UMEÅ UNIVERSITY MEDICAL DISSERTATIONS
New Series No. 1342 ISSN 0346-6612 ISBN 978-91-7264-975-0
NEUROPROTECTION AND AXONAL
REGENERATION AFTER PERIPHERAL
NERVE INJURY
DAG WELIN
Department of Integrative Medical Biology
Section for Anatomy
Department of Surgical and Perioperative Sciences
Section for Hand & Plastic Surgery
Umeå University, Sweden, 2010
Responsible publisher under Swedish law: the Dean of the Medical faculty
Copyright©Dag Welin
ISBN: 978-91-7264-975-0
ISSN: 0346-6612
Cover: The peripheral nervous system:by Theodor de Bry 1621
Printed by: Print and Media, Umeå, Sweden 2010
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TABLE OF CONTENTS
ORIGINAL PAPERS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
ABBREVIATIONS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Clinical background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Classification of nerve injuries . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Factors influencing functional recovery after peripheral nerve injury . . . . . . . . .
Degeneration and regeneration after nerve injury. . . . . . . . . . . . . . . . . . . . . . . . .
Surgical repair after peripheral nerve injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Neuroprotection after peripheral nerve injury. . . . . . . . . . . . . . . . . . . . . . . . . . . .
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SPECIFIC AIMS. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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MATERIALS AND METHODS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Experimental animals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Labeling with retrograde fluorescent tracers. . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Sciatic nerve injury and repair . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Ventral root rhizotomy and avulsion. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Experimental groups . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
N-acetyl-cysteine treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Tissue processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Neuronal counts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Laser microdissection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
RNA isolation and quantitative RT-PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Image processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Statistical analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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RESULTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Peripheral nerve injury induces delayed loss of cutaneous sensory neurons . . . .
Spinal motoneurons degenerate after ventral rhizotomy and avulsion . . . . . . . . .
Nerve repair improves survival of axotomized sensory neurons . . . . . . . . . . . . .
Regeneration of sensory neurons and motoneurons after nerve repair . . . . . . . . .
Regenerating sensory neurons up-regulate peripherin mRNA expression . . . . . .
Sensory neurons down-regulate expression of NF-H mRNA after injury . . . . . .
Regenerating sensory neurons up-regulate ATF3 mRNA expression . . . . . . . . .
N-acetyl-cysteine promotes survival of spinal motoneurons . . . . . . . . . . . . . . . .
N-acetyl-cysteine promotes survival of cutaneous sensory DRG neurons . . . . . .
Effects of N-acetyl-cysteine and nerve grafting on neuronal regeneration . . . . .
Effect of N-acetyl-cysteine and nerve grafting on axonal sprouting . . . . . . . . . .
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Neuroprotection and axonal regeneration after peripheral nerve injury
DISCUSSION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Fluorescent retrograde tracers for neuronal quantification . . . . . . . . . . . . . . . . . .
Mechanism of retrograde cell death after peripheral nerve injury . . . . . . . . . . . .
Response of spinal motoneurons to distal and proximal axotomy . . . . . . . . . . . .
Sensory neurons after peripheral nerve injury . . . . . . . . . . . . . . . . . . . . . . . . . . .
Neuroprotective effect of nerve grafting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Regulation of peripherin and ATF3 genes in primary sensory neurons . . . . . . . .
N-acetyl-cysteine supports neuronal survival . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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CLINICAL IMPLICATIONS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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CONCLUSIONS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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ACKNOWLEDGEMENTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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REFERENCES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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PAPERS I-IV. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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ABSTRACT
Following microsurgical reconstruction of injured peripheral nerves, severed axons are able to
undergo spontaneous regeneration. However, the functional result is always unsatisfactory
with poor sensory recovery and reduced motor function. One contributing factor is the
retrograde neuronal death which occurs in the dorsal root ganglia (DRG) and in the spinal
cord. An additional clinical problem is the loss of nerve tissue that often occurs in the trauma
zone and which requires “bridges” to reconnect the injured nerve ends. The present thesis
investigates the extent of retrograde degeneration in spinal motoneurons and cutaneous and
muscular afferent DRG neurons after permanent axotomy and following treatment with Nacetyl-cysteine (NAC). In addition, it examines the survival and growth-promoting effects of
nerve reconstructions performed by primary repair and peripheral nerve grafting in
combination with NAC treatment.
In adult rats, cutaneous sural and muscular medial gastrocnemius DRG neurons and spinal
motoneurons were retrogradely labeled with fluorescent tracers from the homonymous
transected nerves. Survival of labeled neurons was assessed at different time points after
nerve transection, ventral root avulsion and ventral rhizotomy. Axonal regeneration was
evaluated using fluorescent tracers after sciatic axotomy and immediate nerve repair.
Intraperitoneal or intrathecal treatment with NAC was initiated immediately after nerve injury
or was delayed for 1-2 weeks.
Counts of labeled gastrocnemius DRG neurons did not reveal any significant retrograde cell
death after nerve transection. Sural axotomy induced a delayed loss of DRG cells which
amounted to 43-48% at 8-24 weeks postoperatively. Proximal transection of the sciatic nerve
at 1 week after initial axonal injury did not further increase retrograde DRG degeneration, nor
did it affect survival of corresponding motoneurons. In contrast, rhizotomy and ventral root
avulsion induced marked 26-53% cell loss among spinal motoneurons. Primary repair or
peripheral nerve grafting supported regeneration of 53-60% of the motoneurons and 47-49%
of the muscular gastrocnemius DRG neurons at 13 weeks postoperatively. For the cutaneous
sural DRG neurons, primary repair or peripheral nerve grafting increased survival by 19-30%
and promoted regeneration of 46-66% of the cells. Regenerating sural and medial
gastrocnemius DRG neurons upregulate transcription of peripherin and activating
transcription factor 3. The gene expression of the structural neurofilament proteins of high
molecular weight was significantly downregulated following injury in both regenerating and
non-regenerating sensory neurons. Treatment with NAC was neuroprotective for spinal
motoneurons after ventral rhizotomy and avulsion, and sural DRG neurons after sciatic nerve
injury. However, combined treatment with nerve graft and NAC had significant additive
effect on neuronal survival and also increased the number of sensory neurons regenerating
across the graft. In contrast, NAC treatment neither affected the number of regenerating
motoneurons nor the number of myelinated axons in the nerve graft and in the distal nerve
stump.
In summary, the present results demonstrate that cutaneous sural sensory neurons are more
sensitive to peripheral nerve injury than muscular gastrocnemius DRG cells. Moreover, the
retrograde loss of cutaneous DRG cells taking place despite immediate nerve repair would
still limit recovery of cutaneous sensory functions. Experimental data also show that NAC
provides a highly significant degree of neuroprotection in animal models of adult nerve injury
and could be combined with nerve grafting to further attenuate retrograde neuronal death and
to promote functional regeneration.
Key words: Dorsal root ganglion; Motoneuron; Axonal reaction; Peripheral nerve graft; Nerve
regeneration; N-acetylcysteine.
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Neuroprotection and axonal regeneration after peripheral nerve injury
ORIGINAL PAPERS
This thesis is based on the following papers which are referred in the text by Roman
numerals.
I.
Welin, D., Novikova, L.N., Wiberg, M., Kellerth, J-O. and Novikov, L.N.
Survival and regeneration of cutaneous and muscular afferent neurons after
peripheral nerve injury in adult rats. Experimental Brain Research, 186,
315-323, 2008.
II.
Reid, A.J., Welin, D., Wiberg, M., Terenghi, G., and Novikov, L.N.
Peripherin and ATF3 genes are differentially regulated in regenerating and
non-regenerating primary sensory neurons. Brain Research, 1310, 1-7,
2010.
III.
Zhang, C-G., Welin, D., Novikov, L., Kellerth, J-O., Wiberg, M. and Hart,
A.M. Motorneuron protection by N-acetyl-cysteine after ventral root
avulsion and ventral rhizotomy. British Journal of Plastic Surgery, 58, 765773, 2005.
IV.
Welin, D., Novikova, L.N., Wiberg, M., Kellerth, J-O. and Novikov, L.N.
Effects of N-acetyl-cysteine on the survival and regeneration of sural
sensory neurons in adult rats. Brain Research, 1287, 58-66, 2009.
All published papers are reproduced with the permission of the copyright holders.
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Dag Welin
ABBREVIATIONS
ATF3
DRG
FB
FG
FR
i.m
i.p
i.t
L4/L5/L6
MG
NAC
NF-H
NG
PBS
PFA
ROS
SC
S.E.M
SUR
TB
Activating transcription factor 3
Dorsal root ganglion
Fluorescent tracer Fast Blue
Fluorescent tracer Fluoro-Gold
Fluorescent tracer Fluoro-Ruby
Intramuscular
Intraperitoneal
Intrathecal
Lumbar segments 4/5/6
Medial gastrocnemius nerve
N-acetyl-cysteine
Neurofilament proteins of high molecular weight
Peripheral nerve graft
Phosphate buffer solution
Paraformaldehyde
Reactive oxygen species
Spinal cord
Standard error of the mean
Sural nerve
Fluorescent tracer True Blue
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Neuroprotection and axonal regeneration after peripheral nerve injury
INTRODUCTION
Clinical background
Peripheral nerve lacerations are common injuries and often cause long lasting
disability (Jaquet et al. 2001) due to pain, paralyzed muscles and loss of adequate
sensory feedback from the nerve receptors in the target organs such as skin, joints
and muscles (Lundborg and Rosen 2007). Normal function will not be regained even
if the nerve is repaired (Lundborg 2000a). Nerve injuries typically affect young
adults and the majority of injuries is domestic or occupational accidents with glass,
knifes of machinery (McAllister et al. 1996; Rosberg et al. 2005) but road traffic
accidents, iatrogenic injuries, assault, self-inflicted injuries are other known causes.
The severity of the injuries varies from minor, such as digital nerve injury to major,
such as brachial plexus injury. Nerve injuries affect both the individual patient and
the society, for example, an injury to the ulnar or median nerve in the forearm has
major social consequences (Jaquet et al. 2001) and results in a sick leave for a
median time of 157 or 273 days, respectively. The total cost per patient with ulnar or
median nerve injury has been estimated to be EUR 51 238 and EUR 31 186,
respectively (Rosberg et al. 2005).
Classification of nerve injuries
The typical peripheral nerve is composed by efferent motor axons originated from
spinal motoneurons and afferent sensory axons from dorsal root ganglion (DRG)
neurons. An injury to the peripheral nerve, therefore, will affect both motor and
sensory function and induce retrograde (axon) reaction in spinal cord and DRGs.
There are different types of nerve injury and the most common include nerve
transection, compression, crush, traction and avulsion.
To facilitate clinical work, Seddon et al. (1943) classified nerve injuries into three
main groups: neurapraxia, axonotmesis and neurotmesis. Neurapraxia is the mildest
type of nerve injury and is often associated with transient ischemia without physical
disruption of the axons or myelin sheath. Although this form of nerve injury blocks
nerve impulse transmission, it does not require any surgical intervention and
recovery usually occurs within a few weeks. Axonotmesis refers to the disruption of
several axons and myelin sheath but without significant damage to the connective
tissue layers (Seddon et al. 1943). Axons distal to the injury degenerate but there is
possibility for spontaneous regeneration without surgical treatment. Neurotmesis is
usually associated with partial or complete severance of the nerve with disruption of
the axon, myelin sheath and the connective tissue elements. It is the most severe
type of nerve injury and no functional recovery is possible without surgical nerve
reconstruction. Another common grading system is the Sunderland’s classification
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which contains five degrees of severity of the nerve injury. However, it requires
histological analysis of the injured nerve (Birch 2005).
