2,28 = 8.6, P = 0.002) with
cV1q-1w significantly reducing the torque deficit in both
mdx and C57 mice (P < 0.001). The acute cV1q-4h treatment, examined in mdx mice, also significantly reduced
the torque deficit following EIMD (P < 0.05).
3.2.3. Assessment of cV1q treatment on necrosis and
sarcolemmal damage after EIMD
Necrosis was quantitated on H&E stained sections
(Fig. 6): since necrosis was not observed in any C57 mice,
only data from mdx mice are presented (Fig. 7). The % area
of necrosis in the TA muscle of tested limbs was compared
to the non-tested contralateral limb. Significant main
effects of leg (F1,28 = 19.0, P < 0.001) (whether dynamometer tested or contralateral un-tested limb) and treatment
(F2,28 = 4.0, P = 0.029) were observed on muscle necrosis
in mdx mice. Post-hoc analysis revealed that for all treatment groups the level of necrosis was significantly greater
in the TA muscle of the tested limb compared to the untested limb (P < 0.05). Furthermore, exposure to cV1q
for 1 week (cV1q-1w) significantly reduced the level of
myofibre necrosis in the TA muscles of both the tested limb
(Fig. 6) and the un-tested limb, compared to untreated mdx
Fig. 5. Mean (±SE) peak torque deficit (contractile dysfunction) in
treated and untreated mdx and C57 mice after the dynamometer EIMD
protocol. # significantly different from untreated; *significantly different
from corresponding mdx group; P < 0.05.
Fig. 4. Mean (±SE) data for treated and untreated mdx and C57 mice recorded prior to the EIMD protocol for (A) peak isometric torque normalised to
body weight. (B) Optimal joint angle. *significantly different from corresponding mdx group; P < 0.05.
Please cite this article in press as: Piers AT et al., Blockade of TNF in vivo using cV1q antibody reduces contractile dysfunction of skeletal muscle in
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Fig. 6. Typical examples of myofibre necrosis in H&E stained transverse sections of TA muscles subjected to the dynamometer EIMD protocol. (A)
Untreated mdx muscle showing myofibre necrosis, fragmented sarcomeres and inflammatory cell accumulation. Similar necrosis is observed in (B) cVaM
(control IgG) treated mdx muscle. (C) mdx muscle treated with cV1q (anti-TNF IgG) for 1 week prior to EIMD protocol showing areas of centrally
nucleated myofibres but far less myofibre necrosis. Scale bars represent 0.1 mm.
Fig. 7. Mean (±SE) area of myofibre necrosis in dynamometer tested and
un-tested (control) TA muscles of mdx mice sampled 48 h after the
dynamometer EIMD protocol. *significantly different from corresponding
test leg; # significantly different from untreated group; P < 0.05.
mice (P < 0.05). However, the level of necrosis in the acute
cV1q-4h treated mdx mice was not significantly different
from the un-treated mdx mice (in either tested or un-tested
limbs). These results show that 1 week exposure to cV1q in
mdx mice significantly reduced necrosis in both the tested
and un-tested (control) limbs compared to untreated mdx
mice. The effects of cV1q-1w and cVaM treatments on sarcolemmal damage were evaluated by the analysis of EBD
positively stained myofibres (Fig. 8). Neither cV1q-1w
nor cVaM treatments had any significant effect on the number of EBD positive fibres in mdx mice (F2,15 = 0.52,
P = 0.60). EBD staining was not performed in cV1q-4h
treated mice.
Fig. 8. TA cross sections showing positive staining for Evans Blue Dye (EBD) in myofibres 48 h after the EIMD protocol for (A) untreated mdx, (B) mdx
mice treated with cV1q for 1 week, and (C) mdx mice treated with cVaM for 1 week. Scale bars represent 500 lm. (D) Mean (±SE) area of EBD positively
stained fibres in TA muscles of mdx mice sampled 48 h after the dynamometer EIMD protocol. Note: There were no (<1%) EBD positive fibres observed
in C57 mice.
Please cite this article in press as: Piers AT et al., Blockade of TNF in vivo using cV1q antibody reduces contractile dysfunction of skeletal muscle in
response to eccentric exercise in dystrophic mdx and normal mice, Neuromuscul Disord (2010), doi:10.1016/j.nmd.2010.09.013
ARTICLE IN PRESS
8
A.T. Piers et al. / Neuromuscular Disorders xxx (2010) xxx–xxx
Calculation of the ‘recovery score’ shows a correction by
cV1q-1w treatment of the mdx values (control untreated
value as 0%) towards those for normal C57 mice (considered as 100%) of 78% for torque deficit and 56% for myofibre necrosis. The recovery score for the torque deficit in
this study is similar to the 81% recovery reported by Gillis
[38] for isolated EDL muscles of transgenic mdx mice overexpressing a truncated form of utrophin. The recovery
score for mdx mice exposed to a single cV1q-4h injection
was 56% for torque deficit but only 5% for necrosis.
4. Discussion
The contractile dysfunction of dystrophic mdx muscles
(compared with normal controls) measured by the dynamometer in this study is similar to that for the GRMD
dog model of DMD [5] and for transgenic mdx mice that
over express insulin-like growth factor-1 [34]. The main
findings of this study support our hypothesis that blockade
of TNF (using cV1q antibody) would significantly reduce
the extent of contractile dysfunction and (in some cases)
subsequent myofibre necrosis in mdx mice following a bout
of damaging eccentric exercise. These data further support
the use of cV1q administration to protect dystrophic muscles from initial and longer-term damage.
Exposure to cV1q for 1 week before EIMD significantly
reduced the initial contractile dysfunction (recovery
score = 78%) and the subsequent myofibre necrosis (recovery score = 56%) in response to EIMD. The beneficial
effects of cV1q in reducing myofibre necrosis has also been
demonstrated in mdx mice exposed to voluntary wheel running [11] with similar effects observed for other anti-TNF
treatments (e.g. Remicade and Enbrel [12–14]). Interestingly, TNF blockade was also beneficial and reduced the
initial contractile dysfunction in C57 mice, although to a
lesser extent than for mdx mice. This observation suggests
that, in addition to its contribution to myofibre necrosis,
TNF also plays a crucial role in mediating the initial functional weakness following EIMD in both normal and dystrophic muscles: this accords with ex vivo studies on nondystrophic muscles [25].
