Department of Food Technology
University of Helsinki
Finland
EKT series 1445
Properties of intramuscular connective tissue in
pork and poultry with reference to weakening of
structure
Liisa Voutila
ACADEMIC DISSERTATION
To be presented, with the permission of the Faculty of Agriculture and Forestry of the
University of Helsinki, for public examination in lecture room XII, University main
building,
on 03 April at 12 noon.
Finland 2009
Custos and supervisor
Professor Eero Puolanne
Department of Food Technology
University of Helsinki
Helsinki, Finland
Reviewers
Dr. Jacques Lepetit
Quality of Animal Products,
INRA Clermont-Ferrand UR 370
Saint Gene`s Champanelle, France
and
Dr. Ian Richardson
Division of Farm Animal Science
University of Bristol
Langford
United Kingdom
Opponent
Associate Professor Jim Claus
Meat Science & Muscle Biology Laboratory
University of Wisconsin-Madison
Madison, WI, USA
ISBN 978-952-10-5378-8 (paperback)
ISBN 978-952-10-5379-5 (PDF)
ISSN 0355-1180
Helsinki University Print
Helsinki 2009
Voutila, L. 2009. Properties of intramuscular connective tissue in pork and poultry with reference
to weakening of structure (dissertation). EKT series 1445. University of Helsinki. Department of
Food Technology, 94 pp.
Abstract
The most common connective tissue research in meat science has been conducted on the
properties of intramuscular connective tissue (IMCT) in connection with eating quality of
meat. From the chemical and physical properties of meat, researchers have concluded that
meat from animals younger than physiological maturity is the most tender. In pork and
poultry, different challenges have been raised: the structure of cooked meat has weakened.
In extreme cases raw porcine M. semimembranosus (SM) and in most turkey M.
pectoralis superficialis (PS) can be peeled off in strips along the perimysium which
surrounds the muscle fibre bundles (destructured meat), and when cooked, the slices
disintegrate. Raw chicken meat is generally very soft and when cooked, it can even be
mushy.
The overall aim of this thesis was to study the thermal properties of IMCT in porcine
SM in order to see if these properties were in association with destructured meat in pork
and to characterise IMCT in poultry PS. First a 'baseline' study to characterise the thermal
stability of IMCT in light coloured (SM and M. longissimus dorsi in pigs and PS in
poultry) and dark coloured (M. infraspinatus in pigs and a combination of M. quadriceps
femoris and M. iliotibialis lateralis in poultry) muscles was necessary. Thereafter, it was
investigated whether the properties of muscle fibres differed in destructured and normal
porcine muscles.
Collagen content and also solubility of dark coloured muscles were higher than in light
coloured muscles in pork and poultry. Collagen solubility was especially high in chicken
muscles, approx. 30 %, in comparison to porcine and turkey muscles. However, collagen
content and solubility were similar in destructured and normal porcine SM muscles.
Thermal shrinkage of IMCT occurred at approximately 65 °C in pork and poultry. It
occurred at lower temperature in light coloured muscles than in dark coloured muscles,
although the difference was not always significant. The onset and peak temperatures of
thermal shrinkage of IMCT were lower in destructured than in normal SM muscles, when
the IMCT from SM muscles exhibiting ten lowest and ten highest ultimate pH values were
investigated (onset: 59.4 °C vs. 60.7 °C, peak: 64.9 °C vs. 65.7 °C). As the destructured
meat was paler than normal meat, the PSE (pale, soft, exudative) phenomenon could not
be ruled out. The muscle fibre cross sectional area (CSA), the number of capillaries per
muscle fibre CSA and per fibre and sarcomere length were similar in destructured and
normal SM muscles. Drip loss was clearly higher in destructured than in normal SM
muscles.
In conclusion, collagen content and solubility and thermal shrinkage temperature vary
between porcine and poultry muscles. One feature in the IMCT could not be directly
associated with weakening of the meat structure. Poultry breast meat is very homogenous
within the species.
Acknowledgements
This study was carried out at the Department of Food Technology, University of Helsinki
during the years 2003-2009. Funds were obtained from the Finnish Ministry of
Agriculture and Forestry, Marie Curie Fellowship Association, Finnish Animal Breeding
Association, Finnish meat industry, and from the Helsinki University Foundation. Their
financial support is gratefully acknowledged.
I wish to express my warmest gratitude to my supervisor, Professor Eero Puolanne,
who gently guided me through this process of conducting scientific research and
constructing the Ph.D. thesis. I also would like to thank co-supervisor Dr. Marita
Ruusunen, with all the practical advice you gave me in scientific work and writing. You
made my life so much easier. I thank Dr. Marjo Halonen for the advice in writing. Dr.
Tuula Sontag-Strohm, Päivi Kanerva and Outi Brinck are acknowledged for helping me
with the protein analyses and Dr. Riitta Kivikari and Dr. Pirjo Salo-Väänänen for
methodological efforts in the beginning of this study.
Part of the experiments was carried out at the Ashtown Food Research Centre
(formerly known as The National Food Centre), in Dublin, Ireland. I would like to express
my gratitude to my Irish supervisor, Dr. Anne Maria Mullen for the kind co-operation in
the research we conducted and also for introducing me the scientific and systematic way
of working at Ashtown Food Research Centre. I also would like to thank Dr. Paul Allen
and Mr. Declan Troy for all the co-operation and help during my stay at the research
centre.
I am grateful for the reviewers of this thesis, Dr. Ian Richardson and Dr. Jacques
Lepetit. The discussing tone in the feedback, all the constructive criticism and advice
given were very helpful.
My special thanks go to Irja Korhonen whom I could always rely on when it was
matter of analysing samples. I also thank Kaisa Rautapalo, Kirsti Noro, Olavi Törmä,
Tapio Antila, Esko Petäjä-Kanninen (†), Maria Ylä-Ajos, Jutta Varis, all the post docs,
postgraduate students and others at the Department of Food Technology I was entitled to
work with, for all the advice and enjoyable discussions.
All my friends deserve my warm gratitude for understanding me during the process of
this thesis. Special thanks go to my hilarious volleyball team, no matter what, I was
always in good mood after the 2 hours training. All the support from families Kuikka and
Åberg during this process of thesis is highly appreciated.
Finally, I express my deepest gratitude to my parents Arja and Jussi for the endless
care and support during these years. Thank you also for understanding that despite this
process sometimes drew my attention far away from home, there is no place like home for
me. I am grateful also to my brother Juho and his fiancée Elina and to my sister Leena for
all the encouragement I got, and for all the non-scientific activities which you made me to
participate. All these elements, my family, balanced wonderfully this piece of scientific
work.
Helsinki, March 2009
Liisa Voutila
Contents
Abstract
3
Acknowledgements
4
List of original publications
9
Research input and authorship of papers (I-IV)
10
Abbreviations
11
1 Introduction
13
2 Review of the literature
17
2.1 Composition of connective tissue.............................................................................17
2.1.1 Proteins.............................................................................................................17
2.1.1.1 Collagen................................................................................................17
2.1.1.2 Elastin...................................................................................................19
2.1.2 Carbohydrate containing components of connective tissue..............................20
2.1.2.1 Proteoglycans.......................................................................................20
2.1.2.2 Glycoproteins.......................................................................................21
2.2 Collagen formation and degradation........................................................................21
2.2.1 Biosynthesis of collagen in fibroblasts.............................................................21
2.2.2 Degradation of collagen....................................................................................22
2.3 Cross-links in collagen.............................................................................................23
2.3.1 Formation of cross-links...................................................................................23
2.3.2 Immature cross-links.........................................................................................25
2.3.2.1 Aldimine and oxoimine........................................................................25
2.3.2.2 Disulphide cross-link............................................................................25
2.3.3 Mature cross-links.............................................................................................25
2.3.3.1 Pyridinoline..........................................................................................25
2.3.3.2 Ehrlich chromogen...............................................................................26
2.3.3.3 Histidino-hydroxylysinonorleucine......................................................26
2.3.3.4 Histidino-hydroxymerodesmosine.......................................................26
2.4 Structure of intramuscular connective tissue............................................................27
2.4.1 Perimysium.......................................................................................................27
2.4.2 Endomysium.....................................................................................................29
2.4.3 Basal lamina, a few special glycoproteins and a proteoglycan.........................31
2.5 The effect of cold storage on intramuscular connective tissue.................................33
2.6 The effect of post mortem treatments of meat on the properties of intramuscular
connective tissue and observed structure .......................................................................35
2.6.1 Heating..............................................................................................................35
2.6.2 Acidic marinating and bacterial collagenase marinating..................................36
2.6.3 Pressure treatments...........................................................................................36
2.6.4 Calcium chloride injections..............................................................................36
2.6.5 Thermal shrinkage temperature of collagen.....................................................37
2.7 General properties of intramuscular collagen in farm animals.................................38
2.7.1 Collagen content ..............................................................................................38
2.7.2 Collagen solubility............................................................................................40
2.7.3 Collagen cross-links..........................................................................................41
2.7.4 Chemical properties of collagen and meat structure.........................................42
2.7.5 Structure of intramuscular connective tissue and meat tenderness...................43
2.7.6 The effect of growth rate on the properties of connective tissue and observed
meat tenderness..........................................................................................................44
2.8 Motivation of the present work.................................................................................44
3 Objectives of the study
46
4 Materials and methods
47
4.1 Animals ....................................................................................................................47
4.2 Sampling...................................................................................................................47
4.3 Connective tissue analyses.......................................................................................47
4.3.1 Differential scanning calorimetry (DSC) .........................................................47
4.3.2 Selected SDS-PAGE analysis on the DSC samples ........................................49
4.3.3 Collagen content and collagen solubility .........................................................49
4.4 Meat quality traits.....................................................................................................50
4.4.1 The ultimate pH................................................................................................50
4.4.2 Colour ..............................................................................................................50
4.4.3 Drip loss ...........................................................................................................51
4.5 Muscle fibre properties in pork and poultry.............................................................52
4.6 Statistical analyses....................................................................................................53
5 Primary findings
54
5.1 Collagen content of porcine and poultry muscles.....................................................54
5.2 Collagen solubility in porcine and poultry muscles.................................................54
5.3 Thermal shrinkage of intramuscular connective tissue and ultimate pH in pork and
poultry.............................................................................................................................56
5.3.1 The ultimate pH and thermal shrinkage of the porcine muscles ......................56
5.3.2 The ultimate pH and thermal shrinkage of the poultry muscles ......................57
5.4 Properties of meat and muscle fibres in pork and poultry........................................57
5.4.1 Meat quality traits ............................................................................................57
5.4.2 Muscle fibre properties ....................................................................................58
5.4.3 The relationships between and within the meat quality and the connective
tissue parameters .......................................................................................................58
6 Discussion
60
6.1 Methodological and material aspects........................................................................60
6.1.1 Notes on sampling and materials......................................................................60
6.1.2 Notes on the methodology ...............................................................................60
6.2 The amount of collagen in different porcine muscles..............................................61
6.3 Thermal stability of intramuscular connective tissue in the porcine muscles..........62
6.3.1 Collagen solubility in the porcine muscles.......................................................62
6.3.2 Thermal shrinkage temperatures of the porcine muscles in relation to ultimate
pH...............................................................................................................................63
6.3.3 DSC sample moisture of the porcine intramuscular connective tissue.............64
6.4 Properties of the intramuscular connective tissue in poultry....................................65
6.4.1 The amount of collagen in the poultry muscles................................................65
6.4.2 Collagen solubility of the poultry muscles.......................................................66
6.4.3 Collagen solubility with animal age and anatomical location..........................66
6.4.4 Thermal shrinkage temperatures of intramuscular connective tissue in the
poultry muscles in relation to ultimate pH.................................................................68
6.4.5 DSC sample moisture and enthalpy of thermal shrinkage in poultry...............69
6.4.6 The possible role of non-collagenous molecules of intramuscular connective
tissue in the DSC sample moisture and weakened structure......................................69
6.5 Muscle fibre properties in pork and poultry.............................................................70
6.5.1 Muscle fibre CSA.............................................................................................70
6.5.2 Could the destructured appearance of meat be associated with weakness of
single muscle fibres?..................................................................................................71
6.5.3 Sarcomere length in destructured SM muscles.................................................72
6.6 Concluding remarks..................................................................................................73
7 Conclusions
74
8 Suggestions for further studies
75
9 References
76
Appendix A. Figures of combined data
90
List of original publications
This thesis is based on the following publications:
I
Voutila, L., Mullen, A. M., Ruusunen, M., Troy, D. Puolanne, E. (2007).
Thermal stability of connective tissue from porcine muscles. Meat Science,
76(3), 474-480.
II
Voutila, L., Ruusunen, M., Puolanne, E. (2008). Comparison of the thermal
characteristics of connective tissue in loose structured and normal structured
porcine M. semimembranosus. Meat Science 80(4), 1024-1030.
III
Voutila, L., Ruusunen, M., Jouppila, K., Puolanne, E. (2009) Thermal
properties of connective tissue in breast and leg muscles of chickens and
turkeys. Journal of the Science of Food and Agriculture 89(5), 890-896.
IV
Voutila, L., Perero, J., Ruusunen, M., Jouppila, K., Puolanne, E. Muscle
fibre properties and thermal stability of intramuscular connective tissue in
porcine M. semimembranosus. (submitted)
The publications are referred to in the text by their Roman numerals.
9
Research input and authorship of papers (I-IV)
This thesis is a summary of the research reported in the four (I to IV) appended papers. The
research input and authorship of the papers are as follows:
I L. Voutila, A.M. Mullen, M. Ruusunen, D. Troy, E. Puolanne (2007). Thermal stability of
connective tissue from porcine muscles. Meat Science, 76(3), 474-480.
The planning of this study was carried out by M.Sc. Liisa Voutila and Prof. Eero Puolanne in
Finland and by M.Sc. Liisa Voutila and Dr. Anne Maria Mullen in Ireland. The experimental
study, including empirical work, data analysis and preparation of the manuscript was carried out
by M.Sc. Voutila. The study was supervised by Dr. Anne Maria Mullen, Mr. Declan Troy, Dr.
Marita Ruusunen and Prof. Puolanne, all giving comments and suggestions and preparing the
manuscript.