There are different techniques used to repair injured peripheral nerves. For example,
“epineural repair” indicates that micro sutures were placed in the epineurium to
align the nerve ends whereas in “fascicular repair” individual nerve fascicles are
repaired. Nerve reconstructions could be also subdivided into “primary repair” if it
is performed within 48 hours after injury, “early secondary repair” if performed
between 2 days and 6 weeks and “late secondary repair” (Green 1998). In this thesis,
all nerve injuries are complete transections of the peripheral nerves and, therefore,
can be classified as “neurotmesis”. For nerve reconstruction and grafting only
primary repairs with epineural suturing technique were used.
Factors influencing functional recovery after peripheral nerve injury
The current management of a lacerated peripheral nerve is based on either direct
microsurgical repair where the cut ends are approximated with sutures, or
autologous nerve grafting where an existing defect is bridged with a harvested donor
nerve. Even if the injured nerve is meticulously repaired the outcome is often
unpredictable and disappointing (Jaquet et al. 2001; Ruijs et al. 2005; Lundborg and
Rosen 2007). Several factors have been identified that influence the outcome after
nerve repair. Young age is a main factor for recovery and could be explained by
shorter regeneration distance, greater regeneration potential and brain plasticity
(Lundborg 2000a; Ruijs et al. 2005). It has also been shown that delay until surgical
repair is unfavorable for recovery (Ruijs et al. 2005) which is suggested to result
from neuronal loss, distal nerve stump fibrosis and Schwann cell atrophy (Gordon et
al. 2003; Lundborg and Rosen 2007). The proximity of the injury is a significant
predictor for motor recovery due to the long regeneration distance leading to
irreversible muscle atrophy before target reinnervation (Ruijs et al. 2005). Good
cooperation and motivation of the patient, cognitive capacity and specialized hand
therapy are other factors contributing to the outcome after nerve repair (Ruijs et al.
2005; Lundborg and Rosen 2007). A vital component for normal motor control is
adequate proprioceptive feedback (Westling and Johansson 1984). Therefore,
sensory regeneration is important for the recovery of motor function. Future
modifications in surgical techniques are not likely to significantly improve the
outcome after a nerve injury. Thus, attention has been focused on the neurobiology
following nerve injury and how various components of axon reaction and Wallerian
degeneration can be modulated.
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Neuroprotection and axonal regeneration after peripheral nerve injury
Degeneration and regeneration after nerve injury
Axon reaction
After nerve injury signaling through injury-induced discharge of axonal potentials,
interruption of target-derived factors and retrograde injury signals transported from
the site of injury to the cell body enables the neuron to respond to the trauma.
Proposed injury signals of importance consist of microtubule-dependant axonal
transport of mitogen-activated protein kinases (MAPK) including Erk and JNK, and
local release of cytokines LIF, IL-6 and CNTF leading to activation of the JAKSTAT pathway (Abe and Cavalli 2008). Axotomized neurons respond by upregulation of regeneration-associated genes in association with conversion of the
neuron from a transmitting to a growth state (Boyd and Gordon 2003).
Nerve transection produces morphological changes in the neuronal perikarya known
as chromatolysis or axon reaction. The changes include swelling of the cell body,
nucleolar enlargement, displacement of the nucleus to the periphery and dissolution
of Nissl bodies (Kreutzberg GW 1995). Chromatolysis is explained as a change in
the neurons metabolism aiming at an increased regenerative potential but may also
be a sign of severe trauma with loss of a large axoplasmic volume. Axon reaction
could result in both neuronal survival and regeneration or neuronal death (Lundborg
2000a). The principal determinant of the extent of neuronal death after axotomy
seems to be the loss of target-derived neurotrophic factors (Terenghi 1999).
For many years it was recognized that experimental nerve injuries result in a loss of
primary sensory neurons and more recently it was confirmed by direct observation
of DNA fragmentation in vivo (Groves et al. 1997; Oliveira et al. 1997; Hart et al.
2002a). The extent of the loss ranges from 7 to 51% (Rich et al. 1989; Liss et al.
1994; Liss et al. 1996; Groves et al. 1997; Tandrup et al. 2000; Hart et al. 2002a; Ma
et al. 2003; Jivan et al. 2006), depending on the experimental model used. Since the
first prerequisite for axonal regeneration is the survival of the neuron following
injury, it is likely that this neuronal cell death is of great importance for the outcome
of the axonal regeneration and target organ reinnervation (Fu and Gordon 1997).
Motoneurons seem to be more resistant to peripheral axotomy (Novikova et al.
1997) however, 20 to 30% of motoneurons die after proximal C7 axotomy or lumbar
rhizotomy (Gu et al. 1997; Ma et al. 2001; Zhang et al. 2005; Jivan et al. 2006), and
more than 50% degenerate after ventral root avulsion at 4-8 weeks after injury
(Koliatsos et al. 1994; Novikov et al. 1995).
Posttraumatic neuronal death is believed to be an apoptotic response comprising
pathways and mediators that are activated in various neuropathological conditions
(Becker and Bonni 2004). It is possible that different cell types encode different
apoptotic and survival genes, and also that cells may activate different apoptotic
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pathways in response to different external stimuli (Pettmann and Henderson 1998).
Following axotomy, apoptosis might be initiated by alterations in electrical activity,
neurotoxic inflammatory products and loss of target derived neurotrophic support
(Ambron and Walters 1996; Fu and Gordon 1997). Central components in the
present context are the mitochondria. Pro-apoptotic mediator Bax and pro-survival
Bcl-2 interact at the level of the mitochondria and it has been suggested that their
ratio determines the fate of the cell by governing mitochondrial outer-membrane
permeability (Gillardon et al. 1996). Bcl-2 has been shown to heterodimerize with
Bax which counteracts the death-repressor activity of Bcl-2. When Bax exerts
dominance over Bcl-2, pores are formed in the mitochondrial membrane which
allows release of apoptotic molecules (cytochrome-c) into the cytosol, thus
triggering the execution caspase-cascade. This culminates in the activation of
caspase-3, an ‘effector caspase’ which is directly involved in the proteolytic action
on cells, e.g. digestion of structural proteins in the cytoplasm and degradation of
chromosomal DNA (Becker and Bonni 2004). Recently, a different response in
regulation of Bax, Bcl-2 and caspase-3 has been indentified in cutaneous and
muscular subpopulations of sensory DRG neurons after injury. The Bcl-2:Bax ratio
in sensory DRG neurons projecting to muscles increases whilst the ratio for sensory
DRG neurons projecting to the skin decreases. Simultaneously the levels of caspase3 have been markedly down-regulated in the muscular afferent neurons and
progressively up-regulated in the cutaneous afferent neurons (Reid et al. 2009).
Wallerian degeneration
After a nerve transection, a well-defined cascade of cellular changes occurs in the
distal nerve segment and to the first node of Ranvier in the proximal nerve stump.
The process is called Wallerian degeneration and involves degeneration of all
affected axons, degradation of their myelin sheaths and invasion of macrophages to
remove debris. Wallerian degeneration creates a favorable microenviroment for
regeneration of surviving neurons (Navarro et al. 2007). The Schwann cells divide
by mitosis and line up within each basal lamina tube to form the bands of Büngner,
providing guidance for regenerating nerve fibers (Frostick et al. 1998; Terenghi
1999; Hall 2001). During the initial phase of regeneration an axon in the proximal
stump sends out multiple sprouts, averaging five axons, which regenerate into
multiple endoneurial tubes in the distal stump, potentially leading to complex
mismatch in reinnervation of denervated targets. If the axons lack a supportive
structure they will, together with connective tissue, form a neuroma. After target
reinnervation all axons from a neuron, but one, die-back (Gordon et al. 2003).
Regenerating axons in the distal nerve stump enlarge in diameter and reach normal
size when they make connections with the target organs (Sulaiman and Gordon
2000).
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Neuroprotection and axonal regeneration after peripheral nerve injury
Schwann cells and peripheral nerve injury
The Schwann cells promote survival and regeneration by increased synthesis of
surface cell adhesion molecules (CAMs), by providing a basement membrane with
extracellular matrix proteins laminin and fibronectin, and by producing various
growth factors (Frostick et al. 1998; Lundborg 2000a; Lundborg and Rosen 2007).
After injury, Schwann cells in the distal stump proliferate and convert their function
from myelination of electrically active axons to growth support for regenerating
nerve fibers (Fu and Gordon 1997). Optimal expression of growth-supportive
molecules by Schwann cells is associated with macrophages infiltrating the distal
stump from peripheral circulation (Gordon et al. 2003). The presence of viable
Schwann cells in the distal nerve stump is essential for regeneration after nerve
injury (Fu and Gordon 1997). It has been demonstrated that prolonged axotomy
leads to rather limited regeneration due to atrophy of Schwann cells and decline in
expression of neurotrophic factors (Sulaiman and Gordon 2000; Hoke et al. 2002;
Gordon et al. 2003; Jivan et al. 2006).
Neurotrophic factors and their receptors after peripheral nerve injury
It is mainly agreed that neurons rely on neurotrophic support for survival. Trophic
factors can be divided into neurotrophins (NGF, BDNF, NT-3 and NT-4/5),
neuropoetic cytokines (Il-6, LIF and CNTF) and the Glial cell-lined derived
neurotrophic factor family (GDNF, Neurturin, Persephin and Artemin). In the
uninjured neuron the trophic factors are produced by the target organ and delivered
to the neuron by retrograde axonal transport. Neurons sensitive to neurotrophins
express the high-affinity tyrosine kinase (Trk) receptors and the low-affinity p75
receptor. Neurons sensitive to the GDNF family express the GFRα receptors and the
common signal transduction unit, ret receptor. Neuropoetic cytokines binds to
specific receptors and the common signal transduction receptor subunit, gp130
(Boyd and Gordon 2003). Different subpopulations of neurons have been identified
and characterized by overlapping but yet distinct trophic requirements (McMahon et
al. 1994). A large proportion of the primary sensory afferents expresses the TrkA
receptors for NGF, whereas trkC receptors for NT-3 are mainly expressed by large
diameter proprioceptive neurons (Lindsay 1996). Motoneurons express TrkB and
TrkC receptors and bind BDNF, NT-4/5 and NT-3. A part of the neurons in the
dorsal root ganglion lacks Trk receptors and express GFRα receptors which bind
members of the GDNF family. After a sciatic crush, the Schwann cells in the distal
end up-regulate their expression of neurotrophins NGF, BDNF and NT-4/5 and
trophic factors GDNF, LIF and IL-6. Motoneurons up-regulate trkB and GFRα1
receptors. Neither intact nor injured motoneurons express GDNF (Boyd and Gordon
2003). In primary sensory afferents all Trk receptors are down-regulated after
axotomy (Bergman et al. 1999) but the GFRα1 receptor is dramatically up-regulated
in large-diameter sensory afferents (Bennett et al. 2000). It is possible that surviving
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peripheral axotomy sensory and motor neurons express receptors which match the
profile of neurotrophic factors produced by Schwann cells in the distal nerve stump.
Regenerative response after axonal injury
The functional goal of regeneration is to replace the distal nerve segment lost after
injury, to reinnervate the target organs and to restitute their function. A peripheral
nerve injury activates a complex molecular response in the neuron. The biochemical
changes develop within hours after axotomy, triggered by the signaling mechanism
mentioned previously, and is overall thought to induce a decreased synthesis of
products for neurotransmission and increased synthesis of growth associated
proteins and components of the membrane. Signaling through the axotomy-activated
kinases lead to up-regulation or activation of several transcription factors including
c-Jun, cAMP responsive element binding protein (CREB), STAT-3, Akt and
Nuclear Factor kB (NFkB). This upregulation leads to changes in gene expression in
a large number of genes, many of which the function is unknown but involves
changes in transcription factors, cytoskeletal proteins, cell adhesion and guidance
molecules, trophic factors and receptors, cytokines, neuropeptides and
neurotransmitter synthesizing enzymes, ion channels and membrane transporters
(Navarro et al. 2007).