It is worth noting that the exposure to cV1q for 1 week
reduces not only necrosis after EIMD (in the test leg) but
also the background pathology in the mdx mice as indicated by a significant (albeit small) reduction in necrosis
in the un-tested limb. To distinguish between possible
longer-term beneficial effects of cV1q on the muscle environment compared with more short-term effects of blocking TNF, we conducted experiments with a single cV1q
injection administered 4 h before EIMD. We chose a 4 h
time-point to ensure that the cV1q had sufficient time to
take effect following the IP injection. This short-term exposure to cV1q in mdx mice also reduced the contractile dysfunction following EIMD (recovery score = 56%).
However, this protective effect on force production did
not extend to myofibre necrosis, which was still elevated
in the TA of the tested mdx limb (recovery score = 5%).
Furthermore, the level of necrosis in the un-tested limb
was not significantly different from that of untreated mdx
mice which would indicate that the background level of
pathology was unaffected. Therefore, at least the protective
effect of cV1q on contractile dysfunction must be mediated
by the short-term blockade of TNF rather than some major
adaptation of the mdx environment. This is consistent with
the beneficial effects of cV1q in normal C57 mice in reducing the contractile dysfunction after EIMD. Therefore, we
propose that elevated levels of TNF in response to EIMD
have direct effects on contractile function.
TNF has been widely implicated as a possible mediator
of the muscle weakness observed in conditions of chronic
and acute inflammation (e.g. muscle damage/trauma,
chronic obstructive pulmonary disorder, AIDS, congestive
heart failure etc.). Elevated systemic TNF in mice results in
loss of body weight (by 3 weeks) and decreased muscle fibre
cross-sectional area (by 5 weeks) [9] and, in tissue cultured
myotubes, chronic TNF exposure (up to 3 days) results in
muscle protein catabolism [41]. Acute exposure of isolated
muscle preparations to TNF (for 4 h), however, causes
contractile dysfunction and weakness, without evidence
of catabolism [25]. Our dynamometer results are in accord
with the experiments of Reid et al. [25] who demonstrated
that TNF administration to isolated mouse muscles in vitro
depressed tetanic force without altering the intra-cellular
Ca2+ concentration ([Ca2+]i) in diaphragm and limb muscles of non-dystrophic mice. Other in vivo experiments by
Hardin et al. [42] show that IP administration of TNF acts
quickly (within 1 h) via the TNF-1 receptor to increase
ROS production and decrease specific force in non-dystrophic mouse diaphragm. These effects were abolished by
treatment with the antioxidant Trolox leading to the suggestion that muscle-derived oxidants are essential postreceptor mediators of TNF-induced force loss [42].
Altered cytosolic Ca2+ homeostasis has also been implicated as a putative pathway in development of the dystropathology [43]. Elevated [Ca2+]i may arise from transient
tears in the fragile sarcolemma of dystrophic myofibres,
and/or through mechanosensitive (stretch-activated) calcium channels. The uptake of EBD following the in vivo
eccentric exercise protocol was limited to 5% of myofibres
in mdx mice, which is slightly less than that reported by
Whitehead et al. (8.6%) from in vitro experiments on
EDL muscles from mdx mice [44]. Interestingly, the level
of EBD staining was unaffected by cV1q treatment suggesting that the beneficial effects of cV1q reported in this study
are not mediated by reducing the sarcolemmal tearing in
dystrophic myofibres. This does not preclude the possibility
that the contractile dysfunction is mediated by altered calcium homeostasis as numerous recent studies have implicated the contribution of transient receptor potential
channels (TRPC) to the dystropathology [45,46] and elevated expression of TRPC1 has been reported in mdx muscle [47,48]. Regardless of the source of entry, the influx of
Ca2+ into cells has been linked to ROS production and
activation of the nuclear factor-kappa B (NF-jB) pathway
Please cite this article in press as: Piers AT et al., Blockade of TNF in vivo using cV1q antibody reduces contractile dysfunction of skeletal muscle in
response to eccentric exercise in dystrophic mdx and normal mice, Neuromuscul Disord (2010), doi:10.1016/j.nmd.2010.09.013
ARTICLE IN PRESS
A.T. Piers et al. / Neuromuscular Disorders xxx (2010) xxx–xxx
[43,49] that regulates pro-inflammatory cytokine expression, in particular TNF [43]. Thus, a putative mechanism
involved in the exercise-induced contractile dysfunction in
mdx mice may involve a TNF mediated positive feedback
relationship with NF-jB [30,50] and ROS [51]. The protective effect of TNF blockade using cV1q might be mediated
(at least in part) by altering this positive feedback cycle,
although further quantitative studies are required to clarify
this hypothesis.
5. Summary
We have shown that cV1q administration (1 week before
EIMD) protects dystrophic skeletal myofibres against initial contractile dysfunction and subsequent myofibre necrosis in adult mdx mice subjected to a damaging eccentric
contraction protocol in vivo using a custom built mouse
dynamometer. This technique enables the precise in situ
quantitation of the initial physiological damage of muscles
and evaluation of the impact of cV1q mediated TNF blockade on both contractile dysfunction and myofibre necrosis.
The results from this study indicate that TNF acts quickly
(within 30 min) to cause muscle weakness. Short term (4 h)
administration of cV1q protects against TNF-mediated
contractile dysfunction but not necrosis of mdx muscles,
indicating that different mechanisms contribute to these
events. The controlled EIMD produced by the dynamometer, with precise timing of damage combined with the ability to repeatedly measure functional parameters in vivo
(including force production, muscle fatigue and adaptation
[34]) makes this a powerful tool to test the efficacy of various therapeutic interventions and to more precisely define
the molecular mechanisms responsible for initial and
longer-term damage of dystrophic muscles.