II L. Voutila, M. Ruusunen, E. Puolanne (2008). Comparison of the thermal characteristics of
connective tissue in loose structured and normal structured porcine M. semimembranosus. Meat
Science, 80(4), 1024-1030.
The planning of this study was carried out by M.Sc. Liisa Voutila and Prof. Eero Puolanne.
Most of the experimental study, including empirical work, data analysis and preparation of the
manuscript was carried out by M.Sc. Voutila. Dr. Marita Ruusunen participated in the empirical
work. The study was supervised by Dr. Ruusunen and Prof. Puolanne, both giving comments
and suggestions and preparing the manuscript.
III L. Voutila, M. Ruusunen, K. Jouppila, E. Puolanne. Thermal properties of connective
tissue in breast and leg muscles of chickens and turkeys. Journal of the Science of Food and
Agriculture (submitted)
The planning of this study was carried out by M.Sc. Liisa Voutila and Prof. Eero Puolanne.
Most of the experimental study, including empirical work, data analysis and preparation of the
manuscript was carried out by M.Sc. Voutila. Dr. Marita Ruusunen participated in the empirical
work. The study was supervised by Dr. Ruusunen and Prof. Puolanne, both giving comments
and suggestions and preparing the manuscript. Dr. Kirsi Jouppila participated in preparation
of the manuscript by giving comments and suggestions.
IV L. Voutila, J. Perero, M. Ruusunen, K. Jouppila, E. Puolanne. Muscle fiber properties
and thermal stability of intramuscular connective tissue in porcine M. semimembranosus. Journal
of the Science of Food and Agriculture (submitted)
The planning of this study was carried out by M.Sc. Liisa Voutila and Prof. Eero Puolanne.
The sampling was carried out by M.Sc. Voutila and Mr. Juan Perero. The muscle fiber analyses
and corresponding data analyses were carried out by Mr. Perero. Other parts of the experimental
study, including empirical work, data analysis and preparation of the manuscript were carried
out by M.Sc. Voutila. The study was supervised by Dr. Marita Ruusunen and Prof. Puolanne,
both giving comments and suggestions and preparing the manuscript. Dr. Kirsi Jouppila
participated in preparation of the manuscript by giving comments and suggestions.
10
Abbreviations
ADD (muscle)
M. adductor
AH
aldolhistidine
BF (muscle)
M. biceps femoris
DHLN
dihydroxylysinonorleucine
DSC
differential scanning calorimetry
EC
Ehrlich chromogen
ECM
extracellular matrix
(fibre) CSA
muscle fibre cross sectional area
ER
endoplasmic reticulum
GAG, GAGs
glycosaminoglycan(s)
GAS (muscle)
M. gastrocnemius
GS (muscle)
M. gluteus superficialis
HHL
histidinohydroxylysinonorleucine
HHMD
histidinohydroxymerodesmosine
HLN, ∆HLN
hydroxylysinonorleucine, dehydrohydroxylysinonorleucine
HMD
hydroxymerodesmosine
HP
hydroxylysylpyridinoline
IMCT
intramuscular connective tissue
IS (muscle)
M. infraspinatus
ITL (muscle)
M. iliotibialis lateralis
λ
lambda, wave length
LD (muscle)
M. longissimus dorsi
LL
M. longissimus lumborum
LN
lysinonorleucine
LP
lysylpyridinoline
LT
M. longissimus thoracis
MAS (muscle)
M. masseter
MMP, MMPs
matrix metalloproteinase(s)
pHu
ultimate pH
PIF (muscle)
M. puboischiofemoralis
11
PJP
perimysial junctional plate
PS (muscle)
M. pectoralis superficialis
PSE
pale, soft, exudative meat
QF (muscle)
M. quadriceps femoris
SDS-PAGE
Sodium dodecyl sulphate polyacrylamide gel electrophoresis
SEM
scanning electron microscopy
SM (muscle)
M. semimembranosus
ST (muscle)
M. semitendinosus
TB (muscle)
M. triceps brachii
TIMP, TIMPs
tissue inhibitors of matrix metalloproteinases
Tm
temperature of denaturation of single collagen molecule
To
onset temperature of thermal shrinkage of intramuscular
connective tissue
Tp
peak temperature of thermal shrinkage of intramuscular
connective tissue
Ts
shrinkage temperature of intramuscular connective tissue
UTS
ultimate tensile strength
12
1 Introduction
The most important driving force to study the properties of intramuscular connective
tissue (IMCT) in meat technology has been the variability of texture of beef. Collagen
content (Mitchell & Hamilton, 1933; Dikeman et al., 1986), collagen solubility (Hill,
1966; Nishimura et al., 1999), thermal shrinkage temperature of IMCT (Tp) (Torrescano et
al., 2003) and collagen cross-links (Shimokomaki et al., 1972) have been measured in raw
meat in order to explain the changes in texture of cooked meat. However, it seems that one
single feature does not explain the variation of cooked meat texture. Pork and poultry have
also gained some attention but nowadays, pork or poultry toughness, as long as
commercial pig breeds and broiler chickens are concerned does not seem to be a problem.
The most studied muscles are the M. longissimus dorsi (LD muscle) in cattle, pigs and
sheep and M. pectoralis superficialis (PS muscle) in poultry.
A newer problem that seems opposite of toughness of meat has been recognised
(Figures 1A-C): cooked pork and poultry meat disintegrate. In extreme cases raw pork
(ham) and turkey (breast) meat can be peeled along the perimysial connective tissue
surrounding the muscle fibre bundles (destructured meat) (Figure 1A). Most of the slices
of cooked pork and turkey meat disintegrate too easily (Figures 1B and 1C). In normal
meat the strip will break when pulled by hand (Figure 1A). It seems that all chicken breast
meat provided nowadays is extremely soft, and when cooked, even mushy. However, the
selection criteria established by the industry for the destructured turkey breast meat has
not been effective to find the meat that will cause disintegration in cooked products
(Claus, 1998) The destructured raw meat and disintegration of cooked ham (M.
semimembranosus, SM muscle) has been reported by Franck et al. (1999). The
disintegration of cooked turkey meat has been reported by Swatland (1990) and overall
tenderness (mushy structure to the author’s opinion) of chicken meat has been reported by
Allen et al. (1998) and Makkonen (2003). The mushy structure of cooked chicken breast
meat can be perceived so that the need for chewing before swallowing the piece of meat is
minimal.
13
B
A
C
Figure 1 A) On the left, a strip of muscle fibre bundles in destructured meat. The segment
of a line shows the first part of the strip cut by knife. The second part of the strip of
muscle fibre bundles was generated by pulling the first part by fingers. On the right,
normal meat where strip broke when pulling by hand. B) Disintegrating cooked pork on
the plate. C) Disintegrating sliced cooked turkey breast meat.
The disintegrating cooked turkey breast meat products might, according to Swatland
(1990), be a consequence of too fast growth of the muscle fibres in relation to connective
tissue; thus the muscle fibres according to Swatland (1990) 'outgrow' of their connective
tissues in turkeys. Several authors have studied the possible causes of structural defects in
turkey muscles at single muscle fibre level in association with focal myopathy (Siller &
Wight, 1978; Sosnicki et al., 1988; Sosnicki et al., 1989), ischemia (Sosnicki et al.,
1991a; Sosnicki et al., 1991b) and with pale soft and exudative (PSE) meat (Pietrzak et
al., 1997; Dransfield & Sosnicki, 1999). However, to the author’s knowledge, the
properties of perimysial connective tissue in association with destructured raw meat and
disintegrating cooked meat products has not been reported. Fang et al. (1999) showed that
during pig’s growth the thickness of perimysial connective tissue increases approximately
13 % per month. In addition, Oshima et al. (2009) proved that the increase in muscle
collagen content is due to the development of perimysium. The hypothesis driving the
present study is that the weakening of meat structure is a consequence of gradual increase
in growth rate and decrease in slaughter age achieved by breeding in pigs and poultry
(Figure 2).
According to Hammond (1971) domestic pigs grow generally faster than wild boars
and the hind quarter is larger in adult domestic pigs than in adult wild boars. Franck et al.
(2002) reported that the destructured meat occurs in porcine LD and SM muscles, which
indeed are located in the large hind quarter of pigs. In addition, commercial chickens reach
1.5 kg body weight in 38 days whereas Thai indigenous chickens need 16 weeks to reach
the same weight (Wattanachant et al. 2004). Breeding for fast growth has also been
successful in turkeys (Havenstein et al., 2007).
14
Figure 2. A diagram of structurally weakened meat that in extreme cases can be seen as
structural defects in raw and cooked meat.
According to Hugenschmidt et al. (2007) the percentage of disintegrated slices in
Swiss ham industry is 3-7 % of total production. In a preliminary study Ruusunen et al.
(2008) on the cooking properties of hams made of destructured SM muscles it was shown
that the cooked ham made of destructured meat released more purge between the casing
and the product than the cooked ham made of normal meat (22 g/kg vs. 3 g/kg). O'Neill et
al. (2003) in turn, found that only 41% of the cooked hams made from carcasses classified
as PSE (measurements on LD muscles, though) were successfully sliced whereas 77% of
the hams made from normal carcasses produced acceptable slices.
Cooking of meat causes denaturation and shrinkage of proteins, of which myosin (29
% of total protein in general), collagen (the major protein of IMCT, 5 % of the total
protein) and actin (13 % of the total protein) (Lawrie, 1998, p. 59) are most often
discussed. According to Brunton et al. (2006) the denaturation of myosin occurs at
approximately 59 °C when measured from ground whole meat with ultimate pH (pH u) of
5.6-5.8. As different types of collagens located in the basement membrane, endomysium,
perimysium and epimysium, shrink at different temperatures, the collagen related fluid
loss according to Bendall and Restall (1983) covers the temperature range from 60 to 94
°C. This range includes also the core temperatures of the cooked ham products. Cooking
also causes fluid loss from meat. However, the most dramatic loss of water binding
capacity during cooking occurs already at approximately 35-50 °C (Hamm, 1972).
Franck et al. (2000) and Minvielle et al. (2003) reported that destructured meat of
hams comes from muscles with rapid pH decrease after slaughter, is pale in colour and it
has according to Minvielle et al. (2001) wider space between muscle fibres than normal
15
meat. According to Hugenschmidt et al. (2009) the destructured meat has less total and
insoluble collagen. Franck et al. (1999), Franck et al. (2003), Laville et al. (2005), Franck
et al. (2007) have concluded that the changes in the meat are of PSE type.
Swatland (1993) has pointed out that softness is the least understood part of pale, soft
and exudative meat. It is therefore possible that in the muscles where PSE most often
occurs (including porcine LD muscles and SM muscles and breast muscles in poultry),
weakening of IMCT might be one sign of the expressed 'softness'. In addition, Purslow
(1985) showed with cooked beef that when pulling meat perpendicular to the muscle
fibres, the fracture is detected at perimysial level. Indeed, the properties of IMCT and
possible relationship between muscle fibre characteristics and properties of IMCT in
destructured pork in general have not been extensively studied. Also the properties of
IMCT in poultry have gained too little attention. It was, thus considered important to carry
out a study in order to characterise the porcine IMCT from raw meat in order to suggest
possible candidates for differences in the cohesiveness of raw pork, to investigate the
physical and chemical properties of IMCT, and to study whether the properties of IMCT
were in association with the muscle fibre properties. Also poultry meat was investigated in
order to produce baseline type information on the state of poultry IMCT.
16
2 Review of the literature
2.1 Composition of connective tissue
Connective tissue consists of proteins, complex polysaccharides and water as different
mixtures depending on the type of tissue. In IMCT the main protein is collagen and
another important protein is elastin. Polysaccharides, which often are bound to protein,
form together with water the ground substance (of IMCT). The IMCT in meat is in three
hierarchical levels: epimysium is the layer surrounding the whole muscle, perimysium
surrounds bundles of muscle fibres, and individual muscle fibres are surrounded by the
endomysium (Bailey & Light, 1989; Alberts et al., 2002)
2.1.1 Proteins
The major protein in connective tissues is collagen and different types of collagen
comprise up to 25-30 % of total protein in mammal bodies (Bailey & Light, 1989).
Collagen forms rather inelastic fibrous structures, but also filamentous and network-like
structures in order to hold the structure of organs, to provide a framework for transforming
the force developed in the contractile filaments and to give the environment for cell
proliferation and growth (Bailey & Light, 1989; Kjaer, 2004). To date, 29 different types
of collagen have been identified (Brown & Timpl, 1995; Prockop & Kivirikko, 1995;
Fitzgerald & Bateman, 2001; Koch et al., 2001; 2003, Myllyharju & Kivirikko, 2001;
2004; Hashimoto et al., 2002; Sato et al., 2002; Banyard et al., 2003; Boot-Handford et
al., 2003; Pace et al., 2003; Söderhäll et al., 2007).
2.1.1.1 Collagen
The family of collagen types can be divided into fibre forming collagens (I, II, III, V and
XI), network forming collagens (IV, VIII and X), with special attention to type IV
collagen in basement membranes, fibre associated collagens with interrupted triple helix
(IX, XII, XIV, XIX and XXI), filamentous collagen (VI), anchoring fibres forming
collagen (VII), collagens with transmembrane domain (XIII and XVII) and the collagens
that have been only partly characterised (Bailey & Light, 1989; Prockop & Kivirikko,
1995). Most abundant types of collagen in IMCT are, however, the types I, III, IV, and V
(Light & Champion, 1984). Molecules of all collagen types are similar in basic structure
which is triple helix composed of three polypeptide chains (α-chains) (Figure 3). This
folding is not as common in protein as α helix or β sheet folding. Thus, in this way
collagen is special. The molecules of the most abundant fibre forming collagen, type I
(approximately 300 nm long), align next to each other forming striated fibrils (Figure 4).
These striated fibrils form the collagen fibres (Alberts et al., 2002; Bailey & Light, 1989;
Orgel et al., 2001).
17
Figure 3. Diagrammatic illustration of globular domains of the collagen molecule (Bailey
& Light, 1989).1
Figure 4. Illustration of the 4.4 D over-lap structure of the collagen fibre demonstrating
the derivation of the banding pattern of a negatively stained collagen fibre as seen in the
electron microscope (Bailey & Light, 1989).1
The difference between collagen types comes from the amino acid composition of the
α-chains and either the similarity or difference of the three α-chains in one molecule
(Bailey & Light, 1989; Prockop & Kivirikko, 1995; Alberts et al., 2002). The amino acid
compositions of the most abundant types of collagens in meat are shown in Table 1.