Recently, the roles of neuronal intermediate filaments and activating transcription
factor 3 (ATF3) in the regenerative response have been examined. Neuronal
intermediate filaments are crucial components of the neuronal cytoskeleton
comprising the neurofilament triplet proteins of low (68 kDa), medium (150 kDa),
and high (NF-H, 200 kDa) molecular weight, in addition to α-internexin (66 kDa)
and peripherin (57 kDa) (Helfand et al. 2004; Lariviere and Julien 2004). Peripherin
and NF-H define two distinct subpopulations of sensory DRG neurons (Fornaro et
al. 2008). It has been shown that after nerve injury sensory neurons in DRGs and
spinal motoneurons up-regulate peripherin (Oblinger et al. 1989; Troy et al. 1990;
Wong and Oblinger 1990; Terao et al. 2000) and down-regulate NF-H (Wong and
Oblinger 1990; Muma et al. 1990).
Surgical repair after peripheral nerve injury
The gap between the two ends of a divided nerve can vary from a few millimeters to
several centimeters depending on the type of injury and the timing of surgery. A
small nerve gap is traditionally repaired with primary nerve suturing. However, if
the peripheral nerve injury is associated with a significant tissue loss, bridging
strategies are needed to provide a physical substrate for axonal growth. Defects in
the peripheral nervous system is most commonly bridged by autologous nerve grafts
obtained from “less important” sensory nerves from the patient’s legs. So far, nerve
grafts have been superior to other substitutes since they provide appropriate
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Neuroprotection and axonal regeneration after peripheral nerve injury
alignment and contain the cellular constituents of normal peripheral nervous tissue.
The technique is not optimal, however, since the patient will suffer from loss of
sensation, scarring and sometimes pain in the donor region (Wiberg and Terenghi
2003). In case of dorsal and ventral root avulsion from the spinal cord, transfer of
previously uninjured nerves may be considered. If the distal nerve end is removed
from the muscle, direct implantation of the nerve back into the muscle, a so called
neurotisation, can be performed (Birch 2005). Retrograde neuronal degeneration
after peripheral nerve injury can experimentally be reduced by early nerve repair
(Ma et al. 2003; Jivan et al. 2006; Zhang et al. 2006) possibly by reestablishment of
endogenous neurotrophic support from the distal nerve stump (Boyd and Gordon
2003; Low et al. 2003). Clinical results from repair of the brachial plexus also
demonstrate that early nerve reconstruction provides better functional outcome when
compared to delayed nerve repair (Jivan et al. 2008).
Recent studies have also demonstrated that in contrast to sensory nerve autografting,
more efficient axonal regeneration could be obtained with motor nerve grafts
(Moradzadeh et al. 2008; Chu et al. 2008). Improved regeneration with motor
grafting may be a result of the nerve's Schwann cell basal lamina tube size
(Moradzadeh et al. 2008) or reflect different expression of neurotrophic factors from
Schwann cells in the sensory and motor nerve grafts (Hoke et al. 2006; Chu et al.
2008). However, the obvious problems from the donor region limit the potential
clinical use of grafts from motor nerves.
Neuroprotection after peripheral nerve injury
Several experimental studies have shown that permanent axotomy induces
significant but delayed retrograde cell death of sensory neurons projecting mainly to
skin and does not affect survival of sensory neurons projecting to muscle (Tandrup
et al. 2000; Hu and McLachlan 2003; Welin et al. 2008). Therefore, a therapeutic
window exists where immediate neuroprotective treatment could be initiated to
attenuate retrograde degeneration before nerve repair could be performed.
Neurotrophic factors
It is well known that neurotrophic factors regulate the development, maintenance
and function of the vertebrate nervous system. Studies in vitro have shown that
activation of PKA, Ras/PI-3K/Akt (PI-3K/Akt), or Ras/Raf/ MAPK/ERK
(MAPK/ERK) signaling pathway by neurotrophins and GDNF could promote both
neuronal survival and enhance neurite growth in neuronal populations (Hetman et al.
1999; Soler et al. 1999; Liot et al. 2004; Chierzi et al. 2005). In contrast, the JAKSTAT signaling pathways are activated by cytokines and can mediate regenerative
sensory axon growth (Liu and Snider 2001).The depletion of neurotrophic factors
from target organs and Schwann cells plays a significant role in the induction of
15
Dag Welin
retrograde degeneration (Boyd and Gordon 2003). Administration of exogenous
neurotrophic factors has shown to reduce neuronal loss in sensory neurons (Bennett
et al. 1998; Ljungberg et al. 1999) and motoneurons (Novikov et al. 1995).
Neurotrophic factors can also promote axonal regeneration (Boyd and Gordon 2002;
McKay et al. 2003) and counteract development of axon reaction (Boyd and Gordon
2003). However, the specificity of neurotrophic factors for neuronal subpopulations
(Terenghi 1999), unpredictable interactions (Novikova et al. 2000a) and side effects
(Martin et al. 1996; McArthur et al. 2000; Apfel 2002) makes clinical use difficult.
Antioxidants
During past decades, significant number of various neuroprotective agents has been
tested both in vitro and in vivo. For example, treatments with acetyl-L-carnitine
(Hart et al. 2002b; Wilson et al. 2007), the monoamine oxidase inhibitor deprenyl
(Hobbenaghi and Tiraihi 2003), the cytokine modulator linomide (Ekstrom et al.
1998) and thyroxine (Schenker et al. 2003) have been shown to protect sensory
DRG neurons from retrograde cell death .
It is well established that following injury to the nervous system, mitochondrial
impairment could lead to generation of reactive oxygen species and activation of
apoptotic cascades. Given the important role that oxidative stress plays in promoting
neuronal death, administration of antioxidants may be potentially attractive as
clinically applicable neuroprotective agents (Merenda and Bullock 2006). N-acetylcysteine (NAC), a thiol-containing compound, has a broad range of actions which
includes antioxidant activity, enhancement of intracellular glutathione levels,
inhibition of proliferation, and stimulation of transcription (Holdiness 1991; Yan
and Greene 1998; Arakawa and Ito 2007). NAC has been used in clinical practice
for many years as a mucolytic agent for treatment of congestive and obstructive lung
diseases and as the drug of choice for paracetamol intoxication. Recently it has been
shown that NAC can rescue sensory and motor neurons from retrograde
degeneration (Hart et al. 2004; Zhang et al. 2005; West et al. 2007b)
NAC has a broad range of actions potentially relevant to its neuroprotective effects.
Most of the results are from in vitro experiments and the mechanism of NAC
neuroprotective effect in vivo is largely unknown. NAC has a direct reductant and
free radical scavenging effects, interacting with reactive oxygen species (ROS) and
is neuroprotective in multiple neuronal models in vitro (Yan et al. 1995; Ferrari et
al. 1995). NAC can also enhance neuronal biosynthesis of glutathione (Dringen and
Hamprecht 1999), the principal renewable free radical scavenger within neurons
(Cooper and Kristal 1997; Heales et al. 1999). Depletion of intracellular, and
especially mitochondrial glutathione leads to neuronal death in vitro (Wullner et al.
1999). Increased glutathione levels by NAC prevented cytochrome c release from
the mitochondria, thereby preventing the apoptotic cascade (Kirkland and Franklin
2001). However, the neuroprotective effect of NAC has also been shown to be
16
Neuroprotection and axonal regeneration after peripheral nerve injury
glutathione-independent (Yan et al. 1995) and other antioxidants fail to mimic its
neuroprotective effect (Ferrari et al. 1995). It has been reported that NAC can signal
through the neurotrophic factor-activated Ras-ERK pathway and stress-activated
JNK pathways but does not activate the PI3-K/Akt survival pathway (Xia et al.
1995; Park et al. 1996; Yan and Greene 1998). These findings strongly suggest that
NAC can share with neurotrophic factors the capacity to maintain neuronal survival
and could be used as a neuroprotective agent to rescue sensory and motor neurons
after peripheral nerve injury.
17
Dag Welin
SPECIFIC AIMS
The aims of this study were:
•
To compare the retrograde degeneration and axonal regeneration of sensory
DRG neurons and spinal motoneurons projecting to different target organs
(Paper I).
•
To examine the gene expression of neuronal intermediate filaments and
activating transcription factor 3 (ATF-3) in cutaneous and muscular
sensory DRG neurons (Paper II).
•
To evaluate the neuroprotective effect of N-acetyl-cysteine treatment on
spinal motoneurons after ventral rhizotomy and ventral root avulsion
(Paper III).
•
To evaluate the neuroprotective and growth-promoting effects of N-acetylcysteine treatment and nerve grafting on sensory DRG neurons and spinal
motoneurons after peripheral nerve injury (Paper IV).
18
Neuroprotection and axonal regeneration after peripheral nerve injury
MATERIALS AND METHODS
Experimental animals
The experiments were performed on adult (8-12 weeks; n=228) female SpragueDawley rats (Möllegaard Breeding Center, Denmark). The animal care and
experimental procedures were carried out in accordance with the standards
established by the NIH Guide for Care and Use of Laboratory Animals (National
Institutes of Health Publications No. 86-23, revised 1985) and the European
Communities Council Directive (86/609/EEC). The study was approved by the
Northern Swedish Regional Committee of Ethics in Animal Experiments.
All surgical procedures were performed under general anesthesia with iterated
intraperitoneal injections using a mixture of ketamine (Ketalar, Parke-Davis) and
xylazine (Rompun, Bayer). Benzylpenicillin (Boehringer Ingelheim; 60 mg i.m.)
was given after each surgical procedure.
Labeling with retrograde fluorescent tracers
In order to study neuronal survival and regeneration after peripheral nerve injury,
DRG neurons and spinal motoneurons projecting to the sural or medial
gastrocnemius nerves were retrogradely labeled with fluorescent tracers (Novikova
et al. 1997). Under an operating microscope, the nerves were transected at the same
level in the popliteal fossa of the hind limb and the proximal nerve end was
introduced into a small polyethylene tube containing two microlitres of Fast Blue
(FB, 2% aqueous solution, EMS-Chemie GmbH, Germany; Papers I, II and IV) or
True Blue (TB, 2% aqueous solution, Molecular Probes; Paper III). The tube was
fixed to the surrounding muscles using Histoacryl® glue (B.Braun Surgical GmbH,
Germany) and sealed with a mixture of silicone grease and vaseline to prevent
leakage. Two hours later the tube was removed, the nerve rinsed in saline and the
wound closed in layers. Fast Blue and True Blue produces a very efficient retrograde
labeling of neurons at 1 week after tracer application and the staining remains
constant for at least 6 months (Novikova et al. 1997; Houle and Ye 1999). However,
since it has been reported that axotomized adult DRG neurons may start to die
already after 1 week (Hart et al. 2002a), we also used the fluorescent tracer FluoroGold (FG, 2% solution in saline, Fluorochrome) in experiments with short-term
survival (Paper I) since this dye produces very rapid neuronal labeling within 2-3
days (Novikova, unpublished observation) but gradually disappears from the labeled
neurons after 4 weeks (Novikova et al. 1997; Akhavan et al. 2006). In the
experiments dealing with the time course of retrograde degeneration (Paper I), the
sural and medial gastrocnemius nerves were labeled on different sides. To identify
sural and medial gastrocnemius neurons which had regenerated across the repair site
19
Dag Welin
and into the distal stump of the sciatic nerve the fluorescent tracer Fluoro-Ruby (FR,
10% solution in normal saline, Molecular Probes) was used (Papers I, II and IV).