Acknowledgements
We gratefully acknowledge Centocor (USA) for providing the cV1q anti-mouse TNF antibody and the negative
control antibody (cVaM) and Dr. David Shealy for his support and helpful comments regarding this work. We also
thank Professor Peter Hamer (The University of Notre
Dame, Australia) for his contribution to the dynamometer
work. This research was supported by the Raine Medical
Research Foundation, UWA, the Medical Health Research
Infrastructure Fund of WA and the National Health and
Medical Research Council of Australia.
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response to eccentric exercise in dystrophic mdx and normal mice, Neuromuscul Disord (2010), doi:10.1016/j.nmd.2010.09.013
Journal of Biomedical Optics 13共1兲, 011003 共January/February 2008兲
Three-dimensional optical coherence tomography of
whole-muscle autografts as a precursor to morphological
assessment of muscular dystrophy in mice
Blake R. Klyen
Julian J. Armstrong
Steven G. Adie
The University of Western Australia
School of Electrical, Electronic and Computer
Engineering
Optical+Biomedical Engineering Laboratory
35 Stirling Highway
Crawley, Western Australia 6009
Australia
Hannah G. Radley
Miranda D. Grounds
The University of Western Australia
School of Anatomy and Human Biology
Skeletal Muscle Research Group
35 Stirling Highway
Crawley, Western Australia 6009
Australia
David D. Sampson
Abstract. Three-dimensional optical coherence tomography 共3DOCT兲 is used to evaluate the structure and pathology of regenerating
mouse skeletal muscle autografts for the first time. The death of myofibers with associated inflammation and subsequent new muscle formation in this graft model represents key features of necrosis and
inflammation in the human disease Duchenne muscular dystrophy.
We perform 3D-OCT imaging of excised autografts and compare OCT
images with coregistered histology. The OCT images readily distinguish the necrotic and inflammatory tissue of the graft from the intact
healthy muscle fibers in the underlying host tissue. These preliminary
findings suggest that, with further development, 3D-OCT could be
used as a tool for the evaluation of small-animal muscle morphology
and pathology, in particular, for analysis of mouse models of muscular
dystrophy. © 2008 Society of Photo-Optical Instrumentation Engineers.
关DOI: 10.1117/1.2870170兴
Keywords: optical coherence tomography; three-dimensional imaging; muscle;
muscular dystrophy; small-animal imaging.
Paper 07117SSRR received Mar. 30, 2007; revised manuscript received Aug. 30,
2007; accepted for publication Nov. 16, 2007; published online Feb. 25, 2008.
The University of Western Australia
School of Electrical, Electronic and Computer
Engineering
Optical+Biomedical Engineering Laboratory
35 Stirling Highway
Crawley, Western Australia 6009
Australia
1
Introduction
Muscle tissue from experimental animal models is routinely
studied to understand the cause, disease progression, and efficacy of treatment of human Duchenne muscular dystrophy.
This common form of muscular dystrophy, occurring in approximately 1 in 3500 male births worldwide,1 is characterized by the progressive degeneration and loss of skeletal
muscle, leading to death from respiratory or cardiac failure.2
Myofiber 共muscle fiber兲 necrosis 共cell death兲 normally results
in new muscle formation to repair the damaged tissue, but in
Duchenne muscular dystrophy repeated cycles of necrosis
over time result in failed regeneration with the progressive
loss of myofibers and replacement with fibrous and fatty connective tissue.3
The mouse whole-muscle autograft provides a large mass
of tissue with distinctive histological features of muscle necrosis, inflammation, and regeneration 共representing events in
human disease兲: this model is well characterized, highly predictable, and reproducible between animals.4 The model also
Address all correspondence to: Blake R. Klyen, Optical+Biomedical Engineering
Laboratory, School of Electrical, Electronic and Computer Engineering M018,
The University of Western Australia, 35 Stirling Highway, Crawley, Western
Australia 6009 Australia. Tel: +61-8-6488-3916; Fax: +61-8-6488-1319; E-mail:
[email protected]
Journal of Biomedical Optics
provides a convenient in vivo bioassay for assessing pharmacological interventions designed to reduce inflammation associated with necrosis5,6 共as also occurs in muscular dystrophy兲.
Such drug interventions have been shown to ameliorate skeletal muscle necrosis and pathology in the X-chromosomelinked muscular dystrophy 共mdx兲 mouse experimental animal
model for human Duchenne muscular dystrophy,3,6,7 and in
patients with the disease.2,3,8
Histology is conventionally used to study the morphology
of skeletal muscle. Typically 共e.g., for whole-muscle grafts兲,
the muscle tissue is removed from the animal, processed, and
cut into transverse sections that are stained to identify and
measure the amount of healthy and necrotic muscle tissue, the
extent of inflammation, and the formation 共density and size兲
of myotubes 共multinucleated new muscle fibers兲 within the
regenerating region.9 However, the full histological process is
laborious and time consuming and not amenable to examining
large tissue volumes. It requires sacrifice of the animal for
tissue harvesting, tissue processing, and preservation; sectioning and staining; and microscopic examination of many images. It would be preferable to be able to perform detailed 3D
analysis on large volumes of tissue in situ without the need for
muscle biopsy or animal sacrifice, and subsequent sectioning,
1083-3668/2008/13共1兲/011003/7/$25.00 © 2008 SPIE
011003-1
January/February 2008
쎲
Vol. 13共1兲
Klyen et al.: Three-dimensional optical coherence tomography of whole-muscle autografts…
staining, and imaging. Furthermore, longitudinal studies require the sacrifice of multiple animals for each time point,
preventing the observation of sequential development of the
same tissue within an individual animal. In vivo imaging,
without the need for sacrifice, would enable repeated observation on an individual animal 共optimal experimental scenario兲 and decrease the number of animals necessary per experiment 共this is ethically desirable and has the added
advantage of reduced cost兲. The opportunity to track the same
tissue over time would greatly reduce the complication of
biological variation between animals 共at each time point兲 that
is a considerable problem in conventional sampling. Such an
increase in the efficiency and throughput of tissue characterization would signify a major shift in current experimental
technique with wide application to animal studies and potential human tissue analysis.