Hydroxyproline, the special amino acid for collagen, is not frequently found in other
proteins in animal tissues and so the amount of it has been used for decades as a tool to
determine the collagen content of tissues (Neuman & Logan, 1950; Bergman & Loxley,
1963; Alberts et al., 2002). Quantity, solubility and cross-linking of collagen has been
widely studied in relation to meat tenderness as will be discussed later (for example Hill,
1966; Shimokomaki et al., 1972; Fang et al., 1999).
Reprinted from Bailey & Light (1989). Connective tissue in meat and meat products, 1 st ed., Elsevier
Applied Science, London.
1
18
Table 1. Amino acid composition of the α-chains of the main collagen types in meat
(Bailey & Light, 1989).
Amino
acid
Type I
Type III
Type IV
Type V
α1(I)
α2(I)
α1(III)
α1(IV)
α2(IV)
α1(V)
α2(V)
Hyp
102
87
127
146
127
107
108
Asp
43
46
43
48
52
55
51
Thr
17
19
13
18
28
27
22
Ser
39
36
39
34
26
34
26
Glu
74
69
72
77
62
90
99
Pro
139
123
109
79
69
92
119
Gly
352
350
354
318
313
318
320
Ala
121
113
97
31
49
59
46
Val
21
34
14
30
24
30
19
Cys
-
-
2
-
-
-
-
Met
7
5
8
15
12
10
7
Ile
7
15
13
34
42
18
20
Leu
21
33
22
54
59
37
40
Tyr
2
4
3
5
6
-
1
Phe
13
12
8
24
37
12
12
Hyl
5
9
5
61
42
25
36
His
2
11
6
6
8
10
7
Lys
31
23
30
5
6
18
19
Arg
51
53
47
20
45
57
48
2.1.1.2 Elastin
Elastic fibres give the resilience, which is necessary in muscles, but also for example in
skin, blood vessels and lungs to recoil after transient stretch. Elastic fibres contain 90 %
elastin, another important connective tissue protein that forms a core. The core is
surrounded by microfibrils that are composed of distinct glycoproteins. Elastin is highly
hydrophobic and insoluble and it is formed from its soluble precursor tropoelastin via
cross-linking (Alberts et al., 2002; Mithieux & Weiss, 2005). Elastin is also very stable:
its estimated half life is about 70 years (Petersen et al., 2002). Elastic fibres can undergo
several cycles of stretching and recovery and the maximum extension of elastic fibres
before rupture varies between 100-220 % of the original length. The structure of elastic
19
matrices varies among tissues depending on the function of the tissues. However, the
elastic properties of the different tissues come mainly from elastin (Fung et al., 1993;
Mithieux & Weiss, 2005).
Because of the hydrophobic and insoluble nature of elastin, it is also highly resistant to
proteolytic attack. However, an elastin degrading enzyme, elastase, has been established.
Elastase is secreted from macrophages and granulocytes. The inactive form, pre-elastase,
is activated by a trypsin-like enzyme (Bailey & Light, 1989). To the author's knowledge
the exact mechanism of elastin degradation is not completely known.
2.1.2 Carbohydrate containing components of connective tissue
2.1.2.1 Proteoglycans
Proteoglycans are connective tissue macromolecules that provide a hydrated space around
cells. The physical behaviour of proteoglycans, however, is mostly determined by the
common properties of the glycosaminoglycans (GAGs) that are covalently bound to the
core protein (Ruoslahti, 1988). The core proteins of proteoglycans are an extremely
diverse group as only few small families of core proteins have been recognised but no
common structure for the proteins has been identified. Scott (1988) concluded that only a
very simple classification could be done on core proteins: i) small, ii) large and iii) very
large.
The GAGs are unbranched polysaccharide chains that consist of repeating disaccharide
units. One of the disaccharide units is always an amino sugar. A molecule to be titled as
proteoglycan has to have at least one GAG attached to the core protein, but the number of
GAGs bound to the core protein can vary between 1 and over 100 (Ruoslahti, 1988; Scott,
1988; Alberts et al., 2002). The GAGs can be divided into four types: 1) hyaluronan, 2)
chondroitin sulphate and dermatan sulphate, 3) heparan sulphate and 4) keratan sulphate.
Most of the extracellular proteins, such as collagens, fibronectin, and growth factors have
binding sites for GAGs. The GAGs also bind large amounts of water relative to their
weight; therefore also the matrix with GAGs can bind large amounts of water (Ruoslahti,
1988). However, for whole muscle, most of the water is located in the myofibrils between
actin and myosin filaments and thus, the properties of myofibrillar proteins determine the
water holding capacity of whole meat (Lawrie, 1998 pp. 219-220).
Weber et al. (1996) showed that the binding of core protein of decorin, a small
proteoglycan with chondroitin/dermatan sulphate GAGs, to collagen molecule promotes
the formation of correct type of collagen fibril. Velleman et al. (1996) reported that in
chickens with genetic muscle weakness an increased amount of decorin at 20 d of
embryonic development was followed by an increase in the content of mature collagen
cross-link, hydroxylysylpyridinoline (HP, see page 25), in breast muscles at six weeks
post hatch. Based on these, McCormick (1999) suggested that the expression of decorin
would regulate the content of HP via the regulation of alignment of collagen molecules in
during the formation of collagen fibril.
20
2.1.2.2 Glycoproteins
Glycoproteins differ from proteoglycans most clearly by structure and polysaccharide
chain content. Glycoproteins contain typically 1-60% carbohydrate by weight while
proteoglycans can contain up to 95% carbohydrate by weight. In addition, the
oligosaccharide chains of glycoproteins are relatively short and branched, whereas the
GAG chains of proteoglycans are long and unbranched (Alberts et al., 2002).
Fibrillin, a glycoprotein, has been found in muscles in perimysium and in
endomysium, but also in skin, vasculature, kidney and lung. It has been located in the
elastic fibres (Sakai et al., 1986). Fibrillin is one of the many glycoproteins that surround
the elastin core in the elastic fibres. It has been demonstrated that fibrillin together with
collagen VI contribute to the cell adhesion in vitro (Kielty et al., 1992).
Fibronectin, a large glycoprotein, is a dimer that is composed of two large subunits.
Each subunit has functionally distinct domains which are separated with regions of
polypeptide chain. In connective tissue, fibronectin is in insoluble fibrillar form and it acts
as a link between extracellular matrix (ECM) and actin filaments inside the cells (Hynes &
Destree, 1978; Bailey & Light, 1989; Alberts et al., 2002). Nakamura et al. (2007) showed
that in bovine M. masseter (MAS muscle), fibronectin was found in perimysium and in
endomysium. They also reported that the amount of fibronectin increased with increasing
exercise (the time of mastication of the feed) of the MAS muscle.
Laminin (laminin-1) is a large, asymmetric cross-shaped glycoprotein that consists of
three polypeptide chains (Figure 9). It is very flexible and acts as a binding protein in
ECM. Several isoforms of the polypeptide chains make a formation of a large family of
laminins possible (Alberts et al., 2002). Laminins are the major component of basement
membranes of skeletal muscles and critical for muscle fibre differentiation. Foster et al.
(1987) have demonstrated that laminin enhances the myoblast proliferation in vitro.
Laminins bind to for example integrins (see Figure 9). The two laminin variants,
2(α2β1γ1) and 4(α2β2γ1), also called merosin, are the most abundant laminin variants in
developing and adult muscles. These laminins are needed for muscle fibre survival (Belkin
& Stepp, 2000). However, the association between laminins and meat quality has not been
intensively studied.
2.2 Collagen formation and degradation
2.2.1 Biosynthesis of collagen in fibroblasts
Genes and each mRNA only code for one polypeptide chain of collagen. The typical
structure, triple helix of collagen (Figure 3) is formed in the endoplasmic reticulum (ER)
of a fibroblast after a series of enzymatic modifications of the single polypeptide chains
(pro-α-chains) (Bailey & Light, 1989). Hydroxylation of proline and lysine are carried out
by enzymes prolyl-4-hydroxylase and lysyl hydroxylase located in the ER. These
hydroxylations are essential for the stable triple helix formation and also for the
21
bifunctional cross-link formation between collagen molecules after fibre formation.
Glycosylation of some hydroxylysine residues is carried out by galactosyltransferase and
galctosylhydroxylysineglucosyltransferase (Bailey & Light, 1989; Prockop & Kivirikko,
1995). Also molecular chaperones such as protein disulphide isomerase, glucose
regulatory protein 94 and heat shock protein 47 assist collagen molecule formation
(Lamandé & Bateman, 1999).
After triple helix formation the procollagen molecules are secreted to the extracellular
space via Golgi apparatus. In the extracellular space enzymes pC-propeptidase and pNpropeptidase remove the procollagen propeptides from both ends of procollagen
molecules before the spontaneous alignment of the collagen molecules to collagen fibrils
(types I, II and III). Non-fibrous collagen types are not fully processed by the peptidases in
the extracellular space as are the fibrous collagens (Bailey & Light, 1989).
2.2.2 Degradation of collagen
Most of the proteinases cannot cleave collagen triple helix but fibroblasts have been
shown to synthesise collagen degrading enzymes which act under physiological
conditions. Collagen degradation occurs even before the molecules are secreted from the
cells. The amount of collagen degraded before secretion has been estimated as 15 % of the
collagen produced. This has been suggested to be a way of eliminating abnormal pro-αchains (Laurent, 1987; Bailey & Light, 1989). In non-mineralised matrices the collagen
degradation has four pathways: matrix metalloproteinase-dependent, plasmin-dependent,
polymorphonuclear leukocyte serine proteinase-dependent and phagocytic pathways
(Birkedal-Hansen et al., 1993). Degradation of collagen is a series of actions of enzymes
on the normally resistant triple helix and on the non-helical terminal regions (Bailey &
Light, 1989).
In vertebrates there are at least 20 different types of matrix metalloproteinases (MMPs)
(Nagase & Woessner, 1999). According to Kjaer (2004) the collagenases (MMP-1 and
MMP-8) initiate the degradation of collagen types I and III, whereas the gelatinases
(MMP-2 and MMP-9) break the collagen type IV and other compounds in the connective
tissue. However, for example MMP-2 is also able to degrade type I collagen, so the
degradation of connective tissue as a whole seems a very complex process (Kjaer, 2004).
The inhibitors of MMPs are called tissue inhibitors of matrix metalloproteinases
(TIMPs) (Woessner, 1991; Nagase & Woessner, 1999; Kjaer, 2004). Two of the four
identified TIMPs (TIMP-1 and TIMP-2) are able to inhibit the activities of all MMPs,
especially MMP-2 and MMP-9, respectively. The TIMPs are usually activated together
with the MMPs, most likely to control the degradation of collagen (Kjaer, 2004).
Balcerzak et al. (2001) reported activities of MMP-2 and MMP-9 and most likely the
inactive forms of MMP-1 and MMP-3 in bovine MAS muscles and SM muscles. In
addition, they reported mRNA encoding MMP-1, -2, -9, -16, TIMP-1, -2 and -3, meaning
that these enzymes are potentially associated with the degradation of IMCT.
Generally, collagen turnover rate is high during growth but reduces to a plateau by
maturity (Bailey & Light, 1989). Turnover time of collagen is very short for example in
22
periodontal ligament (only few days) but longer in skin and tendon (Laurent, 1987; Bailey
& Light, 1989). For a weanling rat muscle, collagen half life has been reported to be 45 d
(Rucklidge et al., 1992). Mays et al. (1991) showed that in rat M. gastrocnemius (GAS
muscle) collagen turnover had different characteristics at different ages. Collagen
fractional synthesis decreased from 4.9 %/day of 1 month old rats to 1.6 %/day of 6
months old rats. Old (24 months) rats had fractional collagen synthesis of 0.5 %/day. At
the same time the degradation of newly synthesised collagen increased from 26.2 % in 1
month old rats to 91 % in 24 months old rats (Mays et al. 1991). Mays et al. (1991)
suggested that this was due to the decrease of the activity of prolyl-4-hydroxylase
(discussed above in association with collagen synthesis), because the activity of prolyl-4hydroxylase decreases by age and because the under-hydroxylated collagen is susceptible
to degradation.
2.3 Cross-links in collagen
2.3.1 Formation of cross-links
When the collagen molecules have arranged to form the fibrils, cross-link formation
between the collagen molecules starts. Lysyl oxidase, an extracellular enzyme, converts
the lysine residues in N- and C-terminals of each α-chain to aldehyde groups. This enables
the formation of several types of lysine derived cross-links in fibrillar collagens (Figure 5)
(Eyre et al., 1984; Bailey & Light, 1989; Reiser et al., 1992). In general, tissues bearing
heavy loads contain predominantly cross-links formed via the hydroxyallysine route (Eyre
et al., 1984). The pyrrole and pyridinoline cross-links occur in about equal amounts as a
result of maturation of bone collagen. The maturation product of cartilage collagen is
mainly hydroxylysyl pyridinoline (Eyre et al. 2008).
23
Figure 5. Chemical pathway of cross-linking interactions initiated by lysyl oxidase that
predominates in skeletal tissue collagens. Bone collagen features both pyrrole and
pyridinoline cross-links whereas mature cartilage contains only pyridinolines. Lysines of
triple-helix and telopeptide origin are tracked by color to the mature cross-linking
structures.2
2
Reprinted from Methods, 45/1, Eyre,Weis & Wu, Advances in collagen cross-link analysis, Page 68,
Copyright (2008), with permission from Elsevier
24
2.3.2 Immature cross-links
2.3.2.1 Aldimine and oxoimine
If the hydroxylation of lysine residues before triple helix formation is at a low level, an
immature covalent cross-link the hydroxylysino-Δ-norleucine (ΔHLN, an aldimine)
connects two α-chains of collagen molecules. It is a major cross-link for example in the
skin collagen of young post-natal animals. The ΔHLN is formed between lysine aldehyde
and hydroxylysine and is completely stable in physiological conditions (Bailey & Light,
1989). Heating at +73 ºC for 10 min has been shown to completely cleave the aldimine
cross-links (Allain et al., 1978). If the hydroxylation of lysine residues before triple helix
formation is extensive or complete, another type of immature cross-link, oxo-imine
(hydroxylysino-5-ketonorleucine) is predominant. It is formed between hydroxylysine
aldehyde and hydroxylysine and is stable against heating. These cross-links can be formed
only in the non-helical domains of collagen molecules, thus, only a maximum of two
cross-links per α-chain are possible (Bailey & Light, 1989).