Fast Blue tracer was also injected into the medial gastrocnemius muscle at four
different sites using a 10-µl Hamilton syringe (Hamilton, Switzerland) and into the
sural nerve using a fine-glass microelectrode (Paper II). To test the presence of nonspecific labeling, axotomized sural nerve was divided and capped proximal to the
site of Fast Blue administration to prevent dye uptake. In addition, contralateral
DRG were also examined. These control experiments confirmed that there was no
leakage of the tracer.
Sciatic nerve injury and repair
Sciatic nerve transection and repair were performed at 1 week after Fast Blue
labeling of the sural and medial gastrocnemius nerves. The sciatic nerve was
exposed via a muscle splitting incision and the wound held open with retractors,
taking care at all times not to traumatize the nerve. The surgery on the nerve was
performed using sterile micro instruments and an operating microscope (Zeiss, Carl
Zeiss, Germany). The nerve was divided at mid-thigh level. To produce permanent
axotomy, the proximal nerve stump was ligated with 6-0 Prolene, capped with
polyethylene tube or wrapped in Spongostan® (Johnson & Johnson Medicals, UK)
to prevent spontaneous regeneration. For nerve grafting, a 10 mm piece of the sciatic
nerve was excited, reversed and interposed between the nerve stumps. Primary
repair and nerve grafting of the divided sciatic nerve were performed using four 10/0
Ethilon® epineural sutures (S&T Marketing AG, Switzerland) and fibrin glue
(Tisseel®, Immuno, Vienna, Austria) to secure the graft into position. Skin and
fascia were closed with interrupted 3-0 silk veterinary sutures.
Ventral root rhizotomy and avulsion
One week after tracer application to the medial gastrocnemius nerve, a lumbo-sacral
laminectomy was performed and L5-L6 ventral roots of the left side were identified
and transected close to the corresponding DRGs (ventral rhizotomy). To perform
ventral root avulsion, the proximal stump of the root was grasped with jeweller’s
forceps and slowly pulled until it ruptured and came out in its entire length. The root
was found to regularly rupture at its site of exit from the spinal cord. Spinal roots
were covered with Spongostan® and the wound was closed in layers. Survival of
labeled motoneurons was assessed four week after injury (Novikov et al. 2000).
20
Neuroprotection and axonal regeneration after peripheral nerve injury
Experimental groups
Paper I
To study the time course of neuronal loss, three groups of animals with FG-labeled
nerves were killed at 3 days (n=6), 7 days (n=5) and 14 days (n=5). Five groups of
animals with FB-labeled nerves were killed at 1 week (n=5), 4 weeks (n=6), 8 weeks
(n=5), 13 weeks (n=5) and 24 weeks (n=6). To study nerve regeneration, three
groups of animals with FB-labeled medial gastrocnemius nerve (MG) and three
groups of animals with FB-labeled sural nerve (SUR) were used. One week after
retrograde FB-prelabeling of the SUR or MG nerves in the the popliteal fossa, a
sciatic axotomy was performed proximally in the thigh. In the first group (n=6 for
SUR, n=6 for MG) the sciatic nerve was capped during the entire survival period. In
the second group (n=5 for SUR, n=7 for MG), a primary suture of the sciatic nerve
was performed. In the third group (n=6 for SUR, n=5 for MG), the sciatic nerve was
repaired using a peripheral nerve graft. At 12 weeks after sciatic nerve transection
and repair, the nerve was again transected 10 mm distal to the repair site and the
retrograde tracer Fluoro-Ruby was applied to the proximal nerve stump(Jivan et al.
2006). To achieve optimal FR-labeling of the DRG neurons, the animals were left to
survive for one week before termination of the experiment.
Paper II
To study the expression of regeneration-associated genes after nerve injury and
repair, four groups of experimental animals were used: 1) control animals with FB
injection into medial gastrocnemius muscle (n=5), 2) control animals with FB
injection into sural nerve (n=5), 3) animals with FB-labeled medial gastrocnemius
neurons, sciatic nerve transection and immediate primary repair (n=5) and 4)
animals with FB-labeled sural neurons, sciatic nerve transaction and immediate
primary repair (n=5). At 8 weeks postoperatively the sciatic nerve was transected
10mm distal to the repair site and the proximal stump was labeled with Fluoro-Ruby
to reveal regenerating neurons.
Paper III
The medial gastrocnemius (MG) motorneuron pool was unilaterally prelabeled with
a retrograde tracer. The effect of NAC (given either intraperitoneally, or
intrathecally by infusion into the cerebrospinal space) on survival of those
motorneurons was then determined after ventral rhizotomy, or root avulsion. To
study the effects of delayed NAC treatment separate groups of animals were used.
Numbers of animals in each experimental group are detailed in Tables 1-3.
Paper IV
To study whether NAC could protect sural sensory neurons from retrograde cell
death, sciatic nerve was transected at 1 week after neurons were labeled with Fast
21
Dag Welin
Blue and treatment was initiated. The proximal stump was covered with
Spongostan® (Johnson & Johnson Medicals, UK) to prevent regeneration. Controls
(n=5) and treated animals (n=4) were allowed to survive for 8 weeks.
To study NAC effect on nerve regeneration, three groups of animals were used
(control, n=5; nerve grafting, n=6; nerve grafting with NAC treatment, n=8). One
week after Fast Blue application to the transected sural nerve, the sciatic nerve was
divided and repaired with peripheral nerve graft. In the treated group, NAC infusion
was initiated immediately after nerve repair. At 12 weeks postoperatively, sciatic
nerve was again transected 10 mm distal to the graft-nerve anastomosis and the
nerve stump was introduced into a polyethylene tube containing Fluoro-Ruby. The
animals were left to survive for 1 week before the termination of the experiment.
N-acetyl-cysteine treatment
The clinically available L-stereoisomer of N-acetyl-cysteine (NAC, Tika,
200mg/ml) was used in all experiments. In the experimental groups receiving
intraperitoneal injections, treatment was commenced immediately after the
experimental injury (1ml bolus). The rats were given injections once daily until
termination of the experiment. In the ventral root avulsion group of animals, the
dose tested was 150mg/kg/day, but after ventral rhizotomy, we used 150mg/kg/day
and 750mg/kg/day to evaluate any dose response relationship. In order to exclude
any possible toxic effect of NAC on motorneurons, treatment was also given to
control animals for 4 weeks (n=5) and 8 weeks (n=2). Normal controls included
non-injured rats labeled with FG for 5 weeks and with TB for 8 weeks.
In the experimental groups receiving intrathecal infusion, an Alzet 2002 osmotic
mini-pump (Alza Corp., CA, USA) containing NAC was implanted subcutaneously
in the neck and connected to a polyethylene catheter (Intermedic, PE-60, Clay
Adams, NJ) which was inserted into the lower lumbar subarachnoid space after L5L6 laminectomy (Novikov et al. 1997; Zhang et al. 2005). The catheter tip was
gently advanced rostral to the level of L3-L4 DRGs, fixed to the S1 vertebral bone
by Histoacryl® glue, and additionally secured by several sutures to the back
muscles. The implantation site was covered with Spongostan® and the wound
closed in separate layers. The mini-pump infusion speed corresponded to 2.4 mg/day
of NAC. At intervals of 14 days, the emptied mini-pump was replaced by a second
pump containing the same solution. The total time of intrathecal NAC infusion was
8 weeks.
To establish whether NAC would give neuroprotective benefit if treatment would be
commenced well after injury, four further groups underwent ventral root avulsion
and mini-pump implantation. Minipumps and catheters in the first two groups
contained either PBS (“avulsion + intrathecal vehicle”) or NAC (200 mg/ml,
22
Neuroprotection and axonal regeneration after peripheral nerve injury
“avulsion + intrathecal NAC, immediate treatment”), and the mini-pumps were
replaced after two weeks. The third group’s first minipump contained NAC
(200mg/ml), but its catheter was primed with PBS, and its length calibrated at 37oC
such that delivery of NAC into the subarachnoid space was delayed by one week.
The pump was replaced two weeks later to continue delivery of NAC (avulsion +
intrathecal NAC, 1-week delayed treatment). In the final group, the first mini-pump
contained PBS, but was replaced after one week by another pump with NAC
(200mg/ml), such that drug would reach the end of the catheter a week later,
beginning active treatment two weeks after injury (avulsion + intrathecal NAC, 2week delayed treatment). A group of non-injured rats, labeled for 7 days with TB,
served as baseline controls.
Tissue processing
At the end of the survival period, the animals were given an intraperitoneal overdose
of sodium pentobarbital (240mg/kg, Apoteksbolaget, Sweden) and transcardially
perfused with Tyrode’s solution (37ºC) followed by 4% paraformaldehyde (PFA,
pH 7.4). Spinal cord segments L4-L6 and homonymous DRGs were harvested and
post-fixed in 4% PFA overnight. The spinal cord segments were cut in serial 50 µm
thick parasagittal sections on a vibratome (Leica Instruments, Germany), mounted
onto gelatin-coated slides and coverslipped with DPX. The DRGs were
cryoprotected in 20-30% sucrose for 2-3 days at 4°C, embedded in Tissue-Tek
(O.C.T., Miles Inc., Elkhart, IN, USA), frozen at -80C, cut in serial 40µm thick
sections on a cryomicrotome (Leica Instruments), mounted on gelatin-coated slides
and coverslipped with DPX(Novikova et al. 1997). For morphometric analyses and
axon counts (Paper IV), 2 mm long nerve specimens were excised at 3-5 mm
distance proximal and distal to the implantation site and from the middle of the
nerve graft. The nerves were additionally fixed in 3% glutaraldehyde overnight,
postfixed in 1% OsO4 in 0.1 M cacodylate buffer (pH=7.4), dehydrated in acetone,
and embedded in Vestopal. Semithin transverse sections were cut on a 2128
Ultratome (LKB, Sweden) and counterstained with Toluidine Blue. For laser
microdissection (Paper II), L4 and L5 DRG were rapidly harvested without
perfusion and frozen in liquid nitrogen. Serial 10 µm cryosections of DRG were cut
using Bright (UK) 5040 microtome and mounted onto RNase-free UV light-treated
membrane slides: 1 mm polyethylene-tetraphthalate (PET) membrane, PALM
(Zeiss, Germany). Slides were air-dried in the cryostat before being fixed in icecold methacarn (8 parts methanol, 1 part glacial acetic acid) for 10 minutes, dipped
into ice-cold RNase-free PBS to remove excess OCT, and then in serial ethanol
(70/96/100% for 2 minutes/2 minutes/3 minutes) respectively for tissue dehydration.
23
Dag Welin
Neuronal counts
Nuclear profiles of labeled neurons were counted in all sections at x250
magnification in a Leitz Aristoplan fluorescence microscope using filter block A
(Fast-Blue and Fuoro-Gold) and N2.1 (Fluoro-Ruby). The total number of nuclear
profiles were not corrected for split nuclei, since there was a uniformity in nuclear
size and since the nuclear diameters were small in comparison with the section
thickness (Ma et al. 2001). Furthermore, in estimations of retrograde cell death the
accuracy of this technique is similar to that obtained by using physical disector (Ma
et al. 2001) or by counting neurons reconstructed from serial sections (Novikova et
al. 1997). In Paper III, the cross-sectional soma area of the labeled neurons was
measured with a Eutectic Neuron Tracing System (Raleigh, North Carolina) at ×250
magnification. In Paper IV, myelinated axons in the proximal and distal nerve
stumps, and in the middle of the nerve graft were counted at x1000 final
magnification using Stereo Investigator™ 6 software (MicroBrightField, Inc.,USA).