Biomedical imaging modalities that could potentially be
used to characterize skeletal muscle pathology, ex vivo and in
vivo, include ultrasonography, magnetic resonance imaging
共MRI兲, confocal and multiphoton microscopy, and optical coherence tomography. Ultrasonography10 has been used clinically to probe the gross structure of human muscle tissue with
large penetration depth, but is unable to resolve individual
myofibers with an average diameter of 30 to 50 ␮m. MRI has
been used for in vivo studies of muscle in whole live
animals,11,12 but requires an exogenous contrast agent for localized detection of muscle pathology,13,14 and its resolution14
of 125 ␮m is insufficient to image the individual myofibers.
Confocal and multiphoton microscopy, with their superior resolution, can resolve individual myofibers in vivo,15 but are
limited in penetration depths to a few hundred micrometers,
and routinely depend on the detection of fluorescent signals
from exogenous contrast agents.
Optical coherence tomography 共OCT兲 is an emerging imaging modality capable of real-time, noninvasive, crosssectional imaging of thick biological tissue approaching micrometer resolution.16 OCT uses near-IR light to perform
optical ranging analogous to ultrasound B-mode imaging, but
with resolution of two orders of magnitude greater. Typically,
cross-sectional images are generated, which represent the spatially localized intensity of backscattered light with a penetration depth of up to 2 to 3 mm in tissue.17 A third scan dimension can be added to enable 3D imaging of samples.
OCT has been used to investigate a wide range of biological tissues and structures,17 but its use in the study of muscle
tissue has been limited. Recently, Pasquesi et al. reported the
first study directed specifically toward skeletal muscle tissue
and muscular dystrophy. This study18 used polarizationsensitive OCT to measure changes in the birefringence of
skeletal muscle of the mdx mouse model for Duchenne muscular dystrophy, and correlated these changes with myofiber
damage due to dystrophy. Other previous studies with OCT
have involved muscle tissue only incidentally; e.g., measuring
the refractive index of cardiac muscle in humans,19 and determining the birefringence of animal skeletal20–24 and
cardiac25,26 muscle using polarization-sensitive OCT.
The purpose of this study is to determine if 3D-OCT is
capable of imaging large volumes of ex vivo unstained mouse
skeletal muscle in sufficient detail to be able to differentiate
healthy and pathological tissue. Such capability is a precursor
to eventual noninvasive in vivo assessment of muscular dysJournal of Biomedical Optics
Fig. 1 共a兲 Schematic of the fiber-optic OCT system: BBS, broadband
source; PD, photodetector; VOA, variable optical attenuator; FDODL, frequency-domain optical delay line; and PC, polarization controller. 共b兲 Whole-muscle autograft sample orientation. The arrow indicates the alignment of myofibers: EDL, extensor digitorum longus
and TA, tibialis anterior. 共c兲 En face 共x-y兲 OCT image plane indicating
direction of inflammatory response 共arrows兲.
trophy in mouse models. We compare, for the first time, 3DOCT images of whole-muscle autografts with conventional
histology, to identify damaged and regenerating muscle adjacent to healthy undamaged myofibers. In the remainder of this
paper, we describe the imaging method, the anatomy of the
whole-muscle autograft, and the surgical procedure. We
present coregistered OCT images and conventional histology
of the autografted muscle tissue, as well as 3D-OCT volume
reconstructions, and discuss the results and prospects for in
vivo imaging.
2
Methods
2.1 OCT
We performed 3D-OCT imaging using a 3D-scanning optical
fiber interferometer, as shown in Fig. 1共a兲. The input light is
split by a 50/ 50 fiber optic coupler to reference and sample
arms, and a circulator is used to enable dual-balanced detection to reduce optical intensity noise.27 The broadband light
source is based on a semiconductor optical amplifier 共JDS
Uniphase, Milpitas, California兲 with a center wavelength of
1330 nm and a full-width-at-half-maximum bandwidth of
45 nm.
Measurement of the sample in the axial direction 共along
the beam兲 was performed by scanning the path length of the
reference arm, using a frequency-domain optical delay line,28
with a measured resolution in air of 19.4 ␮m. Transverse
scanning in a single dimension was performed by a galvanometer mirror and objective lens setup with a beam numerical aperture of 0.07, measured in-focus transverse resolution
共1 / e2 intensity beam diameter兲 of ⬃11 ␮m and calculated
axial Rayleigh range of ⬃72 ␮m. The dual-balanced photodetected difference signal was bandpass filtered and logarithmically demodulated before analog-to-digital conversion
共PCI-6111E, National Instruments, Austin, Texas兲. The OCT
system had a measured detection sensitivity of −95 dB with
2 mW incident on the sample.
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Fig. 2 共a兲 En face OCT image and 共b兲 corresponding histology of a transverse section of the autografted muscle sample showing the grafted EDL
overlying the host TA muscle, separated by a dashed line added for clarity. A 3⫻ magnified view 共right兲 shows the graft boundary, with healthy
myofibers 共H兲; surviving myofibers 共S兲; and inflammatory cells 共I兲.
Two-dimensional 共2D兲 OCT images 共512⫻ 512 pixels兲
were acquired in 1 s, and the third dimension was obtained by
sequential cross-sectional 共y-z兲 scans at 10-␮m intervals
along the x axis obtained by linear translation of the objective
lens. The 3D data sets consisting of ⬃400 images were acquired in 5 min and measured 4 ⫻ 1.8⫻ 1 mm 共x ⫻ y ⫻ z兲. En
face 共x-y兲 OCT images at various sample depths were reconstructed from the 3D data set.