2.3.2.2 Disulphide cross-link
Collagen types containing sulphuric amino acid residues form disulphide cross-links. Out
of the most common collagen types (I, III, IV, and V, according to Light & Champion,
1984), the types III and IV form disulphide cross-links. In collagen type III, there are three
cysteine residues in the C-terminal end of the triple helix (as the molecule consists of three
α-chains), which in theory can form one intramolecular disulphide cross-link while the
third residue is free to form a cross-link with a cysteine residue from another molecule
(Bailey & Light, 1989). Also the type IV collagen forms intramolecular and
intermolecular disulphide cross-links. The type IV collagen structure is a sheet-like, multilayered network in the basal lamina and there are always N-terminal ends of four
molecules in a short region known as the 7S domain. The disulphide bonds are found
extensively in the 7S domain (Bailey & Light, 1989).
2.3.3 Mature cross-links
2.3.3.1 Pyridinoline
As the animals grow older, the nature of the covalent lysine derived cross-links of
collagen, discussed above, change, and conformation to more complex structures occurs.
This increases the strength and rigidity of collagen fibres (Bailey & Light, 1989).
Hydroxylysylpyridinoline (HP), which is a product of reaction between oxo-imine and
hydroxylysine and lysylpyridinoline (LP), which is derived from two residues of
25
hydroxylysine and one of lysine, has been found as 3-hydroxypyridinium based mature
cross-links (Eyre et al., 1984; Reiser et al., 1992, Eyre et al. 2008).
2.3.3.2 Ehrlich chromogen
Scott et al. (1983) proposed Ehrlich chromogen (EC) as a mature cross-link and Kuypers
et al. (1994) identified the molecular location of it. The EC is also a trivalent cross-link of
pyrrolic structure and two types of this cross-link are known: lysylpyrrole and and
hydroxylysylpyrrole. The hydroxylysylpyrrole is formed from oxo-imine, like the HP and
LP cross-links. The difference between HP and LP cross-links and hydroxylysylpyrrole is
that in HP and LP cross-links oxo-imine has reacted with hydroxylysine based aldehyde
whereas in hydroxylysylpyrrole oxo-imine has reacted with lysine based aldehyde. The
lysylpyrrole in turn, is a reaction product of hydroxylysine aldehyde, lysine aldehyde and
lysine (Kuypers et al., 1994, Eyre et al. 2008).
2.3.3.3 Histidino-hydroxylysinonorleucine
Histidino-hydroxylysinonorleucine (HHL) was characterised by Mechanic et al. (1987) as
a trifunctional cross-link in skin collagen. They reported that it formed between
hydroxylysine, lysine and histidine groups and that as a special feature it contained two
residues from helical regions and only one residue from the non-helical regions of
collagen molecules. For meat research, Avery et al. (1996) found no relationship between
HHL and meat texture in porcine meat. In general, this collagen cross-link has not been as
widely studied as HP and LP cross-links in the field of meat texture research (for example
Horgan et al., 1991; Smith & Judge, 1991; Young et al., 1994; Coro et al., 2000; Shiba et
al., 2000).
2.3.3.4 Histidino-hydroxymerodesmosine
Histidino-hydroxymerodesmosine (HHMD) has been shown to have a structure that is
formed from the derivative of allysine aldol by addition to a histidine, and thereafter
aldimine compound addition to a hydroxylysine. The exact mechanism of this cross-link
formation has been under debate (Bailey et al., 1974; Siegel, 1976; Eyre et al., 1984) and
possibly therefore HHMD has gained only little attention in meat quality research (Avery
et al., 1996; Ngapo et al., 2002).
26
2.4 Structure of intramuscular connective tissue
2.4.1 Perimysium
Perimysium is the layer of IMCT that surrounds the bundles of muscle fibres. It also
contains the large blood vessels and nerves of the muscle (Bailey & Light, 1989; Lawrie,
1998). The scanning electron microscopy photograph of Nakamura et al. (2003) shows
that perimysium can be divided into two different types: primary perimysium surrounding
the muscle fibre bundles and secondary perimysium surrounding the muscle fibre bundles
in larger scale (Figure 6).
PP
Figure 6. Low magnification SEM photograph in the longissimus muscle. P; perimysium,
E; endomysium (Nakamura et al. 2003).3 The PP; primary perimysium was added as an
interpretation from Fang et al. (1999). Bar = 100 µm.
Light and Champion (1984) and Nishimura et al. (1997) showed that perimysium
contains type I collagen as a major collagen type and types III, V and VI as minor
collagen types. Rowe (1981) demonstrated that well-orientated crimped collagen fibres of
criss-cross were the dominant feature of perimysium. The schematic diagram of bovine
3
Reprinted from Meat Science, 64/1, Nakamura, Iwamoto, Ono, Shiba, Nishimura, & Tabata,
Relationship among collagen amount, distribution and architecture in the M. longissiums thoracis and M.
pectoralis profundus from pigs, Page 46, Copyright (2003), with permission from Elsevier
27
muscle perimysium composed by Rowe (1986) shows that collagen forms an organised
network together with elastic fibres (Figure 7).
Figure 7. Schematic diagram of the perimysium of bovine muscles showing the relative
organisation of the crimped collagen fibres in the characteristic criss-cross pattern, the
thin elastic fibres (solid black) approximating the criss-cross of the collagen and the
longitudinally aligned thick elastic fibres at the junctional regions and in the thick
perimysial sheets (Rowe, 1986).4
The measurement of thickness of the whole perimysial sheath seems more challenging
than visualizing only the collagenous part of it, because like most connective tissues,
perimysium is a composite of different types of molecules and also water as discussed
earlier (page 17). Dehydration of sections removes water also from the connective tissue
part of the section and, for example in the method of Ohtani et al. (1988), only the
collagenous parts of IMCT remain for microscopic studies. Therefore the expression
'thickness of perimysium' should be used carefully when all the components of
perimysium are not visualised in the same image.
According to Iwamoto et al. (2001) it is possible to see actually more than one layer of
collagen fibres which run in different directions in different layers of secondary
perimysium. In the thick layers (3.4-6 μm in thickness) of secondary perimysium the
collagen fibres run longitudinally to the muscle fibre direction and in the thin layers (1.14
Reprinted from Meat Science, 17/4, Rowe, Elastin in bovine Semitendinosus and Longissimus dorsi
muscles, Page 310, Copyright (1986), with permission from Elsevier
28
1.3 μm in thickness) of secondary perimysium collagen fibres run circumferentially to the
muscle fibre direction.
In wild boars (carcass weight 60 kg), the thickness of the perimysial septa in M.
quadriceps femoris, (QF muscle), SM muscle and M. biceps femoris (BF muscle) was 2023 μm, when the measurements were done on cross-sections stained with haematoxylin
and eosin (Zochowska et al., 2005). In comparison, cross-sections of muscle of
commercial 6 months old pigs stained with Picro-Sirius method had a ST perimysium
thickness of 23 μm (Fang et al., 1999), whereas the thickness of perimysium in M.
pectoralis major (=PS muscle) and BF muscle of 38 d old commercial chickens was only
4-10 μm (Wattanachant et al. 2005a). According to Swatland (1990), the thickness of
perimysial septa of PS muscles in 15 week old turkeys was approximately 15 μm, when
the cross-sections had been stained with silver.
Passerieux et al. (2006), when studying bovine M. flexor carpi radialis, showed that in
addition to primary and secondary perimysia (Figure 6), perimysial collagen also had a
third hierarchical level that they nominated perimysial junctional plate (PJP). These PJPs
are particular domains that attach directly to the muscle fibres and underneath the PJPs
Passerieux et al. (2006) found accumulation of muscle fibre nuclei and subsarcolemmal
mitochondria.
Nishimura et al. (1997) showed that in addition to different collagen types also
fibronectin, laminin, dermatan sulphate proteoglycans, and decorin and chonrdoitin
sulphate proteoglycans were present in adult cattle perimysial connective tissue. Eggen et
al. (1997) located decorin in the junctions of neighbouring muscle fibres, in the adventitial
layer of a blood vessel, and in the perimysium. They also were able to show decorin at the
interface between perimysium and the muscle fibres with branches extending into
endomysium. Eggen et al. (1997) also showed that fibromodulin, a collagen binding
protein (Heinegård & Oldberg, 1989), and elastin could be clearly detected in cattle
perimysium. According to Pedersen et al. (1999) the total amount of the sulphated GAGs
of proteoglycans is approximately similar in different muscles but the relative proportions
of different GAGs are variable between muscles.
2.4.2 Endomysium
Endomysium is the connective tissue layer surrounding single muscle fibres (Bailey &
Light, 1989). Several authors have shown the structure and orientation of collagen fibres
in endomysium with a scanning electron microscope (for example Fang et al., 1999;
Iwamoto et al., 2001; Järvinen et al., 2002; Nakamura et al., 2004a; Roy et al., 2007). The
common feature of all the figures is that endomysial collagen forms a thin network around
the muscle fibres (Figure 8).
29
Figure 8. Cells invested by the endomysia in the longissimus muscle of pig.
Bar=10 µm (Nakamura et al., 2003).5
Järvinen et al. (2002) divided the endomysial collagen fibres into three separate
compartments: i) collagen fibres located on the surface of the muscle fibres and mostly
running longitudinally to the long axis of the muscle fibres, ii) collagen fibres connecting
two adjacent muscle fibres and running perpendicular to the long axis of the muscle fibres,
and iii) collagen fibres running around the intramuscular capillaries and nerves. The
collagen fibres around capillaries and nerves had contacts also with the collagen fibres
located on the surface of the muscle fibres, basement membrane of the capillaries and
perineurium. Figures of collagen fibres connecting the two adjacent muscle fibres
presented by Järvinen et al. (2002) were very similar to those presented by Nagel (1935)
although Nagel (1935) had had to draw by hand the fibres that he saw in the microscope.
Nishimura et al. (1997) reported that in addition to collagen types I and IV, adult
bovine endomysium contains laminin, fibronectin, decorin, chondroitin, dermatan and
heparan sulphate proteoglycans. Eggen et al. (1997) reported that aggrecan-like
proteoglycans and elastin were present also in endomysium. According to Velleman
(2002) decorin associates with fibrillar collagens and this interaction is needed for the
maturation of collagen fibres into larger fibre networks. Membrane associated
proteoglycans with heparan sulphate GAGs in turn, function as low-affinity co-receptors
for basic fibroblast growth factor. If heparan sulphate GAGs are removed, the basic
fibroblast growth factor no longer functions as an inhibitor of muscle differentiation, and
therefore the presence of proteoglycans with heparan sulphate GAGs is essential for
skeletal muscle development (Velleman, 2002).
5
Reprinted from Meat Science, 64/1, Nakamura, Iwamoto, Ono, Shiba, & Tabata, Relationship among
collagen amount, distribution and architecture in the M. longissimus thoracis and M. pectoralis profundus
from pigs, Page 49, Copyright (2003), with permission from Elsevier.
30
The images of perimysium and endomysium presented in most studies are in two
dimensions and the interpretation of the images relies on the staining of different
compounds. This makes the presentation of the interactions of different compounds in the
perimysium and endomysium very difficult. To the author's knowledge, only Nakamura et
al. (2007) have published three-dimensional images of perimysim and endomysium of red
meat showing them both together as whole in three dimensions, like the scanning electron
microscopic images from collagenous parts of perimysium and endomysium.
2.4.3 Basal lamina, a few special glycoproteins and a proteoglycan
Basal laminae are thin mats of specialised ECM that separate the muscle fibres from the
surrounding connective tissue (endomysium) (Figure 9). The term basement membrane is
often used to describe the composite of basal lamina and the connecting layer of collagen
fibrils. The functions of basal lamina include determining cell polarity, influencing cell
metabolism, organising the proteins in adjacent plasma membranes, promoting cell
survival, proliferation or differentiation and serving as specific pathways for cell
migration (Alberts et al., 2002).
31
Figure 9. A model of the molecular structure of a basal lamina. (A) The basal lamina is
formed by specific interactions (B) between the proteins type IV collagen, laminin, and
nidogen, and the proteoglycan perlecan. Arrows in (B) connect molecules that can bind
directly to each other. There are various isoforms of type IV collagen and laminin, each
with a distinctive tissue distribution. Transmembrane laminin receptors (integrins and
dystroglycan) in the plasma membrane are thought to organise the assembly of the basal
lamina; only the integrins are shown (Alberts et al., 2002).6
Integrins, a family of transmembrane glycoproteins, function as non-covalently linked
αβ heterodimers to mediate interactions from cell to cell and cell to matrix in wide variety
of cells (Figures 9, 10) (Hynes, 1992; Belkin & Stepp, 2000). They make connections
across the cell membrane to the cytoskeleton and activate many intracellular signalling
pathways (Hynes, 2002). According to Hynes (2002) 18 different α and 8 different β
subunits of integrin are known, and these can form at least 24 distinct integrins in
mammals. Integrins bind to for example laminin (Figure 9), paxillin and talin (Figure 10),
and the type α7β1 is one of the most abundant integrins in muscle fibres (Belkin & Stepp,
2000).
Most of the research on integrins is done for medical purposes, but Lawson (2004) and
Zhang et al. (2006) investigated integrins in pork in order to see if the degradation of
6
Reprinted, with permission, from Alberts, Johnson, Lewis, Raff, Roberts & Walter (2002) Molecular
biology of the cell, 4th ed., Garland Science New York, (NY), 06/02/2009 http://www.ncbi.nlm.nih.gov/books/
bv.fcgi?rid=mboc4.figgrp.3581 . Based on Colognato & Yourchenco (2000). Form and function: the
laminin family of heterotrimers, Dev. Dynamics, 18:213-234.
32
integrins post mortem was related to the amount of drip loss. According to Lawson's
(2004) hypothesis calpain mediated degradation of integrins early post mortem, before the
onset of rigor mortis, would lead to an increased drip loss in pork. This was supported by
the results of Zhang et al. (2006). However, Straadt et al. (2008) did not find a correlation
between immunologically stained integrin intensity and drip loss. They did their first
measurements after the onset of rigor mortis, though.