Laser microdissection
Laser capture microdissection was performed on PALM Microlaser Technologies
microbeam microdissection system (Zeiss, Germany). Retrogradely labeled
fluorescent neurons were visualized, marked under fluorescence illumination (FB UV filter, 350 nm excitation; FR – Cy3 filter, 555 nm excitation) and cut. The PET
membrane of the slide is cut simultaneously and provides a ‘back-bone’ to facilitate
laser pressure catapulting - a precisely aimed but defocused high energy laser beam
to catapult the area against gravity into the collecting plastic tube. Individual
neurons were catapulted contact-free into a PALM AdhesiveCap with a total of 90100 cells captured for each group. This procedure allows collection of material in a
contact-free manner, minimizing the risk of contamination which is particularly
important in sensitive downstream applications.
RNA isolation and quantitative RT-PCR
Laser captured samples were submerged in 350 µl lysis buffer (RLT buffer,
QIAGEN, Germany) and 3.5 µl β-mercapotethanol (Sigma-Aldrich, UK) was added
alongside 20 ng carrier RNA before homogenization. The samples were incubated
upside down for 30 minutes, then vortexed thoroughly. RNeasy (Qiagen) Micro
protocol for isolation of total RNA from microdissected cryosections was
undertaken including the optional DNase step. The purified RNA was eluted in 14 µl
RNase-free water. The RNeasy (Qiagen) Mini protocol was used for isolation of
total RNA from whole L4 and L5 DRG in the HPRT validation group. In this group,
total RNA was analysed on a NanoDrop™ 1000 spectrophotometer (Thermo
Scientific, USA) for quality and quantity of product. Five micrograms of each
sample was then used for qRT-PCR to ensure standardized starting total RNA
24
Neuroprotection and axonal regeneration after peripheral nerve injury
quantity. Unfortunately, the quantities of laser-microdissected sample RNA were <
2ng/µl and therefore could not be accurately quantified; however, the quality of
RNA was satisfactory.
All total RNA samples were converted to cDNA using a First Strand cDNA
Synthesis Kit (Superarray, U.S.A.). qRT-PCR was performed with a Rotor-Gene
6000 (Corbett Life Science, Australia) using SYBR™ Green fluorescence mastermix (Superarray, U.S.A.) and analysed using Rotor-Gene 6000 Series Software
version 1.7.61 (Corbett Life Science, Australia). Primers were pre-designed by
Superarray - housekeeping gene HPRT (Genbank Accession No. NM012583,
Catalogue No. PPR44247E); genes of interest peripherin (Genbank Accession No.
NM012633, Catalogue No. PPR45223A), neurofilament triplet proteins of high
molecular weight, NF-H (Genbank Accession No. NM012607, Catalogue No.
PPR42491A) and activating transcription factor 3, ATF3 (Genbank Accession No.
NM012912, Catalogue No. PPR44403A). All reactions had been optimised to work
under the same conditions – initial denaturation/HotStart DNA Polymerase
activation: 95°C for 15 minutes; PCR cycles: 95°C for 30 seconds, 55°C for 30
seconds, and 72°C for 30 seconds repeated for 40 cycles. One PCR run was
performed for each named gene and contained a standard curve, generated from
serial dilutions of cultured DRG neuron cDNA over 3 orders of magnitude; all
experimental samples were assayed in duplicate. A negative control assay was
always included where cDNA template was replaced with RNase-free water. From
the standard curves described above, the C(t) values for the three genes of interest
were used to calculate mRNA levels (arbitrary units) in each sample. For every
animal, the expression level of each gene was normalized to that of HPRT (Reid et
al. 2009). Confirmation of the amplified products was established by performing a
melting curve analysis: 95°C for 1 minute, 65°C for 2 minutes, then 65 - 95°C,
reading every 0.2°C, holding for 1 sec between reads.
Image processing
Preparations were photographed with a Nikon DXM1200 digital camera attached to
a Leitz Aristoplan microscope. The captured images were resized, grouped into a
single canvas and labeled using Adobe Photoshop CS3 software. The contrast and
brightness were adjusted to provide optimal clarity.
Statistical analysis
In Papers I, III and IV, one-way analysis of variance (ANOVA) followed by a post
hoc Newman-Keuls test or Tukey test (Prism®, GraphPad Software, Inc; San Diego,
California) were used to determine statistical differences between experimental
groups. In Paper II, ANOVA followed by Bonferroni’s Multiple Comparison test
25
Dag Welin
and unpaired student t-test were used to determine differences in gene expression.
All data was expressed as mean ± S.E.M. A value of p < 0.05 was considered to be
statistically significant.
26
Neuroprotection and axonal regeneration after peripheral nerve injury
RESULTS
Peripheral nerve injury induces delayed loss of cutaneous sensory neurons
In control rats, 3,340 (± 176 S.E.M.) cutaneous sural DRG neurons and 216 (± 8
S.E.M.) muscular gastrocnemius DRG neurons were labeled by Fast Blue (FB) at 1
week after application of the dye to the homonymous nerves (Table 1, Fig. 1, Paper
I and Tables 1 and 2, Fig. 1, Paper IV). Counts of Fluoro-Gold-labeled neuronal
profiles revealed no significant loss of sural or gastrocnemius DRG neurons at 3
days, 1 week and 2 weeks after axotomy (Table 1, Paper I). Therefore, the numbers
of FB-labeled sural and gastrocnemius neurons at 1 week after tracer application
were used as baseline controls in the subsequent experiments (“Control” in Tables
1-5, Paper I). Counts of FB-labeled sural DRG neurons after axotomy demonstrated
22% cell loss after 4 weeks (P<0.01, Table 1, Paper I) and 45% loss at 8-24 weeks
post-operatively (P<0.001; Table 1, Paper I). In contrast, no significant retrograde
cell loss was found among the gastrocnemius DRG neurons up to 24 weeks after
nerve transection (P>0.05; Table 1, Paper I). Delayed proximal transection of the
sciatic nerve at 1 week after FB-labeling did not further increase retrograde
degeneration of the sural DRG neurons (P>0.05; compare Table 1 and Table 2,
Paper I), nor did it affect survival of the medial gastrocnemius DRG neurons (Table
3, Paper I). There was no loss of spinal motoneurons after sciatic axotomy (Tables 4,
5, Paper I and Tables 1, 2, Paper IV).
Spinal motoneurons degenerate after ventral rhizotomy and avulsion
In control rats, 187 (± 10 SEM) medial gastrocnemius motoneurons were labeled by
True Blue (TB) at 1 week after application of the dye to the transected peripheral
nerve (Table 1, Paper III). Tracing with Fluoro-Gold (FG) labeled 172 (± 5 SEM)
spinal motoneurons. There was no difference in efficacy of these retrograde tracers
(P>0.05). Both ventral rhizotomy and ventral root avulsion induced significant
retrograde degeneration among the axonally injured motoneurons. Ventral
rhizotomy resulted in the death of 26% of medial gastrocnemius motorneurons
within 8 weeks (Figure 1B, Paper III) and a 21% reduction in soma area (rhizotomy
+ no treatment, Table 1, Paper III). The more severe avulsion injury resulted in the
death of 53% spinal motoneurons and a 31% reduction in soma area within 4 weeks
(avulsion + no treatment, Table 2, Paper III).
Nerve repair improves survival of axotomized sensory neurons
Neither primary repair (p<0.05) nor peripheral nerve grafting (p<0.01) prevented
neuronal degeneration among the sural DRG neurons (Table 1, Paper I). However,
primary nerve repair increased survival rate of the sural DRG cells by 30%
(P<0.001; Table 2, Paper I), whereas nerve grafting increased their survival by 19%
27
Dag Welin
(P<0.01; Table 2, Paper I; Table 1, Paper IV). However, there was no statistical
difference between the two nerve repair techniques (P>0.05; Table 2, Paper I). The
numbers of medial gastrocnemius DRG neurons and sural and gastrocnemius
motoneurons were unaffected by nerve repair (Tables 3-5, Paper I; Table 2, Paper
IV).
Regeneration of sensory neurons and motoneurons after nerve repair
Neuronal regeneration across the nerve repair site was quantified using second
labeling with Fluoro-Ruby (FR). At 13 weeks after sciatic axotomy and nerve
grafting, 32% of the sural DRG neurons had succeeded in regenerating their axons
10 mm into the distal stump of the sciatic nerve (Table 2; Fig. 2, Paper I; Table 1;
Fig 1, Paper IV). Primary repair was more efficient and promoted axonal
regeneration of 54% of the sural DRG cells (P<0.001; Table 2, Paper I). This
difference between the two nerve repair methods was not seen among the
gastrocnemius DRG cells, where about 50% of the neurons regenerated across the
lesion site after both primary repair and nerve grafting (Table 3, Paper I). When
calculating the ratio between the number of regenerating (FB+FR)-labeled DRG
cells and the number of FB-labeled DRG neurons remaining at 13 weeks after
axotomy and nerve repair, we found that the proportion of regenerating neurons
(FR+FB/FB ratio in Table 2, Paper I and FR/FB ratio in Table 1, Paper IV) after
primary nerve repair and nerve grafting was similar between the sural and
gastrocnemius DRG populations. With respect to regeneration of the sural and
gastrocnemius motoneurons, 53-59% of these cells became labeled by FR from the
distal sciatic nerve stump (Tables 4, 5; Fig. 2, Paper I and Table 2, Paper IV).
Regenerating sensory neurons up-regulate peripherin mRNA expression
Peripherin mRNA expression in sensory DRG neurons with regenerating axons
(FB+FR double-labeled neurons) was significantly higher than in control noninjured sural and gastrocnemius neurons. Axotomized sural and gastrocnemius
sensory neurons that failed to produce axonal outgrowth (FB-labeled cells) did not
significantly upregulate peripherin and transcript levels were similar to that seen in
non-injured neurons (Fig. 2, Paper II). The relative value of peripherin mRNA was
significantly higher in regenerating sural sensory neurons than in regenerating
gastrocnemius neurons (P<0.05). However, the sevenfold increase in transcripts of
gastrocnemius neurons is a proportionally greater response to injury than the
fourfold increase in sural neurons.
Sensory neurons down-regulate expression of NF-H mRNA after injury
The relative expression of NF-H in control non-injured neurons was significantly
higher in gastrocnemius subpopulation of sensory neurons (Fig. 3, Paper II). Injured
28
Neuroprotection and axonal regeneration after peripheral nerve injury
sural and gastrocnemius sensory neurons with established axonal regeneration
(FB+FR cells) significantly down-regulated NF-H transcripts, whereas injured nonregenerating sensory neurons (FB cells) reduced transcripts levels even further.
There were no significant differences between the regenerating and nonregenerating groups in niether sural nor gastrocnemius neuronal subpopulations.
Regenerating sensory neurons up-regulate ATF3 mRNA expression
There was a significant up-regulation of ATF3 in regenerating sural and
gastrocnemius sensory neurons (Fig. 4; Paper II).There was no significant difference
between the control and non-regenerating groups in the MG neuronal subpopulation.
These results highlight a difference in ATF3 expression between neurons with
proven regeneration and neurons that had failed to regenerate in both subpopulations
of sensory neurons.
N-acetyl-cysteine promotes survival of spinal motoneurons
Following ventral rhizotomy, intraperitoneal N-acetyl-cysteine (NAC) maintained
soma area, and exhibited dose-dependent neuroprotection (Table 1, Paper III). The
effect being statistically significant at the higher dose tested (750mg/kg/day, 7%
neuron death, P<0.05). Intrathecal NAC was slightly more neuroprotective (5%
neuron death, P<0.05; Figure 1C), and also preserved soma area (P<0.001).
When compared to untreated controls intraperitoneal NAC (150mg/kg/day) was also
significantly neuroprotective (40% neuron death, P<0.01) after ventral root avulsion
(Table 2, Paper III), and had some effect to maintain soma area (81% of normal).