2.2 Animal Model and Whole-Muscle Autograft
Surgery
Three female C57BL/10ScSn 6-week-old mice were obtained
from the Animal Resources Centre, Murdoch, Western Australia. The mice were housed and treated according to the
Western Australian Prevention of Cruelties to Animals Act
共1920兲 and the Australian National Health and Medical Research Council guidelines, under protocols approved by the
University of Western Australia Animal Ethics Committee.
For whole-muscle autograft surgery, the mice were anaesthetized with 1.5% 共v/v兲 Rodia halothane 共Merial, Parramatta,
Australia兲, N2O, and O2 and transplantation of the whole intact extensor digitorum longus 共EDL兲 muscle was performed
on one or both legs, described in detail elsewhere.9,29 Briefly,
the EDL muscle with both tendons attached was removed
from the anatomical bed of the hindlimb of the mouse and
transplanted onto the anterior surface of the tibialis anterior
共TA兲 muscle of the same leg. The tendons of the EDL were
sutured onto the host TA muscle, the skin closed, and the mice
allowed to recover. Five days after the surgery, the mice were
anaesthetized and sacrificed via cervical dislocation, and the
entire TA muscle with attached EDL muscle was excised and
placed in phosphate-buffered saline 共PBS兲 on ice for transporJournal of Biomedical Optics
tation for ex vivo 3D-OCT imaging.
2.3 OCT Imaging Protocol
All muscle samples were imaged within 2 h of animal sacrifice. A number of imaging geometries were investigated on
the first two samples, and the optimum protocol, used on subsequent samples, is described in the following. The sample
was bisected along the midregion to expose the transverse
face of both the host TA and graft EDL, and then placed on an
imaging window and bathed in PBS to prevent drying, and to
reduce the refractive-index mismatch by displacing any air
gaps between the glass plate and sample. The muscle sample
was oriented with its myofibers parallel to the optical axis and
its transverse face parallel to the en face imaging plane, as
shown in Fig. 1共b兲, and full-volume 3D-OCT scans were obtained. The image axial depth was scaled by 1.4, the approximate refractive index of muscle tissue.30 This sample alignment was preferred as it enabled the gross muscle tissue
morphology and individual myofibers of the transverse plane
of the EDL and TA to be displayed over the large area 共4
⫻ 1.8 mm2兲 of the en face OCT image plane, for comparison
with conventional transverse histology.
2.4 Histological Processing and Image Coregistration
Immediately following OCT imaging, samples were fixed in
4% 共w/v兲 paraformaldehyde 共pH 7.6兲 for 30 min, transferred
to 70% 共v/v兲 ethanol followed by automated standard paraffin
preparation 共Shandon Citadel 1000, Thermo Fisher Scientific,
Inc., Waltham, Massachussets兲. Transverse 5-␮m-thick sections were cut on a microtome and collected on glass slides,
and stained with hematoxylin and eosin 共H&E兲 for light microscopy 共DM RBE, Leica Microsystems GmbH, Wetzlar,
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Fig. 3 共a兲, 共c兲, and 共e兲 En face OCT images, and 共b兲, 共d兲, and 共f兲 the corresponding histology of transverse sections of the autografted muscle sample
at depths of 共a兲 and 共b兲 100, 共c兲 and 共d兲 250, and 共e兲 and 共f兲 500 ␮m. Surviving myofibers 共S兲. The 2⫻ magnified view 共insets兲 show the graft
boundary.
Germany兲 and digital image capture 共DXM 1200F, Nikon
Corporation, Ontario, Canada兲. Nonoverlapping images of the
grafts were obtained with an automated microscope translation stage, and images were tiled and stitched together using
commercial software 共ImagePro Plus 4.5.1, Media Cybernetics Inc., Silver Spring, Maryland兲 on a personal computer.
Coregistration of OCT and light microscopy images was performed by matching for depth and visually correlating structural features of interest.
3
Results
All en face images taken from 3D-OCT data sets obtained
using the protocol described in Sec. 2.3 were found to be
consistent in appearance. The typical results presented in this
section were taken from one mouse. Figure 2 shows a representative en face OCT image and the corresponding H&Estained histological image of the transverse face of the EDL
and TA tissues through the midregion of the muscle sample at
a depth of 100 ␮m. A 3⫻ magnified image 共right兲 shows the
graft boundary 共dashed line兲 and nearby healthy myofibers
共H兲 of the healthy host TA tissue. A mean filter 共1⫻3 pixels兲
was applied to this image to reduce speckle noise. The kernel
size was chosen to roughly match the OCT in-focus transverse
resolution. As a result of surgery, the blood and nerve supply
to the EDL muscle is severed, causing the grafted tissue to
undergo
necrosis,
subsequent
inflammation,
and
regeneration.5 The histology in Fig. 2共b兲 shows inflammatory
cells 共I兲 infiltrating the grafted EDL: this is associated with
the revascularization of the graft in part from the host TA. In
the peripheral zone of inflammation, the necrotic tissue is phagocytozed, although some myofibers survive at the periphery
Journal of Biomedical Optics
共S兲, and myoblasts 共precursor muscle cells兲 are activated and
fuse to form myotubes.5 The centrally located area of persisting necrotic tissue 共N兲 remaining within the EDL is termed
the necrotic core, and constitutes approximately 50% of the
entire area of the graft in these samples 共at 5 days兲.
In the OCT image in Fig. 2共a兲, the grafted EDL tissue is
readily distinguished from the host TA tissue, meeting at a
clearly defined boundary 共dashed line兲. The TA tissue displays
a regular pattern of circular features, in marked contrast to the
EDL graft, which lacks any such prominent features, instead
exhibiting a more uniform texture corrupted by the speckle
noise typical of OCT images.16 The 3⫻ magnified image
共right兲 highlights the distinction between the two muscle
types, and the circular features of the OCT image in Fig. 2共a,
right兲 are of a size and shape that correlate well with the
healthy myofibers 共H兲 of the corresponding histology 关Fig.