Figure 10. A diagram of the integrin adhesion complex on the cell surface. Integrins are
made up of two transmembrane domains (α and β). Also present in integrin containing
adhesion complexes are a large number of signalling proteins including talin, α-actinin,
focal adhesion kinase (FAK), vinculin (Vin), Paxillin (Pax), integrin linked kinase (ILK),
non-receptor kinase (Src), p130 docking protein (Cas) and adaptor protein (Crk)
(Lawson, 2004).7
Other glycoproteins, such as nidogen (Kohfeldt et al., 1998) and glypican (Velleman
et al., 2006) have not been studied intensively in the field of meat technology. Hannesson
et al. (2005) reported that post mortem tenderisation of beef as a result of 7 days cold
storage was associated with loss of a proteoglycan, perlecan (Larraín et al. 1997).
2.5 The effect of cold storage on intramuscular connective tissue
Immediately after slaughter the temperature in muscles is high. However, as the blood
circulation has stopped, the temperature decreases and the muscles become anaerobic.
This has not been proven to affect the properties of IMCT (Bailey & Light, 1989), but too
fast or too slow chilling are known to cause meat quality defects. Storage of meat at
temperatures of 0-5 °C for days to a few weeks (ageing) has been used to tenderise meat
structure and to improve eating quality. The mechanism of tenderisation of meat structure
during cold storage is well described, but the fine controls are still to be elucidated.
7
Reprinted from Meat Science, 68/4, Lawson, The role of integrin degradation in post-mortem drip loss
in pork, Page 560, Copyright (2004), with permission from Elsevier.
33
As the ability of muscle fibre sarcoplasmic reticulum to accumulate Ca2+ ions
decreases after slaughter, the post mortem proteolysis of myofibrillar proteins responsible
for at least partly the softening of meat has been suggested to originate from the increase
of the concentration of free Ca2+ ions in the myofibrillar compartment of muscles (Lawrie,
1998). Koohmaraie (1996) and Koohmaraie and Geesink (2006) have reviewed research
on the calpain mediated meat tenderisation so it will be introduced very briefly.
According to Etherington et al. (1987), Koohmaraie et al. (1987) Koohmaraie (1996) and
Koohmaraie and Geesink (2006) meat tenderisation occurs with the help of a Ca2+
activated proteolytic enzyme, μ-calpain which starts degrading the myofibrillar proteins
after the sarcoplasmic reticulum has released the Ca2+ ions after slaughter. Other enzymes
of the calpain family, m-calpain and calpain 3 do not seem to be involved to meat
tenderisation as strongly as μ-calpain (Koohmaraie & Geesink, 2006). Calpastatin, the
inhibitor of μ-calpain and m-calpain, has been shown to limit post-mortem proteolysis and
thus meat tenderisation. Another Ca2+ based theory about meat tenderisation is discussed
by Ji and Takahashi (2006). According to their theory Ca2+ ions bind directly to the
myofibrillar proteins, and no enzymes are involved with Ca2+ mediated meat tenderisation.
Already Takahashi (1992) suggested that it is the paratropomyosin that weakens the rigor
linkages during cold storage and thus softens the structure of meat. In addition, Takahashi
(1992) showed that in Ca2+ treated myofibrils connectin, a link protein between myosin
filament and Z-disk, was degraded due to the binding of Ca2+ ions to connectin. Ji and
Takahashi (2006) also pointed out that the pH and temperature optimums for the μcalpain and m-calpain are around 7.5 and 25 °C, respectively, which is very different from
the conditions in meat during cold storage (pH 5.5-5.7 and temperature 3-5 °C).
Dransfield (1997) modelled meat with a strip of aluminium foil referring to the muscle
fibre (stiff element) and with foam rubber referring to the ECM (extensible material). He
concluded that in meat the 'glue' (chemical bonds) between the stiff and extensible
material is extensible, too. During extension, fibre (foil in the model) breaks in several
places at regular intervals due to the stress transferred through the glue and the matrix.
Similar fracture patterns were found in meat after cold storage of days (beef), and also
after 24 hours of cooking (rabbit muscle) and after vacuum massage (processed pork)
(Dransfield, 1997).
Stanton and Light (1988) showed that cold storage of meat at 0-3 °C caused proteolytic
damage to perimysial connective tissue. They identified similar peptides in perimysium
from cold stored meat (14 days) and in perimysium treated with cathepsin. In addition,
Stanton and Light (1987, 1990) demonstrated that ageing of beef increased the solubility
of perimysial and endomysial collagen. This was supported by the results of Sylvestre et
al. (2002) on lamb, when studying the effect of cold storage on the lamb LD muscles and
SM muscles. Sylvestre et al. (2002) also found a positive correlation between the level of
active MMP-2 at slaughter and the increase in free hydroxyproline during the next 21 days
cold storage. They also reported a positive correlation between the level of active MMP-2
at slaughter and the collagen solubility at slaughter and at 21 d post mortem. Based on
this, Sylvestre et al. (2002) suggested that the activity of MMP-2 might play a role in the
collagen degradation during post mortem cold storage of meat and therefore eventually
affect meat quality.
34
Pierson and Fox (1976), in turn, reported that ageing of bovine LD muscle increased
meat tenderness but did not influence the amount of salt or acid soluble collagen. There
are conflicting results also about the effect of post mortem ageing on peak temperature of
thermal shrinkage (Tp) of IMCT: Judge and Aberle (1982) reported a decrease in T p of
IMCT in beef with 7 d ageing but Findlay and Stanley (1984) and McClain et al. (1970)
did not find a decrease in Tp of IMCT with up to 8 d ageing of beef. For the destructured
porcine SM muscles, there are no ageing studies reported. However, as the ageing was
associated with enhanced meat tenderness and lowered Tp of IMCT, at least in part of the
studies, and destructured meat is considered already too weak in structure, it would not
seem relevant to study the effect of ageing on destructured meat. Still, the possibility of
the above mentioned ageing effects to take place faster in the destructured than in normal
porcine SM muscles exists.
2.6 The effect of post mortem treatments of meat on the
properties of intramuscular connective tissue and observed
structure
2.6.1 Heating
When collagen is heated, denaturation, an irreversible conversion from the triple helix
form to a more random coiled peptide chain structure occurs. Denaturation causes the
collagen fibres to shrink to one fourth of the original length, which creates force. This is a
specific feature for collagen in comparison to other proteins. The thermal shrinkage of
collagen fibres in mammals occurs at approximately at 65 °C (Bailey & Light, 1989).
When this conversion from triple helix to random coiled form is complete, collagen
behaves according to the theory of rubber-like elasticity (Lepetit, 2008).
When meat is heated over 65 °C, a gradual increase in collagen solubility occurs
between 70 and 100 °C. At temperatures above 65 °C part of the collagen swells and
softens and finally disintegrates and forms gelatin (Lawrie, 1998). In the scanning electron
microscope, the denaturation and disintegration of perimysium and endomysium are seen
in the range 80-100 °C (Wattanachant et al., 2005b). According to Wattanachant et al.
(2005b) increasing the cooking temperature results also in the increase of the shear force
of cooked meat. Christensen et al. (2000) in turn, reported that as the shear force of
cooked meat increased, the tensile breaking strength of perimysial connective tissue along
the muscle fibre direction decreased at temperatures above 50 °C. The decrease of the
tensile strength of perimysial connective tissue above 50 °C would also fit in the
disintegrating cooked ham and disintegrating cooked turkey breast meat products: the core
temperature of such type products was 72 °C in the study of O'Neill et al. (2003). In
addition, the disintegrating meat is most obvious when the slices are across the muscle
fibre direction, and thus, the breakage of perimysium occurs along the muscle fibre
35
direction. According to Swatland (1990) and Hugenschmidt et al. (2007) the slices fall
apart.
2.6.2 Acidic marinating and bacterial collagenase marinating
Arganosa and Marriott (1989) and Aktas and Kaya (2001) reported that treating meat
or only IMCT with organic acids lowers the Tp and enthalpy (ΔH) of IMCT. Arganosa and
Marriott (1989) also reported an increased collagen solubility with acid treatment. Aktas
and Kaya (2001) suggested that the acid treatment of IMCT disrupted the non-covalent
intermolecular bonds (hydrogen bonds and dipole or ion-pair interactions and
intermolecular cross-links). This, according to Aktas and Kaya (2001), enhanced the
swelling effect of IMCT and decreased the Tp of IMCT. Also treatments of collagen with
different bacterial collagenases seem to decrease the Tp and ΔH of IMCT (Bernal &
Stanley, 1986; Tunick, 1988; Beltran et al., 1990). All the authors indicated that acidic
marinating or bacterial collagenase treatments of meat/IMCT could be associated with
improved meat tenderness. Destructured porcine SM muscles have not been studied after
low pH marinating, but interestingly, the destructuration occurs more often in muscles
with low ultimate pH (pHu <5.5) than in muscles with pHu considered normal (pHu 5.65.7). However, commercial marinating aims at pH of 4.5 in meat, at least in Finland, and
the studies on the pH effect on the IMCT/collagen are often approximately in the range
4.5<pH<7.0; thus the low pH effect on the IMCT/collagen is easy to detect as low Tp
when the range is so wide.
2.6.3 Pressure treatments
The reports about effects of pressure treatments on meat tenderness seem contradictory:
Beef hardness (measured with Texture Profile Analysis) has been successfully reduced
with high pressure treatment by Ma and Ledward (2004), but according to Macfarlane et
al. (1981) pressure treatment did not lower the Warner Bratzler shear force values. Ma and
Ledward (2004) suggested that the high pressure treatment accelerated the proteolysis
post mortem in meat but did not affect the structure of meat itself. The Tp of IMCT or
solubility of intramuscular collagen does not seem to be affected by pressure treatments
(Suzuki et al., 1993; Ma & Ledward, 2004).
2.6.4 Calcium chloride injections
The injection of CaCl2 solution into prerigor or postrigor meat has also been shown to
improve beef tenderness. Koohmaraie et al. (1987) showed that the activity of proteolytic
enzyme requiring high concentration of Ca2+, m-calpain, does not decrease during ageing
of meat, but the activities of μ-calpain, low Ca2+ concentration requiring proteolytic
enzyme, and calpastatin, the inhibiting enzyme for the calpains, decreased during ageing
36
of meat. Whipple and Koohmaraie (1993) concluded that with CaCl2 injection to 5 days
aged meat it was possible to activate the m-calpain to degrade the myofibrillar proteins
during the next 48 hours of ageing of meat and therefore improve meat tenderness. This
was supported by the results of Wheeler et al. (1997) and Jaturasitha et al. (2004).
Watanabe et al. (1996) in turn, showed that with a ZnCl2 injection to muscle after the
onset of rigor mortis it was possible to prevent the post-mortem proteolysis. According to
Koohmaraie et al. (1989), Wheeler et al. (1993) and Jaturasitha et al. (2004) it is possible
to improve meat tenderness with pre-rigor injections of CaCl2, but Jaturasitha et al. (2004)
also showed that the collagen content and solubility did not change with CaCl2 injection.
The CaCl2 marinating of IMCT extracted from meat lowered the Tp of the IMCT (Aktas &
Kaya, 2001).
The role of Ca2+ in the formation of destructured meat might be of interest. As the
porcine destructured meat has been described as PSE-like meat (Laville et al. 2005), could
the abnormally rapid flush of Ca2+ from sarcoplasmic reticulum to the myofibrillar
compartment of the muscle immediately after slaughter, which is considered typical for
formation of PSE meat, also affect IMCT in vivo?
2.6.5 Thermal shrinkage temperature of collagen
Most of the work conducted to study the thermal shrinkage of collagen has focused on
describing the thermal shrinkage temperature of collagen in different muscles of domestic
animals and through this knowledge has tried to understand the variability of texture of
meat especially of ruminants. As discussed above, marinating and ageing of meat have
been shown to lower the thermal shrinkage temperature of collagen. However, in these
studies the IMCT/collagen has been immersed in different solutions, but the effect of the
ultimate pH of meat where the IMCT/collagen is taken from, on the thermal shrinkage of
collagen has been rarely studied. Comparison of the results of different studies has to be
done carefully, as the size of the sample analysed and the heating rate of the samples
affect the results of the temperatures of thermal shrinkage of collagen.
As discussed above, denaturation causes collagen to shrink irreversibly. However, the
exact temperature of shrinkage of collagen (expressed as Ts, Tm or Tp in literature) is
determined by different factors. The thermal shrinkage temperature of collagen is low
when the intrafibrillar water content of collagen is high. The intrafibrillar water content in
turn, is determined by the cross-links of collagen: the more cross-links, the less the fibres
are able to swell and the higher the thermal shrinkage temperature. It seems that the
mature cross-links draw the collagen molecules closer together than the reducible crosslinks (Miles et al., 2005).
The thermal shrinkage temperature of collagen varies between muscles (King, 1987;
Torrescano et al., 2003) as does also the cross-link content of collagen (Horgan et al,
1991). The Tp differs also between animal species (McClain et al., 1971) and increases
with animal age (Horgan et al., 1991). This seems to be at least partly due to the
maturation of the collagen cross-links. However, low pH increases water binding of
connective tissue (Puolanne & Ruusunen, 1981) and also lowers the thermal shrinkage
37
temperature (Finch & Ledward, 1972; Aktas & Kaya, 2001). For destructured meat
measuring the Tp might provide information on the presence of this phenomenon in
connective tissue compartment of the porcine SM muscles.
2.7 General properties of intramuscular collagen in farm animals
2.7.1 Collagen content
The most common IMCT study in meat technology has probably been on beef, but also
IMCT of pigs, chickens and turkeys have gained some attention. From Table 2 it can be
seen that bovine muscles contain slightly more collagen than muscles of pigs, chickens
and turkeys although variation in collagen content between muscles is high. Hill (1966)
and Nakano and Thompson (1980) have shown that the muscle collagen content is highest
in young animals (cattle and pigs) and decreases as the animal grows older. Oshima et al.
(2009) showed that the variation of muscle collagen content is due to the variation in the
state of development of perimysium.