Intrathecal delivery was even more effective, reducing neuron death to only 30%
(P<0.001), and significantly maintaining soma area (92% of normal, P<0.001). Even
when intrathecal NAC treatment was delayed by two weeks after avulsion (Table 2,
Paper III), it still conferred a highly significant degree of neuroprotection (42%
neuron death vs. 56% death with sham treatment, P<0.001). The protective effect
was significantly greater (P<0.05 vs. 2-week delay) when treatment was delayed for
only one week and interestingly there was no loss of neuroprotection as compared to
immediate treatment.
No neurotoxic effect of intrathecal or intraperitoneal treatment with NAC was
demonstrated (Tables 1 and 2, Paper III). Similarly, the mean soma area of spinal
motoneurons neurons was unaffected by either route of treatment (Tables 1 and 2,
Paper III). No adverse effects attributable to treatment or minipump implantation
were encountered during this study.
29
Dag Welin
N-acetyl-cysteine promotes survival of cutaneous sensory DRG neurons
Eight weeks of continuous intrathecal treatment with NAC resulted in 78% survival
of the sural sensory DRG neurons (P<0.01; Table 1; Fig. 1A-C, Paper IV).
However, when nerve grafting was combined with intrathecal infusion of NAC for 8
weeks, survival of DRG neurons was increased by 38% (p<0.01; Table 1; Fig.
1B,D,G, Paper IV) and was not different from the baseline control at 1 weeks after
FB labeling (p>0.05, Table 1, Paper IV). Therefore, nerve grafting and intrathecal
NAC infusion have highly significant additive effects on survival of sensory DRG
neurons after peripheral nerve injury. Survival of sural motoneurons was not
affected by nerve grafting and NAC treatment (p>0.05; Table 2, Paper IV).
Effects of N-acetyl-cysteine and nerve grafting on neuronal regeneration
When nerve grafting was combined with intrathecal NAC treatment, the proportion
of regenerating neurons was statistically increased to 42% (p<0.05; Table 1; Fig. 1E,
H; Paper IV). Thus, improved survival of the sural sensory neurons resulted in better
outcome of neuronal regeneration across the nerve graft. However, when
calculating the ratio of regenerating FR-labeled and remaining FB-labeled sural
sensory neurons at 13 weeks postoperatively, we found that the proportions of
regenerating neurons were similar after nerve grafting alone (46.3%; Table 1, Paper
IV) and nerve grafting followed by intrathecal NAC treatment (45.6%; Table 1,
Paper IV). NAC treatment had no growth-promoting effect on axotomized spinal
motoneurons (p>0.05; Table 2, Paper IV).
Effect of N-acetyl-cysteine and nerve grafting on axonal sprouting
All examined specimens from the sciatic nerve and graft contained numerous
myelinated fibers. However, there were marked differences in appearance and axon
number between the experimental groups as well as between specimens taken from
the nerve stumps and grafts (Figure 2; Table 3, Paper IV). Proximal nerve stumps
contained 11,041 ± 302 (mean ± S.E.M.) and 11,092 ± 543 myelinated fibers in the
groups with nerve graft and nerve graft with NAC treatment, respectively. Although
the number of myelinated axons was significantly increased in the middle of the
graft by 54.2% (p<0.001) and 47.1% (p<0.001) in untreated and NAC-treated
animals, respectively, there was no statistical difference between the experiments
(p>0.05). The numbers of myelinated fibers in the distal nerve stump were
significantly decreased (p<0.001) and were comparable with the counts in the
proximal stump of the nerve (105-109%; Table 3, Paper IV). The axons in the graft
and in the distal stump also appeared smaller in diameter and had thinner myelin
sheath. Thus, NAC treatment had no effect on sprouting of myelinated axons in the
graft and distal nerve stump.
30
Neuroprotection and axonal regeneration after peripheral nerve injury
DISCUSSION
Fluorescent retrograde tracers for neuronal quantification
It has been suggested that an “assumption-free” stereological method is one of the
most accurate techniques for neuronal quantification (Coggeshall and Lekan 1996;
West 1999). However, there are also opposing views indicating that 3-D studies
could have potentially huge biases (Guillery and Herrup 1997; von Bartheld 2001;
Guillery 2002). For example, using “unbiased” counts of DRG neurons after sciatic
nerve transection highly significant variability in the timing and amount of cell loss
has been reported (Tandrup 1993; Tandrup et al. 2000; Hart et al. 2002a). In
previous investigations in our laboratory, physical disector has been compared with
both long-term retrograde Fast Blue-labeling (Ma et al. 2001) and serial
reconstructions of labeled profiles (Novikova et al. 1997) and we have found that
Fast Blue-labeling provides accurate estimates of retrograde cell death. The
existence of different cell populations in DRG (Papers I, II and IV) should also be
taken into account when designing experiments to study retrograde degeneration
after peripheral axotomy. If only certain subpopulations of sensory neurons
degenerate after axotomy, then quantification of total number of DRG neurons even
with modern stereological methods (Coggeshall and Lekan 1996; West 1999) will
underestimate the degree of retrograde cell death. In addition, only about 50% of all
neurons in the L4 and L5 DRGs project their axons to the sciatic nerve (Swett et al.
1991).
Moreover, in the present thesis, regeneration could be examined only by using a
double labeling technique. For this purpose, Fast Blue (FB) was used as a long-term
marker to assess neuronal survival (Novikova et al. 1997), while Fluoro-Ruby (FR)
was used to evaluate axonal regeneration across the injury site (Novikova et al.
1997; Ma et al. 2003). Since Fast Blue requires about one week to provide effective
retrograde labeling of neurons, the more rapidly transported tracer Fluoro-Gold (FG)
was used to study possible retrograde cell degeneration during the first couple of
days after peripheral nerve injury (Novikova et al. 1997). To study gene expression
in DRG neurons (Paper II), Fast Blue was also injected into the medial
gastrocnemius muscle and sural nerve. Although this technique labels only limited
number of sensory and motor neurons, the main goal was to minimize axon reaction
in control neurons.
Mechanism of retrograde cell death after peripheral nerve injury
Neurons may die by either passive necrosis, or an active metabolic process akin to
apoptosis whose intracellular events show a high degree of homology irrespective of
the initiating event (Koliatsos and Ratan 1999). In traumatic injury necrosis is more
31
Dag Welin
associated with direct trauma to the cell body, and occurs too rapidly to be amenable
to therapeutic intervention, but active cell death tends to be a more progressive
secondary phenomenon, giving a potential therapeutic window. Apoptosis is a
complex process involving a variety of different signaling pathways and after
peripheral axotomy it could be initiated by interrupted retrograde transport of targetderived neurotrophins, GDNF family members and neuropoetic cytokines signaling
through the Ras-ERK pathway, the PI3-K pathway and the JAK-STAT pathway
(Kaplan and Miller 2000; Cui 2006; Abe and Cavalli 2008). Initially, axotomy
induces transcription of regenerative as well as cell death genes (Nunez and del Peso
1998), the neuron’s fate lying in the ultimate balance of these gene products
(Gillardon et al. 1996). How the neuron finally commits to cell death rather than
regeneration is not fully understood, but has been associated with abortive entry into
the cell cycle (Freeman 1999), and mitochondria are thought to play a key role (Al
Abdulla and Martin 1998; Budd and Nicholls 1998). It is likely that the normal
electron transport mechanisms are damaged by excess reactive oxygen species
(ROS), possibly in response to unphysiological elevation of intramitochondrial nitric
oxide (Heales et al. 1999; Elliott and Snider 1999). As a result oxidative metabolism
is impaired, mitochondrial homeostasis fails, and they release pro-apoptotic
molecules (Budd and Nicholls 1998) that trigger a caspase cascade culminating in
active cell death.
Response of spinal motoneurons to distal and proximal axotomy
Several factors including the age of the experimental animals, the distance from the
lesion site to the cell body and the type of the injury can determine the severity of
the retrograde axon reaction. Injury to the sciatic nerve in newborn rodents almost
completely eliminates the corresponding spinal motoneurons (Snider et al. 1992;
Vejsada et al. 1995; Li et al. 1998), while the same type of injury in adult animals
has no effect on motoneuron survival (Carlson et al. 1979; Vanden Noven et al.
1993; Novikova et al. 1997) although it induces cell atrophy, synaptic shedding and
degeneration of dendrites (Kreutzberg GW 1995; Carlstedt and Cullheim 2000). The
present results are in line with these observations and show that neither sural nor
gastrocnemius motoneurons degenerate after a division of the distal nerve branch or
permanent sciatic axotomy. However, proximal injuries do induce retrograde
motoneuronal death, such that 20-30% die within 16 weeks after spinal nerve
transection (Ma et al. 2001; Ma et al. 2003; Jivan et al. 2006) and more than 50%
die six weeks after C7 ventral rhizotomy (Gu et al. 1997), and after ventral root
avulsion (Wu and Li 1993; Koliatsos et al. 1994; Novikov et al. 1995; Elliott and
Snider 1999). The cell bodies of motoneurons become atrophic as a reaction to
various types of axonal injury, and this change correlate with the severity of trauma.
After proximal axotomy, an early hypertrophy (12% after 4 weeks) is followed by
atrophy (20% at 16 weeks)(Ma et al. 2001) while after ventral root avulsion,
32
Neuroprotection and axonal regeneration after peripheral nerve injury
significant 50% cell atrophy occurs at 4 weeks postoperatively (Novikov et al.
2000). The present results (Paper III) are consistent with these findings.
Sensory neurons after peripheral nerve injury
In contrast to motoneurons, adult DRG neurons undergo significant retrograde cell
death also after distal peripheral nerve injury. Recent studies have shown that
permanent sciatic transection mainly induces degeneration of small diameter DRG
neurons projecting to skin and expressing neurotrophin receptors trkA and trkC as
well as GDNF receptors (McMahon et al. 1994; Bennett et al. 1998; Karchewski et
al. 1999; Tandrup et al. 2000; Hu and McLachlan 2003). Studies with fluorescent
tracers have shown that permanent sciatic axotomy induces significant but delayed
retrograde cell death of sural sensory neurons projecting mainly to skin and does not
affect survival of medial gastrocnemius sensory neurons projecting to muscle (Hu
and McLachlan 2003). The loss of cutaneous afferent neurons is likely to contribute
to the poor sensory recovery observed clinically after nerve lesions, since neuronal
survival is essential for regeneration (Fu and Gordon 1997), and since the quality of
sensation depends upon the number of primary neurons and the size and degree of
overlap of their receptive fields (Lundborg 2000b). It is not known why cutaneous
afferent neurons are more vulnerable than muscular afferent neurons, since
peripheral nerve injury up-regulates a variety of neurotrophic factors both in the
proximal and distal nerve stumps (Gillen et al. 1997; Terenghi 1999; Boyd and
Gordon 2003). One explanation could be a mismatch between the neurotrophic
factors expressed after injury and the receptors present in the axotomized sensory
neurons. It is also possible that the amounts of retrogradely transported neurotrophic
factors (Curtis et al. 1998) are insufficient for activating the intracellular pathways
responsible for survival of cutaneous DRG neurons. When peripheral axotomy is
treated with exogenous neurotrophins, almost complete survival of sensory DRG
neurons can be achieved (Ljungberg et al. 1999; Kuo et al. 2005).