2共b, right兲兴. In the entire 3D-OCT data set, we observed the
grafted EDL tissue to be clearly distinguished from the host
TA tissue with a distinct boundary between the two.
Three representative en face OCT images at different
depths within the whole-muscle autograft are presented in
Fig. 3, along with coregistered histology. At all depths, the
OCT images 关Figs. 3共a兲, 3共c兲, and 3共e兲兴 correspond well in
overall structure with the histology 关Figs. 3共b兲, 3共d兲, and 3共f兲兴.
The pattern of circular structures in the TA tissue persists
throughout the 3D-OCT data set, with a shape and size similar
to that expected for healthy myofibers. In the grafted EDL
tissue, however, a similar pattern is lacking throughout. The
tissue generally appears more uniform in texture, although
there are sparsely distributed small regions of high intensity,
which resemble the circular features found in the TA tissue.
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The contrast between the two muscle tissue types is more
apparent, particularly at the graft boundary 共see Fig. 3 insets兲,
as the image depth increases. Another difference between the
grafted EDL and host TA tissue is that the average signal
intensity, particularly at the graft boundary as seen in Fig.
3共a兲, 3共c兲, and 3共d兲 共insets兲, is attenuated to a greater extent in
the grafted EDL tissue than in the neighboring region of TA
tissue at the same depth. In the histological sections, the necrotic core within the EDL graft is obvious in all sections, and
constitutes approximately 50% of the entire area of the graft.
However, this feature is not evident in the corresponding OCT
images. The average myofiber diameter of this region is
smaller than those of the host TA muscle.
The 3D-OCT cutaway reconstructions of the autografted
muscle sample reveal the transverse face of the graft EDL
tissue and host TA tissue at three depths into the sample in
Fig. 4. The reconstructed en face plane of the 3D-OCT data
set is by far the most informative section for revealing the
muscle morphology. The x-z and y-z image views in Fig. 4
show far fewer readily identifiable features.
4
Discussion
This preliminary investigation has demonstrated that 3D-OCT
is a promising tool for examining the morphology of muscle
pathology and identifying individual muscle fibers. Previous
studies involving muscle tissue using conventional B-scan
oriented OCT have presented images that do not show individual muscle fibers or reveal significant morphological detail
of the muscle tissue.18,20,21,24,31 A possible exception is the
study by Hsiung et al.32 of the hamster cheek pouch at ultrahigh resolution 共1.5 ␮m at 800 nm兲, wherein, within the superficial 共250 ␮m兲 and deep muscle layers 共750 ␮m兲, structures resembling muscle fibers are observed, although the
origin of these structures was not discussed. Similarly, Guo et
al. presented a B-scan OCT image of rabbit muscle in which
structure is evident, but the nature of this structure was not
discussed, nor was there comparison with histology.22
In working with excised muscle, we were able to optimally
orientate the sample to match the largest scan area 共en face兲 to
the transverse face of the whole-muscle autograft, which is
the orientation of most interest to experimental biologists and
conventionally examined in histological sections. We have
found the availability of 3D data indispensable in identifying
gross and fine features in the autografted muscle sample. The
availability of alternative 2D sectional views provided by the
3D data set, as well as the ability to visualize the muscle
sample in cut-away 3D movies 共not presented here兲, proved
invaluable in orientating the samples and presenting the en
face images shown here. By contrast, our early work 共not
reported兲 recording only 2D sections was less successful. En
face OCT images at all depths were found to be significant for
visualizing the transverse-face whole-muscle autograft structure and features of interest; in contrast, images from the x-z
or y-z planes were less informative.
The en face OCT images generated from the 3D-OCT data
set reveal a distinct difference between the muscle structure of
the grafted EDL and the host TA, which correlates well with
the corresponding histology. We observed that the muscle tissue features were smaller in the histology than in the OCT
images. This shrinkage is due to dehydration of the tissue
Journal of Biomedical Optics
Fig. 4 Three 3D-OCT reconstructions of the autografted muscle
sample, with cut-away sections revealing the transverse face at depths
of 共a兲 100, 共b兲 250, and 共c兲 500 ␮m into the tissue. A dashed line
added for clarity separates the EDL and TA tissues.
during the histological process, as previously reported.33 The
most noticeable feature of the OCT images, confirmed by histology, is the observation of healthy myofibers 共H兲 in the TA
muscle, and their general absence in the graft EDL 关Figs. 2共a兲,
3共a兲, 3共c兲, and 3共e兲兴 with a few exceptions. Surviving myofibers 共S兲 were identified in OCT images of the EDL 关Figs.
2共a兲, 3共a兲, and 3共c兲兴 共confirmed in the corresponding histological sections兲 by their distinctive high-intensity circular
shape comparable with the appearance of the healthy myofibers of the TA, which differentiated them from the generally
featureless and lower intensity surrounding regions containing
necrotic tissue and inflammatory cells. The 3D-OCT volume
enhanced identification of surviving myofibers, as they could
be tracked in depth and seen in successive en face OCT images reconstructed along the z axis 关Figs. 3共a兲 and 3共b兲兴.
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By 5 days after transplantation, revascularization of the
EDL muscle and infiltration of inflammatory cells into the
grafted EDL tissue is well advanced. The inflammatory tissue
in the en face OCT images of the EDL has an appearance
similar to those features observed in the study by Cobb et al.
of wound healing in mouse skin, particularly the granulation
tissue in the dermis.34 The presence of the inflammatory cells
in the EDL with the breakdown of necrotic muscle tissue
significantly increases the density of cell nuclei is this region,
staining dark blue in the histology of Fig. 2共b兲. These nuclei
共⬃1 to 2 ␮m in diameter兲 are around an order of magnitude
smaller than the resolution of the OCT system and therefore
cannot be visualized. This is a plausible explanation for the
more uniform appearance of the EDL muscle graft, which is
especially obvious at the boundary of the two tissues, as seen
in Fig. 3 共insets兲. The presence of inflammatory cells and
subsequent increase in the density of nuclei in the EDL tissue,
particularly at the graft boundary, should also affect the attenuation of the OCT signal with depth 共A-scan profile兲. The
expected relatively stronger scattering in the EDL is confirmed in Fig. 3 共insets兲, which shows that the average signal
strength in the EDL tissue is lower than in the neighboring TA
tissue, increasingly so at greater depth 关Fig. 3共c兲兴.