In poultry, Coro et al. (2000) reported that collagen content of PS muscles from laying
hens slightly increased by age but not after the age of 42 days. According to Nakamura et
al. (2004b) collagen content of breast muscles and leg muscles in broiler chickens is
highest at the ages of two weeks (14 days) and five weeks (35 days), respectively,
although the differences were small.
Table 2. Collagen content of most studied muscles in cattle, pigs, chickens and turkeys.
Reference
Animal breed
Animal
age/weight
Muscle
Collagen content
mg/g
Hill, (1966)
Fresian
9 weeks
ST
9.2
Hill, (1966)
Aberdeen Angus
15 years
ST
21.9
Goll et al. (1963)
Hereford
13-17 months
BF
9.4
Cross et al. (1973)
Hereford
10 months14 years
LD
6.1
Raes et al. (2003)
Limousin
Commercial, not
defined
SM
5.8
Raes et al. (2003)
Limousin
Commercial, not
defined
LD (LL)
6.9
Cross et al. (1984)
Charolais
6-18 months
LD
5.7
Cattle
38
Table 2 continued
Reference
Animal breed
Animal
age/weight
Muscle
Collagen content
mg/g
Li et al. (2007)
Simmental x Luxi
17-19 months
ST
7.0
Li et al. (2007)
Simmental x Luxi
17-19 months
LD
4.4
Wheeler et al. (2000)
White composite
66 kg skinned
carcass
LD
4.1
Wheeler et al. (2000)
White composite
66 kg skinned
carcass
SM
4.5
Boutten et al. (2000)
(LW x P x) x (LW x Commercial, not
LD), P76 x (LW x defined
LD)
SM
2.6
Boutten et al. (2000)
(LW x P x) x (LW x Commercial, not
LD), P76 x (LW x defined
LD)
BF
5.0
Correa et al. (2006)
Duroc x (Landrace x 115 kg live weight LD (LT)
Large White)
4.0
Therkildsen et al. (2002)
Duroc x Landrace x 68-79 kg carcass
Large White
weight
LD
3.1
Roy et al. (2007)
Red Cornish x New 80 d
Hampshire
ITL
4.7
Roy et al. (2007)
Red Cornish x New 80 d
Hampshire
PIF
3.3
Nakamura et al. (2004b)
Red Cornish x New 77 d
Hampshire
PS
3.3
Nakamura et al. (2003)
Silky
360 d
PS
2.9
Nakamura et al. (2003)
Silky
360 d
ITL
8.1
Nakamura et al. (2004a)
Commercial
53 d
PS
1.7
Liu et al. (1996)
Rhode Island Red
210 d
BF
8.3
Pigs
Chickens
Coro et al. (2000)
Laying hens
42 d
PS
5.2
Sakakibara et al. (2000)
White Leghorn
1-2 years
PS
6.5
BUT line 32
112 d
PS
5.8
Turkeys
Fernandez et al. (2001)
39
2.7.2 Collagen solubility
Collagen solubility has been shown to decrease as animals grow older (Hill, 1966; Nakano
& Thompson, 1980; Fang et al., 1999; Coro et al., 2000). From Table 3 it can be seen that
collagen solubility varies between muscles. In chickens the collagen solubility is higher
than in cattle, pigs or even in turkeys. However, the age of slaughter in relation to puberty
of the animal differs among the species. According to Fang et al. (1999) and Ruusunen et
al. (2007) the general slaughter age of pigs farmed for meat production is around 5-6
months which is slightly before the age of puberty in pigs (6-7 months, Hafez & Hafez,
2000). For cattle, the age of slaughter in Finland is approximately 18-20 months (Hannula
& Puolanne, 2004), which is also near the age of puberty of beef cattle (11-15 months for
heifers, until 18-24 months for bulls, Jainudeen & Hafez, 2000). The slaughter age of
turkeys and chickens in Finland is according to study III 110-111 days and 38 days,
respectively. Turkeys reach their puberty at the age of approximately 7-8 months and
chickens at the age of approximately 6 months (Froman et al., 2000).
In respect to the age of puberty and age of slaughter, commercial chickens are
slaughtered well before puberty in comparison to turkeys, pigs and cattle. This most
probably explains the high collagen solubilities in muscles of commercial chickens in
comparison to other species. As according to Oksbjerg et al. (2000) the growth rate of pigs
has increased during 20 years of intensive breeding, and according to Kerr et al. (2001)
also chickens are bred for a high growth rate, the animals reach their slaughter weight at
an earlier age than before. This would suggest that the collagen of fast growing animals is
less matured and more soluble than that of slow growing animals at the same slaughter
weight. Previously, Swatland (1990) suspected that for disintegrating cooked turkey meat,
the fast growing muscle fibres had outgrown their endomysial connective tissue. In the
present work a question is raised as whether the increased intramuscular collagen
solubility in pigs, chickens and turkeys can be associated with the weakening of the
structure of meat (Figure 1A).
Table 3. Collagen solubility in the most studied muscles in cattle (method of Hill, 1966)
Reference
Animal breed
Animal
age/weight
Muscle
Collagen solubility,
%
Smith & Judge (1991)
Holstein
USADA
A-maturity
SM
6.7
Li et al. (2007)
Simmental x Luxi
17-19 months
LD
19.0
Nishimura et al. (1999)
Japanese Black Cattle
9 months
LD
28.0
Nishimura et al. (1999)
Japanese Black Cattle
18 months
LD
19.0
Nishimura et al. (1999)
Japanese Black Cattle
24 months
LD
16.0
Nishimura et al. (1999)
Japanese Black Cattle
18 months
ST
15.0
Dikeman et al. (1986)
Aberdeen Angus
15 months
LD
13.4
Cattle
40
Table 3 continued
Reference
Animal breed
Animal
age/weight
Muscle
Collagen solubility,
%
Nakano & Thompson
(1980)
Hereford
20 months
TB
8.3
Fang et al. (1999)
Landrace
6 months
ST
24.0
Lebret et al. (1998)
Large White x (Large
White x Landrace)
5.5 months
LD
17.4
Lebret et al. (1998)
Large White x (Large
White x Landrace)
5.5 months
BF
18.2
Correa et al. (2006)
Duroc x (Landrace x
Yorkshire)
115 kg live
weight
LD (LT)
12.3
Liu et al. (1996)
Rhode Island Red
210 d
PS
40.0
Liu et al. (1996)
Rhode Island Red
210 d
BF
54.0
Coro, et al. (2000)
Laying hens
42 d
PS
25.7
112 d
PS
24.9 (heating
at 85 °C)
Pigs
Chickens
Turkeys
Fernandez et al. (2001) BUT line 32
2.7.3 Collagen cross-links
The conversion of reducible cross-links (aldimines and oxo-imines) to mature cross-links
(reported mainly in relation with HP and LP) has been shown to be complete as the
animals reach the physiological maturity (Bailey & Shimokomaki, 1971; Moriguchi &
Fujimoto, 1978; Horgan et al., 1991; Zimmermann et al., 1993; Young et al., 1994).
Zimmermann et al. (1993) concluded that training prevented the increase of concentration
of HP in rodent slow twitch muscles. They explained it by the training induced increase in
collagen synthesis and thus, training induced increase in the concentration of immature
cross-links in collagen. They did not measure collagen solubility, but the increase of the
proportion of immature cross-links in collagen might have been interpreted also as
increased collagen solubility.
However, Shiba et al. (2000) did not find an effect of training on the concentration of
HP cross-links in goats. A negative correlation between mature collagen cross-link, HP,
and collagen solubility has been shown to exist but it seems to be weak (Young et al.,
1994; Coro et al., 2000; Smith & Judge, 1991). For structurally weakened meat in general,
and especially in destructured porcine SM muscles and turkey PS muscles, the proportions
of different cross-links in collagen might be of importance, but their role in the mechanical
41
test described in Figure 1A is unclear. Is the content of the mature cross-links in collagen
the differentiating factor that becomes visual in structurally different meats?
2.7.4 Chemical properties of collagen and meat structure
Reducing the variation in meat tenderness and finding the optimal tenderness in meat have
been the ultimate goals in numerous studies; thus, most of the cited reports have this fixed
aspect also for connective tissue data. This means that the connective tissue data has
usually been plotted against tenderness parameters. However, the driving force of the
present study is quite opposite, meat that is already destructured in the raw state. To get an
idea of how different treatments affect connective tissue and if these treatments could be
candidates in predisposing the meat to the defect called ‘destructured meat’, it was
considered necessary to review the reports about connective tissue and meat tenderness. It
should be remembered however, that weakened structure has not been quantified by the
same instruments as meat tenderness, only by eye and by hand, mostly because of the
difficulties in allocating the measurements to the correct component(s) of meat. The most
important difference in the mechanical test (Figure 1A) for meat between destructured and
normal meat is, however, that peeling the strip from normal meat is not possible as the
strip breaks when pulled by hand.
According to Shimokomaki et al. (1972) the total collagen content is not important in
determining meat tenderness. They concluded that the relative proportion of thermally
labile and thermally stable intermolecular cross-links of collagen determine tenderness of
meat. However, Dransfield (1977) reported that total meat collagen content was more
important than heat soluble collagen in determining meat tenderness although neither
sensory panel nor mechanical tests could reliably differentiate the effect of connective
tissue on meat tenderness. For destructured pork, in comparison to normal pork, Minvielle
et al. (2001) showed that the collagen content was similar in both. Negative correlation
values between collagen content and tenderness measurements (either sensory panel or
instrumental) were found for example by Wheeler et al. (2000) in pork, by Torrescano et
al. (2003) in beef and by Young and Braggins (1993) in lamb, but reports by DeVol et al.
(1988) in pork, by Cross et al. (1973) and Dikeman et al. (1986) in beef did not show any
correlation between collagen content and tenderness. To the author's knowledge, there are
no reports about relationship between collagen content and meat tenderness in
destructured meat.
For collagen solubility, researchers seem to have concluded that high intramuscular
collagen solubility can be associated with tender meat in pigs (Fang et al., 1999), in cattle
(Moller,1981) in laying hens (Coro, et al. 2000) and in sheep (Young & Braggins, 1993),
but the relationship is not always clear from muscle to muscle (Boccard et al., 1979) or by
heat treatment (Dransfield, 1977) and can even be absent (Cross et al., 1973; DeVol et al.,
1988; Touraille et al., 1989). In destructured pork Minvielle et al. (2001) found higher
collagen solubility than in normal pork, but the difference disappeared when they made a
comparison between destructured and normal pork with similar pHu values.
42
As high content of collagen cross-link HP has been weakly related to low collagen
solubility (see above), also the relationship between collagen cross-links and meat
tenderness has been measured. Avery et al. (1996) did not find a correlation between any
collagen cross-link and pork tenderness. Ngapo et al. (2002) however, found that double
muscled cattle had less covalent cross-links, HHMD and EC in intramuscular collagen
than normal cattle. They also found that the relative amount of these covalent cross-links
and HHMD had weak positive correlations with Warner-Bratzler shear force measured
from raw meat. Coro et al. (2000) concluded that the increase in toughness of meat from
laying hens was associated with the increase in the content of HP cross-link of collagen.
Berge et al. (1997) made a more complex conclusion on beef: high total collagen content
determined the high content of mature cross-links of collagen and therefore finally also
high meat toughness. The Tp was not related to instrumental tenderness according to
Torrescano et al. (2003).
2.7.5 Structure of intramuscular connective tissue and meat tenderness
The role of IMCT in eating quality of meat has been recently reviewed by Purslow (2004,
2005) so only the main trends will be discussed here. Liu et al. (1996) found that the raw
meat shear force value (in kg) increased linearly with increasing thickness of perimysium
(Picro-Sirius staining) in chicken muscles. Similar types of results were reported by Fang
et al. (1999) in porcine ST muscles (Picro-Sirius staining). These were supported by the
results of Lachowicz et al. (2004) who found a positive correlation between width of
perimysial and endomysial space and instrumental hardness of muscles from wild boars
and domestic pigs. They also reported positive correlations between instrumental hardness
of muscles and muscle fibre cross sectional area.
Totland et al. (1988) found that the superficial parts of bovine ST muscles were more
tender than the deep parts of the same muscles when the tenderness was evaluated by a
sensory panel. They did not, however, find a difference in the volume fraction of
perimysial connective tissue between the superficial and deep parts of muscle. Therefore
Totland et al. (1988) suspected that the difference in tenderness between superficial and
deep parts of muscle was related to the elastin content as the superficial parts of muscle
contained less elastin than the deep parts of muscle. It is also possible that the difference
in the cooling rate between the superficial and deep parts (possibly heat shortened) of
muscle may have led to differences in the tenderness scores. For destructured pork,
Minvielle et al. (2001) reported that the extracellular space measured after hematoxylineosin staining was larger than in normal pork, but they did not find differences in the
properties of connective tissue when studying sections stained with Sirius red.
43
2.7.6 The effect of growth rate on the properties of connective tissue and
observed meat tenderness
The demand for efficiency in farmed animals has led animal breeders to aim for fast
growth and good feed efficiency. With these actions it has become possible to produce
heavier carcasses in shorter periods of time than before. However, to the author's
knowledge, collagen content or solubility have not been monitored during intensive
breeding lasting for decades. Animal diet manipulation during growth has also been
suggested as a tool to increase growth rate and also to improve meat tenderness. However,
Hall and Hunt (1982) did not find a difference between intensively-fed and maintenancelevel-fed cattle for instrumental meat tenderness, meat tenderness evaluated by sensory
panel, collagen content or collagen solubility. This was supported by the results of
McKeith et al. (1985) with beef and results of Correa et al. (2006) with pigs.
However, Therkildsen et al. (2002) reported that with a combination of restricted and
ad libitum feeding of pigs, so that feed was first restricted and then given ad libitum,
collagen solubility in two porcine muscles was higher than that of muscles from pigs on a
fixed diet. Kristensen et al. (2002) in turn reported that collagen solubility was highest in
the muscles of pigs which had been fed with a restricted diet for the whole period from
weaning to slaughter. In both experiments all the pigs were of same age but of different
weight at slaughter. In the study of Minvielle et al. (2001) the most severe destructured
pork and the highest collagen solubility in destructured porcine SM muscles were found in
the heaviest carcasses. They did not report the growth rate of the pigs. This leaves the
question about the relationship between high growth rate and occurrence of
destructuration in porcine SM muscles still open.