Our findings are in agreement with previous observations that DRG neurons
undergo delayed degeneration after peripheral nerve injury (Tandrup et al. 2000;
Jivan et al. 2006), although there are conflicting reports about the time course of
degeneration (Tandrup et al. 2000; Hart et al. 2002a). Although the mechanism for
delayed neuronal degeneration remains unclear, this phenomenon has been
described in cervical motoneurons after brachial plexus injury (Ma et al. 2001; Jivan
et al. 2006), in the red nucleus after cervical spinal cord injury (Houle and Ye 1999;
Novikova et al. 2000b) and in the cerebral cortex after traumatic brain injury (Smith
et al. 1997). One possible explanation for the delayed onset of retrograde reaction
could be that during the first 3-4 weeks after injury, neurons convert from a
“transmitting mode” to a “growth mode”, which results in axonal sprouting and
establishment of transient synaptic contacts. It has been shown that peripheral
axotomy induces formation of supernumerary intramedullary axons in spinal
motoneurons (Havton and Kellerth 1987) as well as collateral sprouting of
33
Dag Welin
myelinated axons in the dorsal horn (Bennett et al. 1996) and dorsal roots (Lekan et
al. 1997). Other possibilities include retrograde transport of neurotrophic factors
from spinal motoneurons to primary afferents via mechanism of transsynaptic
transcytosis (Rind et al. 2005). However, the latter assumption does not agree with
observations that combined transection of peripheral nerve and dorsal roots induces
a similar retrograde cell loss as peripheral nerve injury alone (Coggeshall and Lekan
1996).
Neuroprotective effect of nerve grafting
After peripheral nerve injury, Schwann cells in the distal nerve stump divide and upregulate expression of various neurotrophic factors including NGF, BDNF, NT-4/5,
GDNF and LIF (Boyd and Gordon 2003). Although it has been demonstrated that
Schwann cell proliferation in the distal nerve stump is not required for functional
recovery after nerve injury (Yang et al. 2008), it is well known that expression of
neurotrophic molecules by Schwann cells supports both neuronal survival and
axonal regeneration (Terenghi 1999; Boyd and Gordon 2003). The latter findings
has lead to numerous experimental and clinical studies using peripheral nerve grafts
to prevent retrograde cells death and to direct axonal growth toward peripheral
targets (Rhrich-Haddout et al. 2001; Ma et al. 2003; Bertelli and Ghizoni 2003; Wu
et al. 2004). In previous investigations we have described neurotrophic and
neurotropic effects of primary nerve repair and peripheral nerve grafting on the
survival and axonal regeneration of cervical DRG neurons and spinal motoneurons
(Ma et al. 2003; Jivan et al. 2006). In the latter experiments delayed nerve grafting
promoted motoneuron regeneration more efficiently than regeneration of DRG cells.
In the present study, however, the proportion of regenerating neurons was rather
similar between cutaneous afferent cells, muscular afferent cells and spinal
motoneurons. The reason for improved regeneration is unclear but could be
influenced by the nature of the graft and the timing of nerve repair. Today there is
no real substitution for nerve grafting in patients after peripheral nerve injuries with
long nerve gaps. However, despite the advantages in microsurgical reconstruction of
injured peripheral nerves, the technique of nerve grafting is not optimal and patients
suffer from loss of sensation, scarring and sometimes pain in the donor region
(Wiberg and Terenghi 2003). Moreover, neuroprotection provided by peripheral
nerve graft for sensory DRG neurons in the present study seems to be inferior to the
reported neuroprotective efficacy of systemically administered exogenous
neurotrophic factors (Matheson et al. 1997; Ljungberg et al. 1999; Groves et al.
1999).
Regulation of peripherin and ATF3 genes in primary sensory neurons
Peripheral nerve injury induces a coordinated molecular response which determines
the intrinsic growth state of axotomized neurons through regulation of numerous
transcription factors, growth factors, adhesion molecules and structural components
34
Neuroprotection and axonal regeneration after peripheral nerve injury
(Navarro et al. 2007; Raivich and Makwana 2007). It has been shown that
transcription is necessary for successful axon growth (Smith and Skene 1997) and
changes observed have been likened to a recapitulation of the developmental
phenotype (Costigan et al. 2002; Xiao et al. 2002). In our second study (Paper II),
we demonstrated that, when regenerating across the lesion site, cutaneous sural
DRG neurons and muscular medial gastrocnemius DRG neurons up-regulated gene
expression of peripherin and ATF3.
In the peripheral nervous system, peripherin is expressed mainly in small DRG
neurons with unmyelinated axons (Fornaro et al. 2008) and is necessary for the
proper development of non-peptidergic small unmyelinated sensory cells (Lariviere
et al. 2002). In the present study, we found that peripherin mRNA was expressed in
greater quantities in the sural sensory neurons, which are predominantly small
neurons with unmyelinated axons, than in the medial gastrocnemius sensory
neurons, which are mainly large cells with myelinated axons. Our results suggested
that following nerve injury both subpopulations of sensory neurons significantly upregulate peripherin gene expression, and this is contrary to a previous report
demonstrating peripherin transcript up-regulation mainly in large DRG neurons
(Oblinger et al. 1989). This obvious discrepancy in expression pattern could be due
to differing methodologies since in the previous report authors used axotomy of the
sciatic nerve and in situ hybridization with a labeled cDNA probe to quantify
mRNA in small neurons throughout L5, many of which may not have been
axotomized. In contrast, the present study was based on laser microdissected cells
obtained from an identified subpopulation of sensory DRG neurons. It is also
possible that the response in the sural subpopulation was potentiated by the
conditioning injury 8 weeks prior. However, when compared to corresponding
control neurons, the sevenfold increase in medial gastrocnemius sensory cells was
greater than the fourfold increase observed in sural neurons. It has been shown
previously that peripherin is important for the initiation, extension and maintenance
of neurites in PC12 cells (Helfand et al. 2003). Although the role of peripherin is
unknown, it is clearly a highly dynamic structure that interacts with the other neural
intermediate filaments during development and regeneration (Helfand et al. 2004).
The exact mechanism of action of peripherin also remains elusive, although it may
be a substrate for Akt in potentiating regeneration and phosphorylation is an
important post-transcriptional control (Konishi et al. 2007). Our results
demonstrating low peripherin transcript levels in the neurons that failed to
regenerate may be explained by the previous reports that peripherin expression is
maintained in an up-regulated state for about 8 weeks in axotomized DRG neurons
(Wong and Oblinger 1990). Since our experiments took 9 weeks, it is possible that
we have missed an early response to injury which has subsequently returned to
control levels. Alternatively, the lack of regenerating axons in this population may
be a result of failure to up-regulate peripherin in the initial regenerative response.
35
Dag Welin
Our findings regarding the regulation of NF-H mRNA after nerve injury in DRG
subpopulations are in accordance with previous reports examining whole DRG after
sciatic nerve axotomy which have demonstrated an immediate down-regulation in
expression of this neuronal intermediate filament (Wong and Oblinger 1990; Muma
et al. 1990; Fornaro et al. 2008).
The transcription factor ATF3 is rapidly induced in all axotomized sensory DRG
neurons and immunohistochemistry expression persists for many months if
regeneration is prevented (Tsujino et al. 2000; Kataoka et al. 2007). It has been
speculated that ATF3 may promote neuronal survival (Tsujino et al. 2000;
Nakagomi et al. 2003). Recent studies utilizing over-expression of ATF3 have
demonstrated that ATF3 can promote neurite outgrowth both in vitro and in vivo
(Seijffers et al. 2006; Seijffers et al. 2007). Moreover, it has been found that ATF3
expression following injury is differentially regulated in sensory and motor neurons
(Kataoka et al. 2007) and in neurochemically defined sensory subpopulations
(Averill et al. 2004). However, our results demonstrated that gene expression is
much reduced in sensory neurons that failed to regenerate compared to neurons with
proven regeneration. Neurons with regenerating axons profoundly up-regulated
ATF3 transcripts at 1 week after injury, and it might have been expected that
neurons failing to produce axonal outgrowth would maintain a high ATF3 transcript
levels 9 weeks after injury. The dramatic difference in expression may suggest that
this neuronal population is doomed to regenerative failure because of inadequate
ATF3 levels of expression; however, it is also possible that ATF3 expression in nonregenerating neurons decreases more rapidly, after initial induction, as a survival
response (Kataoka et al. 2007). The exact mechanisms are not entirely understood
but it is believed that ATF3 is involved in determining the cell’s fate after injury
(Hua et al. 2006).
N-acetyl-cysteine supports neuronal survival
Following an injury to the nervous system, mitochondrial impairment could lead to
generation of reactive oxygen species and activation of apoptotic cascades (Merenda
and Bullock 2006). With respect to axon reaction, it has been reported that
degeneration of spinal motoneurons after ventral root avulsion could evolve with
oxidative stress (Martin et al. 1999) and often resemble necrosis (Li et al. 1998). In
vitro experiments also show that an increase of mitochondrial-derived reactive
oxygen species occurs in NGF-deprived sympathetic neurons undergoing apoptotic
death (Kirkland and Franklin 2003).
Since mitochondria and the generation of reactive oxygen species are thought to be
important in the mechanism behind cell death, agents that can stabilize oxidative
36
Neuroprotection and axonal regeneration after peripheral nerve injury
metabolism, or enhance mitochondrial protection from reactive oxygen species
could be neuroprotective. Glutathione is the principle species responsible for this
within neurons, and its depletion, particularly within mitochondria, does increase
susceptibility to toxic stimuli including neurotrophin withdrawal (Cooper and
Kristal 1997; Wullner et al. 1999). Based on these findings, it has been proposed
that glutathione replacement could be neuroprotective (Cooper and Kristal 1997).
However, this agent cannot cross the blood-brain barrier and its principle ratelimiting precursor cysteine is unstable in the CSF (Pan and Perez-Polo 1996; Wang
and Cynader 2000). In contrast, L-stereoisomer of N-acetyl-cysteine (NAC) can
cross the blood-brain barrier, accumulate in the brain and increase intracellular
levels of glutathione (Arakawa and Ito 2007). It has been reported previously that
intraperitoneal treatment with NAC (150mg/kg/day) provides significant rescue of
sensory DRG neurons via mitochondrial preservation (Hart et al. 2004). The
protection of sensory neurons by NAC is dose dependent and effective over a wide
therapeutic range (West et al. 2007b).
N-acetyl-cysteine’s mechanism of action remains unclear, and further investigation
will be required, although putative mechanisms have been proposed (Yan and
Greene 1998; Hart et al. 2004). One possible mechanism is as a glutathione
substrate, thereby acting to maintain intramitochondrial defences against reactive
oxygen species. Other possibilities include a direct reductant effect that may either
equilibrate the increased oxidative stress within the injured neuron, or modulate
signal transduction, including that for neurotrophic pathways (Xu et al. 2002; Droge
2002). Such an effect could alter the cells interpretation of reduced neurotrophic
support, or modulate the response in favor of survival and regeneration, and away
from cell death. This latter effect may explain the action of the D-stereoisomer of
NAC, which cannot be metabolised into glutathione, yet still confers a comparable
degree of neuroprotection to the L-stereoisomer in certain in vitro models (Ferrari et
al. 1995).
Our study on motoneurons (Paper III) demonstrates that immediate systemic
treatment with NAC is neuroprotective for adult motorneurons after ventral
rhizotomy, and that a dose response effect is evident in that survival rose to 93%
when 750mg/kg/day was given. Higher doses would have required the i.p. injection
of an excessive volume and were not tested. Intrathecal (i.t.) administration was
used to test the efficacy of higher tissue levels of NAC, but proved comparable to
the 750mg/kg i.p. dose, suggesting that this dose is supra-maximal, as probably is
the 2.4mg/day intrathecal dose. Although systemic treatment (NAC 150mg/kg/day
i.p.) was less neuroprotective than intrathecal, it still provided a highly significant
degree of neuroprotection, and given the dose response evident in the rhizotomy
model it is likely that a higher dose could be comparably effective. Thus, it is
apparent that NAC can reduce, or possibly eliminate, active motoneuronal death
37
Dag Welin
after proximal axotomy in adults, assuming sufficient tissue levels are attained. The
clinical experience of NAC during the management of paracetamol poisoning is that
it is safe and easily administered systemically even at high doses, making it
clinically applicable even for nerve injuries.