While the morphology evident in the en face OCT images
presented here correlates well with the corresponding histology, several issues must be resolved for the technique to be a
useful replacement for conventional histology. New regenerating myofibers, which are of particular interest in the study
of inflammation and regeneration in muscle tissue, could not
be readily identified in the OCT images. Typically, activated
myoblasts have fused to form myotubes and new myofibers
are visible at the edge of the graft by day 5, becoming evenly
distributed throughout5,9 the EDL around days 7 to 8. Discrimination in histology relies on resolving the nucleus and its
position with respect to the myofiber, as myotubes and regenerated myofibers have centrally located nuclei, while original
myofibers have peripherally located nuclei. Such resolution is
beyond the ability of the OCT setup used in this experiment
but may be attainable with ultrahigh-resolution OCT 共Ref. 32兲
and by utilizing recently published resolution enhancement
methods.35 The distinction between myofiber types may also
be possible through examination of the variation in the
A-scans, but more work is required to establish this. En face
OCT images could not be used to readily identify the necrotic
core within the grafted EDL tissue and, therefore, could not be
used to assess the percentage of this necrotic tissue relative to
the total graft area, an important measure that is used to assess
the effectiveness of drugs aimed at reducing inflammation.5
Future in vivo 3D-OCT imaging of mouse skeletal muscle
tissue must contend with the anatomical orientation of muscle
tissue and the presence of overlying skin tissue. Beyond improvements to the resolution afforded by ultrahigh
resolution32 and high speed,36,37 there are other potential strategies to facilitate in vivo imaging. Optical clearing agents
could potentially be used to increase the imaging depth of
OCT through the skin and cutis to the underlying muscle tissue of interest.38 Less desirably, direct access to muscle could
be made via surgical incision15 or needle incision.39,40
Journal of Biomedical Optics
5
Conclusion
The findings here demonstrate the promise of 3D-OCT as a
tool for the study of the morphology of mouse skeletal
muscle. In the whole-muscle autograft model of damage and
inflammation, OCT clearly shows the boundary between the
tissues of the grafted and host muscles. Further improvements
to enable the distinction between muscle fiber types 共original
undamaged versus regenerated兲 and the detection of the necrotic core, however, are required. Our preliminary results
suggest that OCT may have a future role in the morphological
assessment of the pathology of mouse skeletal muscle with
applications in the study of muscular dystrophy.
Acknowledgments
The authors thank Marilyn Davies for technical assistance
with both the whole-muscle autograft surgery and skeletal
muscle histology, James Ridgley for his contribution to the
histological digital imaging, Thea Shavlakadze for useful discussions on whole-muscle autograft biology, and Timothy
Hillman for preparation of Fig. 1共a兲.
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Vol. 13共1兲
_____________________________________________________________Appendix 1
Appendix 1 – Additional dietary intervention studies.
In addition to the work presented in Chapter 6, dietary interventions involving female
mice, mdx litters, and an Omega-3 enriched diet were also conducted. Due to fatal
flaws in experimental design and diet manufacture each of these studies was
terminated and data were not analysed in full. Each experiment is discussed briefly
below.
Custom diets: All custom diets were manufactured by Mr Warren Potts at Specialty
Feeds Glen Forrest WA.
A) Standard meat free rat and mouse chow – cereal based – 19% protein, 5% fat.
B) AIN93g – semi pure control – 19% protein, 7% fat.
C) High fat – semi pure – 19% protein, 16% fat.
D) High protein – semi pure – 50% protein, 7% fat.
E) Low protein – semi pure – 8% protein, 7% fat.
F) Omega 3 enriched – semi pure – increase Omega -3 content of AIN93G diet up to 7%
(replace canola oil with fish oil).
1) Dietary interventions in female mice:
Dietary intervention studies were initially conducted on female control C57 and
dystrophic mdx mice. Customised diets were fed to female mice from 8-24 weeks of
age: before starting the custom diet at 8 weeks of age all mice were fed a standard
mouse chow diet. There was no significant change in body weight of C57 or mdx
female mice after consumption of the 4 custom diets for 16 weeks. At the time of
sacrifice (24 weeks of age) there were no significant differences in the absolute body
weight (Figure A1) or retroperitoneal fat pad weight between any groups of mice
(Figure A2), however dystrophic mdx mice were significantly heavier than control C57
mice (Figure A1). Similar parallel studies that were being conducted on male mice
showed numerous significant differences in body weight and body composition, and
were therefore pursued in detail (Chapter 6). Due to time constraints detailed analysis
of dietary interventions was only conducted on male mice and the female dietary
intervention studies were not continued (e.g. extent of dystropathology was not
analysed), nor was the difference in gender response examined in detail. The major
differences in response to dietary interventions may be due to sex or less likely, to
differences in the batches of custom diets used for the 2 studies.
I
_____________________________________________________________Appendix 1
Final body weight - 24 weeks (g)
30
25
HF
20
HP
15
LP
AIN93g
10
Chow
5
0
C57Bl/10
mdx
Figure A1. Final body weight (g) of 24 week old C57Bl/10 and mdx female mice. N=8 mice per group.
Bars represent s.e.m.
Std fat pad weight (g fat / g bw)
0.05
0.04
HF
0.03
HP
LP
0.02
AIN93g
Chow
0.01
0
C57Bl/10
mdx
Figure A2. Standardised retroperitoneal fat pad weight (g fat / g bw) of 24 week old C57Bl/10 and mdx
female mice. N=8 mice per group. Bars represent s.e.m.