2.8 Motivation of the present work
To summarise, it has been shown that the properties of intramuscular connective tissue
(IMCT) and collagen content of meat determine the perceived meat structure and
tenderness to some extent but the results are variable. This is despite the systematics in
most studies in which the chemical analyses are performed on unaged meat and
instrumental tenderness and tenderness evaluated by sensory panel are done on meat that
has first been aged and then cooked to a certain core temperature. Similar variety of
studies in association with the destructured meat has not been yet been reported. Methods
for estimating tenderness used in the literature cited, such as Warner-Bratzler shear force,
Texture Profile Analysis and Allo-Kramer shear force, measure meat structure differently.
Increasing sample size in ultimate tensile strength (UTS) test, for example, increases the
UTS result (Lewis & Purslow, 1990). In addition, correlation coefficients are calculated
between parameters measured either from one muscle of many animals or several different
muscles of few animals. It seems that the muscle effect on calculations of correlation
coefficients has not always been controlled, so in theory, the correlation coefficients
obtained from data including several muscles of few animals may include information that
44
disturbs or enhances the correlation coefficients between for example IMCT
characteristics and meat tenderness measured by instrument or sensory panel.
The structure of IMCT obviously changes as animals grow older and it seems that the
increasing thickness of endomysium and perimysiun are related to increasing meat
toughness. However, as a reminder, the meat subjected to analyses in the present study is
far from tough. Weakened structure is a general term describing most of the pork and
poultry nowadays: cooked pork and poultry disintegrate too easily along the direction of
the muscle fibre bundles. In extreme cases the meat is actually destructured already as raw
meat. Destructured meat comes from commercial pigs and turkeys and soft meat from
commercial chickens, which are approximately the same age in the species group as the
normal meat producing animals. All these animals are bred for fast growth. Based on the
literature cited above it was impossible to find one trigger for the weakening of the
structure of raw meat. Therefore a new aspect in connective tissue research was
considered necessary and a few basic methods were chosen.
45
3 Objectives of the study
The ultimate goal of this work was to provide information on the properties of IMCT as a
background for the structural defect in pork and poultry. This was to see whether the
intramuscular connective tissue in pork and poultry had special features, both in content
and thermal stability, which could predispose the cooked meat from light coloured
muscles (SM and LD muscles in pigs, PS muscles in chickens and turkeys) to
disintegrative structure. A comparison with dark coloured muscles of different fibre types
(IS muscles in pigs, QF and ITL muscles in chickens and turkeys) of the same animals
was made. Investigations were supported by meat quality measurements (colour, drip loss
and pH) as the destructured meat often occurs in meat having pale colour and high drip
loss.
The purpose of this study was to investigate the properties of connective tissue in
different muscles of pig, chicken and turkey by focusing on:
1. Collagen content of different porcine muscles (I-II)
2. Solubility of collagen and thermal shrinkage temperature of IMCT in pigs in
association with meat destructuration (I-II)
3. Content and solubility of collagen and thermal shrinkage temperature of IMCT in
chickens and turkeys (III)
4. Possible connections between properties of muscle fibres and IMCT in pigs and
poultry (III, IV)
46
4 Materials and methods
The materials and methods of the present work are described in detail in the original
publications I, II, III and IV.
4.1 Animals
Animals for the present study were obtained from commercial abattoirs. For study I,
samples were taken from 13 pigs representing population A, Ireland, and from 14 pigs
representing population B, Finland. For study II samples were taken from 25 destructured
porcine muscles exhibiting pale colour and 25 porcine muscles exhibiting normal looking
colour and structure. For study III samples were taken from 12 chickens and 12 turkeys.
For study IV samples were taken from destructured muscles of 9 pigs and from normal
looking muscles of 25 pigs.
4.2 Sampling
For study I, approximately 100 g samples were taken from SM muscles, LD muscles
and IS muscles. The locations of sampling were: for the SM muscles the medial surface of
the muscle, for the IS muscles the deep part of the muscle which is on the scapula and for
the LD muscles, the steaks created by cutting the across the muscle between 3 rd and 4th rib.
For studies II and IV, approximately 100 g samples were taken from the deep part of the
SM muscles after visual assessment: Deep SM muscles were characterised as destructured
when a strip of muscle fibre bundles cut by knife could be pulled away by hand, and when
the colour of the same area was lighter than the average colour of the whole muscle
(Figure 1A) (II). For control group normal looking SM muscles, in which the strip cut by
knife broke when pulled by hand, were also taken for further sampling in both studies, II
and IV. For study III, approximately 100 g samples were taken from the middle part of PS
muscles (breast muscles) and as a combination of QF and ITL muscles (leg muscles). For
studies I-III samples were excised from the carcasses at 24 h post mortem and for study IV
samples were excised from carcasses at 48 h post mortem.
4.3 Connective tissue analyses
4.3.1 Differential scanning calorimetry (DSC)
The method for differential scanning calorimetry (DSC) analysis to determine the
onset and peak temperatures and the enthalpy of thermal shrinkage (To, Tp and ΔH,
47
respectively) of intramuscular connective tissue (IMCT) was adopted from Aktas and
Kaya (2001), King (1987) and Macfarlane et al. (1981) (I-IV). The samples for DSC
measurements were frozen 72 h post mortem (I-III) and at 48 h post mortem (IV) and
stored at -20 °C until analysed.
The buffer solution for extracting the myofibril proteins from IMCT was prepared by
mixing the solutions, 0.1 M KCl+0.02 M KH2PO4 (Aktas & Kaya, 2001) and 0.1 M
KCl + 0.02 M K2HPO4 (Macfarlane et al., 1981) to obtain a final buffer pH of 5.75. Three
replicates of each sample were prepared as follows: A 10 g piece of meat was treated for
about 10 s with Ultra-Turrax T25 (IKA® -Werke GmbH & Co, Staufen, Germany) in
100 ml buffer solution (temperature 4–6 °C) at 8000 rpm. The IMCT adhering to the
blades of the mixer was taken and the liquid discarded. The treatment with phosphate
buffer solution was repeated twice more on the IMCT sample. The IMCT was washed by
agitating it twice in 100 ml distilled water for 10 s and the sample was kept in distilled
water (less than 5 min) and quickly blotted on filter paper, which was the only control of
DSC sample moisture done by hand. Approximately 10 mg of sample was placed in a 40
μl aluminium DSC sample pan. The pans were hermetically sealed. The samples were
heated from 10 to 95 °C at a heating rate of 5 °C/min using TA 4000, DSC 30 from
Mettler Toledo AG, Greifensee, Switzerland. The instrument had been calibrated using npentane (−129.7 °C; 116 J/g), n-hexane (−95 °C; 151.8 J/g), mercury (−38.8 °C; 11.4 J/g),
distilled water (0 °C; 334.5 J/g), gallium (29.8 °C; 80 J/g) and indium (156.6 °C; 28.5 J/g).
An empty sample pan was used as a reference.
The To and Tp and ΔH were determined as demonstrated in Figure 1 (II) which is an
example of a thermogram obtained after the extraction described above. Figure 1 (II)
demonstrates that for the samples in the present study, there is only one major peak in the
thermogram, and that the peak is at approximately 65 °C. As according to Kijowski and
Mast (1988) the isolated IMCT produced one major peak in the DSC thermogram at 65.3
ºC, and according to Bailey and Light (1989) the shrinkage temperature of collagen from
mammals is around 65 °C, we considered this sufficient evidence that the peak
temperature (Tp) reported in the present study refers to the IMCT. In the Figure 1 of study
I the tangent of the peak in the thermogram is incorrect.
In study I, the instrument used for DSC analyses of samples from population A was
DSC2010 (TA Instruments, Dublin, Ireland) with a refrigerating cooling system (RCS),
calibrated with mercury (-38.8 °C; 11.4 J/g) distilled water (0 °C; 334.5 J/g) and indium
(156.6 °C; 28.5 J/g). Therefore an intercalibration for the DSC results in study I was
conducted. The intercalibration showed that instrument 2 gave about 1.5 °C higher
readings of Tp than instrument 1. For ΔH, instrument 1 gave about 0.5 J/g higher readings
than instrument 2. However, the standard deviations of the ΔH values in the muscle data
were greater than the difference between the instruments. In the statistical analyses the To,
Tp and ΔH data were handled as if they were obtained with the same instrument.
In studies II-IV, the lid of each sample pan was pierced after the DSC analysis, the
sample was dried overnight at 103 ºC and the moisture content of the sample was
expressed as a percentage of the original DSC sample weight (II) or g kg-1 (III and IV). In
studies III and IV the ΔH values were also expressed as joules per dry matter, thus the ΔH
values were standardised with the DSC sample moisture.
48
4.3.2 Selected SDS-PAGE analysis on the DSC samples
In study II, selected DSC samples obtained from phosphate buffer extraction were
subjected to SDS-PAGE analysis in XCell SureLockTM Mini-Cell (Invitrogen, Nupage,
Carlsbad, USA) according to the manufacturers instructions. The DSC samples
represented four different pHu values (5.30, 5.40, 5.68, 5.90), which covered the pHu range
of destructured and normal SM muscles in the present study. Approximately 40-50 mg of
each DSC sample was used for the analysis.
The SDS sample buffer contained 29 % v/v of SDS (10 % solution), 24 % v/v TrisHCl (0.5 M, pH 6.8), 47 % v/v glycerol and a trace of bromophenol blue. The sample
solution was made as follows: 95 % v/v of SDS sample buffer + 5 % v/v mercaptoethanol.
Mercaptoethanol was used to break the disulphide bonds of proteins. Each piece of DSC
sample (40-50 mg) was incubated in 1 ml of the sample solution for 2 min in boiling water
and then centrifuged for two minutes at 15800 g. The 5 ml injections of the supernatants
containing the soluble proteins were run with a constant current of 76 mA and voltage of
150 V for 82 min in a 12 % acrylamide gel. The the running buffer solution contained 1.5
% w/v TrizmaBase, 7.2 % w/v glycine and 0.5 % w/v SDS. Molecular weight marker
Novex® Sharp Protein Standard for 3.5-260 kDa (Invitrogen, California, USA) was used
in the gel. The gel was stained overnight with an aqueous solution containing 14 % v/v
Coomassie BB R-250 (0.5 %, diluted from Serva Blue R) and 43 % v/v TCA (12 %
solution) and washed in distilled water until the background of the gel was clear.
Most of the IMCT sample was insoluble in the SDS sample solution. Together with
there being one peak in the thermogram, characteristic of the denatured of collagen, it was
concluded that the samples referred to the IMCT.
4.3.3 Collagen content and collagen solubility
The method for total and soluble collagen determination was adopted from Fang et al.
(1999), Hill (1966), Kolar (1990) and Woessner (1961) (studies I-III). Five grams of
homogenized meat was put in a 50 ml centrifuge tube and Ringer’s solution (12 ml) (Fang
et al., 1999) was added and the whole mixed. The mixture was kept in a water bath at
77 °C for 65 min, with stirring every 15 min and centrifuged for 10 min at 3990 g (Sorvall
RC5C, rotor SS-34, DuPont Co., Wilmington, DE, USA). The supernatant was collected
and 8 ml of Ringer’s solution was mixed with the precipitate and centrifuged again for
10 min. After rinsing the precipitate the supernatants from the two centrifugations were
combined. The supernatants and precipitates of all samples were hydrolysed separately in
30 ml of 6 N H2SO4 in a Tecator Digestion System 20-1015 digester (Tecator, Inc.,
Herndon, VA, USA) at 110 °C for 16 h, the hydrolysates were diluted to 250 ml and
neutralised with 1 M NaOH (Woessner, 1961). The hydroxyproline content was
determined by a colorimetric reaction (Kolar, 1990): the hydroxyproline in the samples
was first oxidised with Chloramine-T in an aqueous buffer solution (pH 6.0) which
contained NaOH, citric acid, sodium acetate and 1-propanol. The colorimetric reaction
was carried out with dimetylaminobentsaldehyde (DMABA) in an aqueous solution
49
containing strong (70 %) perchloric acid and 2-propanol. Absorbance of the red colour
was recorded (Perkin–Elmer Lambda 2 spectrometer, Uberlingen, Germany) at λ=560 nm.
A standard calibration for hydroxyproline was established. The percentage of soluble
collagen was calculated from the hydroxyproline concentration in the supernatant. Total
collagen was calculated from the sum of the hydroxyproline concentration in the
precipitate and in the supernatant, using a conversion factor of 7.25 (Goll et al., 1963) and
expressed as mg/g of wet weight.
4.4 Meat quality traits
4.4.1 The ultimate pH
The ultimate pH (pHu) was recorded in duplicate with a pH meter Knick Portamess®
752 (Knick Elektronische Meßgeräte GmbH & Co, Berlin, Germany) attached to an
InLab® 427 electrode (Mettler-Toledo, Urdorf, Switzerland). In study I, the pHu of muscles
from population A was measured on a 1:10 w/v mixture of meat and Na-I-acetate solution
(5 mmol Na-I-acetate + 150 mmol KCl) from samples frozen at 24 h post mortem and
defrosted later for the pHu measurements. From the muscles of population B and in studies
II and III, the pHu was measured from homogenised samples 72 h post mortem. In study
IV the pHu was measured from homogenised samples 96 h post mortem. In study II ten
lowest pHu values were found among the destructured SM muscles and ten highest pHu
values were found among the normal muscles. This drove us to look whether there were
differences in the connective tissue properties between the low pHu and the high pHu SM
muscles (II, Table 4.).
4.4.2 Colour
In study I in population A, colour (lightness, L, and redness, a, was measured as a single
measurement on the medial surface of SM muscles at 24 h post mortem. The colour of LD
muscles of population A was measured on a one inch steak cut across the LD muscle 3 d
post mortem after 3 h blooming. During blooming the LD muscles were covered with a
piece of cling film. Colour meter used for both muscles was Miniscan XE Plus (Hunter
Associates Laboratory Inc., Virginia, USA).
In population B and in study II, colour (lightness, L* and redness, a*) was measured as
an average of four readings across sample surfaces of both sides of the sample with a
Minolta Chromameter CR-200 (Illuminant D65, Minolta Camera Co. Japan), calibrated
with a white plate.