However, for NAC to be an applicable therapy for severe nerve injuries, such as the
brachial plexus injury, it must also be effective if treatment is not commenced
immediately after injury, since that will never be clinically feasible. For sensory
neurons neuroprotective therapy can be delayed for 24 hours after distal axotomy
without any loss of effect, and we found no loss of neuroprotective effect when
NAC treatment was delayed until one week after ventral root avulsion (Table 3,
Paper II), presumably reflecting the slower time-course of active motoneuronal
death. Some loss was found with a two-week delay of therapy, but that will still
leave a highly clinically appropriate therapeutic window for NAC therapy.
Our study on sensory neurons (Paper IV) demonstrates that both nerve grafting and
continuous intrathecal infusion of NAC could attenuate retrograde cell death of sural
DRG neurons after sciatic nerve injury. However, combined treatment with NAC
and nerve grafting significantly improves survival rate and increases the number of
regenerating sural sensory neurons. However, the results also indicate that NAC has
no growth-promoting effect on spinal motoneurons and does not affect axonal
sprouting in the peripheral nerve graft and in the distal nerve stump.
In this investigation we administered NAC via continuous infusion (2.4 mg/day)
into subarachnoid space from osmotic mini-pump since this delivery route and
concentration have been more efficient in rescuing spinal motoneurons than
intraperitoneal NAC injections (Zhang et al. 2005). In contrast to our previous
studies, however, we obtained somewhat lower efficacy of neuroprotection (22%
versus 32%). One possible explanation is that we used different experimental
models of retrograde degeneration, administration routes for NAC and techniques to
quantify the extent of retrograde cell death.
The main finding of this study is that NAC and nerve grafting displayed additive
neuroprotective effect which resulted in nearly 90% survival of axotomized sural
sensory neurons and improved regeneration into the distal nerve stump. Although
the mechanism of this effect is unknown, it has been shown that NAC could work in
synergy with cytokine CNTF to prevent killing of oligodendrocytes by TNF-alpha
(Noble and Mayer-Proschel 1996). Several research groups have also reported that
NAC not only works as an antioxidant, but also causes rapid activation of the RasERK signaling pathway and induces immediate early genes in trophic factordeprived PC12 cells and sympathetic neurons (Yan and Greene 1998), and in
amyloid-beta peptide-stressed neuronal cell cultures (Kuperstein and Yavin 2002;
38
Neuroprotection and axonal regeneration after peripheral nerve injury
Hsiao et al. 2008). Moreover, NAC can suppress p38 kinase activation, caspase-9
and caspase-3 cleavage and block apoptosis in vitro (Floyd 1999; Junn and
Mouradian 2001). Activation of the ERK signaling pathway and suppression of the
caspases cleavage by NAC is similar to the effects exerted by various neurotrophic
factors on sensory neurons and spinal motoneurons in vitro and after peripheral
nerve injury in vivo (Liu and Snider 2001; Boyd and Gordon 2003). It has also been
demonstrated that inhibition of ERK and protein kinase A at the time of injury can
significantly impair the capacity of injured dorsal root ganglia axons to re-initiate
growth cones (Liu and Snider 2001; Chierzi et al. 2005). These observations
demonstrate that NAC could add to neurotrophic factors effects to maintain neuronal
survival through the Ras-ERK signaling pathway.
However, NAC does not completely mimic the effects of neurotrophic factors and it
has been shown that unlike the neurotrophin NGF, NAC does not activate the PI3-K
pathway and has no effect on neurite outgrowth in vitro (Yan and Greene 1998). The
latter findings could explain the lack of growth-promoting effect on spinal
motoneurons and the inability to influence axonal sprouting in the graft and in the
distal nerve stump. Nevertheless, NAC has been shown to attenuate Wallerian
degeneration and hyperalgesia after peripheral nerve injury (Wagner et al. 1998) and
to inhibit functional and structural abnormalities of the peripheral nerve in
streptozotocin-induced diabetic rats (Sagara et al. 1996).
39
Dag Welin
CLINICAL IMPLICATIONS
In current medical practice, the focus on treatment of nerve injuries has been on the
technical aspects of the repair, and much has been written about different techniques
of nerve suturing, but no clear advantage has been shown, for example, between
epineural and fascicular suturing. The goal of nerve repair is still to align the cut
ends as exactly as possible with minimal manipulation. The general idea is that an
end-to-end suture is preferable as long as it can be made without tension. In a fresh
nerve injury the nerve ends elasticity remains, often making it possible to perform
an end-to-end suture as long as no nerve tissue is lost.
The most severe type of nerve injury is the traction injury where an end-to-end
repair is not possible. In those cases, for example brachial plexus injuries, the use of
nerve grafts to bridge the gap are mandatory even in the situation of acute nerve
repair. In the clinical situation, peripheral nerve surgery is often delayed due to late
referrals to specialized units. Surgery can also be further delayed by additional
injuries or due to difficulties making a correct diagnosis. For example, in closed
brachial plexus injuries it can be difficult to distinguish between neurapraxia,
axonotmesis and neurotmesis.
Nerve grafts are used when the cut nerve ends cannot be opposed without undue
tension, whether that is a result from reduced elasticity due to scaring or actual loss
of a segment of nerve tissue. The most commonly used donor nerve is the sural
nerve, but other cutaneous nerves can be considered. The harvested nerve is inserted
into the defect as cable graft to connect the different fascicles. One of the main
difficulties is to find and match the fascicles in the proximal and distal nerve ends,
which is specifically difficult after scar and neuroma formation.
The general results of nerve repair are poor but significant improvement in outcome
has been seen if the repair has been performed early after injury (Jivan et al. 2008).
This improvement may be a result of reduced technical difficulties when nerves are
repaired acutely but is more likely a result from a biological point of view as a nerve
injury initiates a complex pattern of events of anterograde and retrograde
degeneration affecting neuronal cell bodies, axons and myelin sheath. Neurons in
the dorsal root ganglion and in the spinal cord can undergo cell death and atrophy
and the process can only be reduced by reconnecting the injured neurons with their
peripheral target organs.
This thesis has focused on the degenerative changes that occur proximal to the
lesion site after nerve injury. Since the survival of neurons projecting into the
peripheral nerve is a prerequisite for regeneration and target organ reinnervation,
understanding of this process is of importance to improve the results after nerve
40
Neuroprotection and axonal regeneration after peripheral nerve injury
injuries. In the thesis we investigated the survival response of spinal motoneurons
and sensory DRG neurons to different types of nerve injuries and the ability for
these neurons to regenerate their axons after nerve reconstruction.
One of the main and realistic goals of brachial plexus surgery today is to promote
functional motor reinnervation of the shoulder and the arm. Sensory recovery after
these injuries is very poor which could be due to the high vulnerability of cutaneous
sensory DRG neurons to axonal injury. Thus, prevention of retrograde cell death in
sensory ganglia could therefore possibly improve further clinical results after nerve
reconstructions.
In this study we demonstrated experimentally that NAC is an effective
neuroprotective agent which can reduce post-traumatic cell loss in both DRG and
spinal cord. Furthermore, we showed that by reducing the neuronal loss we can
achieve a higher number of regenerating cutaneous sensory neurons. Although
neuroprotective therapy is not used in today’s clinical practice in peripheral nerve
surgery, it has a great potential as neuroprotective treatment with NAC could be
initiated at the first suspicion of a nerve injury and before referral of the patient to a
specialized unit, thereby reducing the negative effect of a delayed nerve repair.
In the present study we used different routes of drug administration including
inthrathecal NAC infusions. However, in a clinical setting, treatment with either
intravenous or oral administration resembling the treatment for paracetamol
intoxication seems to be more realistic. The neuroprotective dose in humans has not
yet been established, but in rats a dose of 30mg/kg/day has been shown to provide
complete protection of sensory neurons in the DRG (Hart et al. 2004). The dose
given for paracetamol intoxication in humans is approximately 200mg/kg/day. The
duration of neuroprotective treatment in humans is not known, however, it seems
reasonable to consider treatment during the period of the most extensive neuronal
death or at least until nerve repair has been performed.
Future evaluation of possible neuroprotective effects of NAC in humans will also
need non biased techniques to evaluate neuronal survival in the DRG and spinal
cord. One promising strategy could be based on recent MRI studies (West et al.
2007a) demonstrating the possibility, by the use of a non-invasive technique, to
quantify retrograde neuronal death following nerve injury.
41
Dag Welin
CONCLUSIONS
The present thesis investigates the efficacy of N-acetyl-cysteine treatment, primary
nerve repair and nerve grafting on the survival and regeneration of sensory DRG
neurons and spinal motoneurons after peripheral nerve injury in adult rats.
On the basis of the experimental data, the following conclusions can be made:
•
Permanent transection of the peripheral nerve induces delayed retrograde
degeneration of sensory neurons projecting mainly to skin but does not affect
survival of sensory neurons projecting to muscles. Distal axotomy does not lead
to any significant cell death among spinal motoneurons.
•
Immediate repair of the divided sciatic nerve with primary suture or peripheral
nerve grafting attenuates retrograde cell death in cutaneous afferent neurons and
supports axonal regeneration. Although the regenerative capacity of surviving
cutaneous afferent neurons is not reduced by the axotomy, recovery of sensory
function could be significantly affected by the cell loss which occurs also after
nerve repair.
•
Subpopulations of regenerating sural and medial gastrocnemius DRG neurons
after immediate primary nerve repair up-regulate transcription of peripherin and
ATF-3, however, expression of the structural neurofilament proteins of high
molecular weight was significantly down-regulated following injury in all
subpopulations of sensory neurons.
•
Continuous intrathecal and intraperitoneal treatment with N-acetyl-cysteine
provides a highly significant protection of spinal motoneurons after ventral
rhizotomy and avulsion injury, and sensory neurons after peripheral nerve
injury.
•
Combined treatment with N-acetyl-cysteine and nerve grafting significantly
improves survival rate and increases the number of regenerating sensory
neurons. However, N-acetyl-cysteine has no growth-promoting effect on spinal
motoneurons and does not affect axonal sprouting in the peripheral nerve graft
or in the distal nerve stump.
42
Neuroprotection and axonal regeneration after peripheral nerve injury
ACKNOWLEDGEMENTS
During the years of research quite a few people have crossed my way and I would
like to thank you all for contributing to this research.
Especially I would like to thank:
Dr Lev Novikov for being a superb supervisor. Without all your help this work
would not have been possible. Thank you for everything.
Professor Mikael Wiberg for supervision and for leading me into research and hand
surgery.
Professor Jan-Olof Kellerth for being my supervisor and for giving me support and
advice.
Dr Liudmila Novikova for teaching me all the practical methods.
Gunnel Folkesson, Maria Brohlin and Gunvor Hällström, Anna-Lena Tallander and
Göran Dahlgren for all the technical assistance.
All colleagues working in parallel in the lab, with nurturing parties and coffee
breaks: Chris West, Sharmila Jivan, Cheng-Gang Zhang, Aleksandra McGrath,
Jonas Pettersson, Christina Ljungberg, Amar Karalija, Paul Kingham, Daniel
Kalbermatten, Frankie and Samuel Jonsson.
I also would like to express my thanks to the people from Manchester University:
Professor Giorgio Terenghi for your helpful comments and suggestions and also
Andy Hart, Adam Reid, Andy Wilson and Alex Hamilton.
And finally, to my dear Tuva for always supporting me.
This study was supported by the Swedish Medical Research Council, Umeå
University, EU, County of Västerbotten, Åke Wibergs Stiftelse, Magn. Bergvalls
Stiftelse, Clas Groschinskys Minnesfond and the Gunvor and Josef Anér
Foundation.
43
Dag Welin
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