2) Dietary interventions in mdx litters:
Dietary interventions were conducted in young mdx mice in attempt to reduce the
acute onset of dystropathology around 3-4 weeks of age. Dietary interventions were
administered in two different ways A) In utero – postnatal day 28, B) postnatal day 15
– 28.
In utero dietary interventions consisted of feeding a custom diet to a female mdx
mouse 2 weeks before mating/conception. The major problem with this experiment
was that litter size was not standardised. Dystrophic mdx mice have a high neo-natal
death rate and litters are quite variable in size. When studying growth and
development of neo-natal mice it is important to standardise litter size as all neo-natal
mice depend on milk provided by the mother for survival and growth (e.g. smaller
litter size = larger pups). The number of mice used in this study (based on 2 litters per
sample group) is summarised in Table 1. Another major problem with this early study
was that all work was conducted using dystrophic mdx mice as mothers: as our
knowledge of the metabolic abnormalities in dystrophic mdx mice increased (Chapter
6 & 7) it became apparent that this work should have been conducted using neo-natal
mdx mice cross-fostered onto non-dystrophic mothers at birth. This would have
eliminated any complications resulting from dystrophic mdx mice being energy
deficient themselves and thus having limited milk producing capacity.
II
_____________________________________________________________Appendix 1
Group / Diet
In-utero – d28
d15 – d28
High fat
N=7 (3, 4)
N=11 (5, 6)
High protein
N=6 (3, 3)
N=8 (6, 2)
Omega-3
N= 9 (4, 5)
N=8 (5, 3)
AIN93g
N=13 (7, 6)
N=11 (5, 6)
Chow
N= 9 (5, 4)
N= 8 (4, 4)
Table 1. Sample group size for dietary interventions in young mdx mice (size of individual litters is shown
in brackets).
There were no significant differences in body weight after dietary interventions in 28
day old mdx mice (Figure A3). This result is heavily influenced by litter size and
different results may have been seen if litters were standardised to a specific number.
Final body weight - 28 day old mice (g)
16
12
HF
HP
8
Omega-3
AIN93g
Chow
4
0
In-Utero
d15-d28
Figure A3). Final body weight (g) for 28 day old mdx mice after dietary intervention. For N, see Table 1.
Bars represent s.e.m.
The extent of dystropathology was assessed histologically on H&E stained muscle
sections. There were no significant differences in myofibre necrosis (Figure A4 - A) or
myofibre regeneration (Figure A4 – B) in the quadriceps muscle of 28 day old mdx mice
after dietary interventions from 15 – 28 days of age. Similar results were seen in the TA
muscle and after consuming the dietary interventions from In Utero – d28. In summary
this was a poorly designed experiment and would need to be completely re-designed
in order to achieve sound scientific results.
III
_____________________________________________________________Appendix 1
Myofibre necrosis (Quad % CSA)
A
16
HF
12
HP
Omega-3
AIN93g
8
Chow
4
0
28 day old mdx mice
Myofibre regeneration (Quad % CSA)
25
B
20
HF
HP
15
Omega-3
AIN93g
10
Chow
5
0
28 day old mdx mice
Figure A4). Myofibre necrosis and regeneration (quadriceps) in 28 day old mdx mice, after consumption
of a dietary intervention from 15 – 28 days of age. A) Necrosis, B) Regeneration. For N, see table 1. Bars
represent s.e.m.
1) Omega-3 enriched diet: An omega-3 enriched diet was fed to dystrophic mdx mice
in attempt to reduce inflammation and thus reduce the extent of dystropathology. The
omega-3 enriched diet was fed to male mdx mice (8-12 weeks +/- voluntary exercise, 824 weeks and 24-40 weeks). Half way through conducting this study on sedentary
adult mice we were notified by the manufacturer that there was a major problem with
the diet composition (failure to add vitamin B), which would result in significant weight
loss in the mice, and that all experiments should cease. This resulted in the loss of 12
weeks of experimental work.
This study was repeated and upon sampling mice that had consumed the omega-3
enriched diet for 16 weeks a large number of mice showed significant weight loss (or
failure to gain weight) and had inflammed and gaseous stomach/bowels. No detailed
analysis of dystropathology was conducted on these mice. Attempts to again repeat
this study encountered major problems with finding a reliable and consistent source of
omega-3 rich fish oil, the oxidation of omega-3 fatty acids in the manufacturing
process, and further oxidation of omega-3 fatty acids in the autoclaving process. Thus
this dietary intervention was subsequently abandoned at the time.
Additional points of interest: There are a few additional points of interest to come out
of these failed dietary intervention studies.
IV
_____________________________________________________________Appendix 1
1) The neo-natal death rate of dystrophic mdx mice is very high (approximately 35%). If
mdx mothers and their new pups are left untouched (e.g cage unchanged and pups not
handled) for at least 10 days after birth, the neo-natal death rate is significantly
reduced to approximately 10%. Based on observations of approximately 50 pups.
2) The litter size of mdx pups is increased on the AIN93G semi-pure diet (19% protein,
7% fat) in comparison to mice fed the standard (cereal based) rat and mouse chow
(19% protein, 5% fat). The average litter size for AIN93G diet is 6.7 pups (n=4 litters)
and the average litter size for standard chow diet is 4.1 (n=4 litters).
3) Both female and male dystrophic mdx mice are significantly heavier than control
C57Bl/10 mice at 8 weeks of age (Figure A5) (that is mdx mice undergo significant
muscle hypertrophy before 8 weeks of age).
mdx female
C57 female
A
25
Bodyweight (g)
20
*
15
10
5
0
n=36
n=36
8 week old mice
mdx male
C57 male
B
Bodyweight (g)
30
25
*
20
15
10
5
0
n=24
n=24
8 week old mice
Figure A5. Absolute body weight (g) of 8 week old mdx and C57Bl/10 mice. A) Female mice, B) Male
mice. Bars represent s.e.m. * indicates significant difference between strains (P<0.05). N indicated on
each graph.
V