In studies III and IV colour (lightness, L* and redness, a*, yellowness b*) was
measured with Minolta CR-400 (D65, Standard observer 2°, Konica Minolta Sensing,
Inc., Daisennishimachi, Sakai Osaka, Japan) that was calibrated with a white plate
50
(Y=93.9 x=0.3157 y=0.3322). The results presented were an average of four readings
across sample surfaces of both sides of the sample.
Colour measurements were done for population B (I) and in studies II and III at 72 h
post mortem. In study IV colour measurements were done at 48 h post mortem
immediately on the cut surfaces. Before the colour measurement meat had been hung in a
sealed plastic bag for drip loss measurement as described below.
Selection by colour lightness was successful as the mean lightness values (L*) of
destructured muscles were higher than those of normal muscles (II, Table 1).
4.4.3 Drip loss
Drip loss was determined by keeping one piece of meat per one muscle sample
(approximately 100 g in study II and 60 g in study IV) in plastic bags at +6 ºC for 2 d after
sampling (modification of the method of Honikel, 1998). In study III duplicate 45 g drip
loss samples were made. Meat was kept away from the drip so that the piece of meat was
sewn into upper part of a plastic bag and hung freely the give time. The plastic bag was in
close contact with meat during hanging. Drip loss was expressed as a percentage of the
initial weight. According to Locker and Daines (1974) the fluid loss during meat cooking
is almost exclusively along the fibres, thus it was thought that this was true also for raw
meat drip. The procedure of cutting of drip loss samples was standardised only within
studies, so the drip loss results of muscles are comparable only within the experiments
(Figure 11).
After the drip loss and colour measurements, most of the meat was homogenised and
pHu was measured as described above. Then pieces of whole meat and the homogenised
meat were frozen (-18 ºC) until differential scanning calorimetry analyses.
51
Colour measurement spots
Drip loss sample
M. adductor
Figure 11. Sampling for meat quality analyses from split porcine M. semimembranosus
(SM muscle). In study II colour was measured on both sides of drip loss sample at the two
spots marked in the figure. In study IV colour was measured on all the four spots marked
in the figure.
4.5 Muscle fibre properties in pork and poultry
Muscle fibre cross sectional area (fibre CSA) was measured from chicken and turkey
breast muscles (III) and from destructured and normal SM muscles (IV). About 0.5 cm x
0.5 cm x 1 cm pieces of the middle section of the chicken and turkey left breast muscles
were quickly frozen in liquid nitrogen at 24 h post mortem and stored at -80 °C until
analysed. The fibre CSA was measured without staining from cross sections of 14 μm
thickness cut with cryostat Reichert-Jung Frigocut 2800 E (Cambridge Instruments
GmbH, Nussloch, Germany) at -24 °C. The fibre CSA2 was calculated from the number of
fibres on a certain measured area and expressed as μm .
The pork samples (IV) were cut into approximately 0.5 x 0.5 x 0.5 cm pieces, frozen
immediately in liquid nitrogen, transported in dry ice and stored at -80 ºC until examined.
Cross sections (14 μm) of all the samples were cut at -24 °C in a cryostat (same as above)
and stained for capillaries according to Andersen (1975).
For sarcomere length and extracellular space measurements (IV), approximately 0.5 x
0.3 x 0.3 cm samples from porcine SM muscles (IV) were put into 3.5 % formaldehyde
solution containing also 39 mM NaH2PO4 and 46 mM Na2HPO4. Samples were stored at
+6 °C until analysed. Sarcomere length was measured according to Cross et al. (1981)
from the samples stored in 3.5 % formaldehyde solution at +6 ºC. Pieces of meat were
homogenised in fresh formaldehyde solution with Ultra-Turrax T25 (IKA® -Werke GmbH
& Co, Staufen, Germany) and the sarcomere length was measured using Novette™ 1507-0
52
helium-neon gas laser (Uniphase, Mantega, USA). Twenty-five measurements were done
on each muscle. Sarcomere length was calculated as follows:
Sarcomere length (μm) =
D = distance (mm) from the specimen-holding device to the screen
T = separation (mm) between the zero and the first maximum band
Samples for extracellular space were mounted in paraffin at 59 °C, cross sections of 4
μm thickness were cut using a microtome Leica RM 2165 (Germany), and stained with
haematoxylin and eosin (Mayer's method). Eight sections per muscle were examined.
Muscle fibre CSA and extracellular space were measured and the capillaries in the
sections were counted with an image analysis system using a computer program (KS300
(Carl Zeiss Vision GmbH, Hallbergmoos, Germany) attached to a microscope Olympus
BH2 (Olympus Optical Company, Tokyo, Japan). A magnification of 150 x was used.
4.6 Statistical analyses
Statistical analyses were carried out with SPSS (SPSS12.0.1 for Windows for studies I-III
and with SPSS13.0 for Windows for study IV). In study I a full factorial General Linear
Model was applied to the data in order to test the differences between three muscles and
also between the two populations. In studies II and IV a Mann-Whitney U-test was used to
test possible differences between destructured and normal SM muscles and Spearman's
correlation coefficients were calculated between the parameters. A linear regression
analysis including a constant term was applied to data in study IV. In study III the data
were analysed with Paired Samples T-test to test the possible differences between the two
types of muscles and with Independent Samples T-test to investigate the possible
differences between the avian species. Also Pearson’s correlation coefficients between
parameters in each muscle were calculated separately.
A linear regression analysis was applied to the combined data of studies II and IV in
order to investigate whether there were relationships between and within meat quality and
connective tissue parameters. A special interest was put on explaining the L* value, which
was one criterion of selection of the destructured samples. Also the possible candidates to
explain variation in To, Tp and DSC sample moisture were investigated. An analysis of
variance was used to test if the collagen content, collagen solubility (I-III), To, Tp and pHu
(I-IV) were different in porcine and poultry muscles.
53
5 Primary findings
5.1 Collagen content of porcine and poultry muscles
Collagen content of the porcine LD, SM and IS muscles varied from 2.8 mg/g to 4.8
mg/g (averages in Table 4) so that it was highest in the IS muscles, and lowest in the LD
muscles. The LD muscles from population A had higher collagen content than the LD
muscles from population B. Collagen content of the SM muscles and the IS muscles were
at the same level in populations A and B (I, Table 2). The destructured SM muscles had
similar collagen content to that of normal SM muscles (Table 4). When the SM muscles
with ten highest pHu values and ten lowest pHu values were compared, collagen content
was similar in both (II, Table 3). The SM muscles in study I had higher collagen content
than the SM muscles in study II (p<0.05, Table 4). The samples for study I were taken
from the medial surface and for study II from the deep SM muscles. Thus, the sampling
spot likely explains the difference between the studies I and II.
Collagen content of the chicken and turkey breast muscles and leg muscles varied from
2.5 mg/g to 4.1 mg/g, with the breast muscles having lower collagen content than the leg
muscles. In both muscle types, collagen content was similar in the chickens and in the
turkeys (averages in Table 4). Collagen content of the avian muscles included in the study
seemed very similar to the porcine muscles studied here.
5.2 Collagen solubility in porcine and poultry muscles
Collagen solubility of porcine muscles varied from 9.2 % to 17.3 % (averages in Table 4).
The highest collagen solubility was found in the IS muscles. Collagen solubility of the LD
muscles was higher than that of the SM muscles in population A, but in population B it
was similar for both muscle types (I, Table 2). Collagen solubility of the destructured SM
muscles was similar to the normal muscles, also when the SM muscles with ten highest
pHu values and ten lowest pHu values were examined (Table 4, II, Tables 3 and 4).
Collagen solubility of the SM muscles in study II was below 10 %, which was lower than
that of two populations in study I (Table 4, p<0.05).
In general, collagen solubilities of the chicken muscles were tremendously higher than
those of the porcine and turkey muscles, for example the collagen solubility of the chicken
breast muscles was higher than that of the turkey breast muscles. However, collagen
solubility of the turkey muscles was near those of the porcine muscles (Table 4).
54
Table 4. Main properties (mean ± s.d.) of meat and connective tissue in studies I-IV. Differences expressed at level p<0.05 by Least Significant
Difference -test.
LD (I)
SM (I)
IS (I)
SM L
(II+IV)
5.60±0.20ad
5.57±0.20ad
6.09±0.22b
5.42±0.07c 5.57±0.13d
(n=27)
(n=27)
(n=27)
(n=34)
59.0±1.2a
57.2±2.5b
59.7±0.9acf
60.2±1.4cefg 60.8±1.2defg
(n=14)
(n=14)
(n=14)
(n=34)
64.8±0.5a
63.5±1.3b
65.4±0.6cehi
65.3±0.7cdh 65.6±0.6efhi
(n=14)
(n=14)
(n=14)
(n=34)
(n=50)
DSC sample 67.9±4.3adegh
moisture, %
(n=14)
68.9±4.8acdegh
64.4±2.1bf
70.4±4.7c
67.5±3.7dg
(n=14)
(n=14)
(n=34)
(n=50)
Collagen
2.8±0.4ae
content, mg/
(n=27)
g
4.1±0.5bf
4.8±0.6c
3.4±0.6d
3.7±0.5dh
(n=27)
(n=27)
(n=25)
(n=25)
Collagen
13.5±2.1agh
solubility, %
(n=27)
12.4±2.8ag
17.3±2.7b
9.2±1.3c
9.3±1.4cd
(n=27)
(n=27)
(n=25)
(n=25)
pHu
To, °C
Tp, °C
SM C
(II+IV)
BR B (II)
(n=12)
BR L (III)
(n=12)
T B (III)
(n=12)
T L (III)
(n=12)
5.96±0.15e
6.62±0.12f 5.86±0.08eg
6.32±0.12h
61.4±0.3deg
60.9±0.6efg 60.2±0.9fg
60.9±0.7g
66.0±0.3fghi
66.4±0.6g
65.5±0.8hi
65.8±0.5i
66.8±2.2defgh 64.0±2.9f
67.3±2.7gh
68.3±1.5cdh
2.5±0.4eg
4.1±0.5fh
2.3±0.3g
3.9±0.5bh
30.3±4.6e
33.7±3.1f
13.4±2.2gh
15.0±1.3h
(n=50)
(n=50)
a,b,c,d,e,f,g,h,i
mean values with different letters as superscripts, differ from each other at p<0.05 level
The bold letters as superscripts indicate an opposite result in the original papers (I and III) in differences in pHu and in Tp because of different statistical analyses in the papers
and for this table of combined data.
LD = M. longissimus dorsi, SM = M. semimembranosus, IS = M. infraspinatus, L = destructured, C = control (normal structured), BR B = chicken breast muscle, BR L =
chicken leg muscle, T B = turkey breast muscle, T L = turkey leg muscle
5.3 Thermal shrinkage of intramuscular connective tissue and
ultimate pH in pork and poultry
5.3.1 The ultimate pH and thermal shrinkage of the porcine muscles
In the porcine muscles, the pHu value was highest in the IS muscle (Table 4). The LD
muscles had pHu values of 5.72 in population A and 5.49 in population B. The pH u value
of the SM muscle varied between 5.30 and 6.25 (I, II and IV).
The To, Tp and ΔH were analysed with different instruments, and the intercalibration
showed that a difference between populations existed: the readings for the same meat
sample of the instrument 2 for population B were 1.5 ºC higher than those of the
instrument 1 for population A. This meant that the difference seen in the Table 2 (I) was
actually larger than the numbers showed. The To and Tp of IMCT were higher in IS
muscles than in LD and SM muscles in population A. In population B a clear difference
existed only between IS and SM muscles (Table 4; I, Table 2). The To, Tp and ΔH were
higher in population A than in population B.
In the destructured SM muscles the pHu was lower than in the normal SM muscles
(Table 4). However, in study IV the mean values of pHu of the destructured SM muscles
and the normal SM muscles were 5.41 and 5.49, respectively (IV, Table 1). Thus, the
difference in pHu between the destructured and the normal categories was smaller in study
IV than in study II.
The To and Tp of the IMCT of the destructured SM muscles were similar to the normal
muscles, when the full data set was analysed (II, IV). However, in the study II, when the
IMCT of the SM muscles with ten lowest pHu values (all earlier classified destructured
muscles) and the IMCT of the SM muscles with ten highest pHu values (all earlier
classified as normal muscles) were analysed, it was found that the To and Tp of the IMCT
were lower in the SM muscles with ten lowest pHu values than in those with ten highest
pHu values. The Tp values of the IMCT of the destructured SM muscles were nearly
similar in studies II and IV. The Tp values of the IMCT of the normal muscles were
practically similar in the two studies (II, IV).
The enthalpy of thermal shrinkage (ΔH) of the IMCT varied in porcine muscles from
3.8 J/g to 8.9 J/g. The IMCT of the IS muscles had the highest ΔH in populations A and B.
However, the ΔH values of the IMCT of the LD muscles were similar to those of the SM
muscles in both populations: in population A approximately 7.1 J/g for both and in
population B approximately 4.5 J/g for both the LD and SM muscles (I, Table 2). Despite
the difference in the To and Tp of the IMCT between the destructured and the normal meat
the ΔH values of the IMCT were similar in study II. However, the IMCT of the
destructured SM muscles had slightly lower ΔH than the normal SM muscles in study IV
(IV, Table 1, 3.8 J/g vs. 4.2 J/g, p<0.05).
56
Interestingly, the DSC sample moisture was higher in the IMCT of the destructured
SM muscles than in the normal SM muscles in studies II and IV (p<0.05, Table 4). When
the ΔH results were standardised with the corresponding DSC sample moisture values, the
IMCT of destructured SM muscles had higher ΔH values per gram of dry matter (IV,
Table 1).
5.3.2 The ultimate pH and thermal shrinkage of the poultry muscles
The pHu of the chicken breast muscles was 5.96 and that of the turkey breast muscles was
5.86. The pHu values of the chicken leg muscles and the turkey leg muscles were 6.62 and
6.32, respectively (Table 4). Only the porcine IS muscles (I) had higher pHu values than
the chicken breast muscles (p<0.05, Table 4). The pHu of the leg muscles of the chickens
and turkeys was higher than that of the porcine muscles included in the studies I, II and IV
(p<0.05, Table 4).
The To of the IMCT was higher in the chicken breast muscles than in the turkey breast
muscles, but in the leg muscles a similar difference between the avian species was not
found. The Tp of the IMCT of the chicken leg muscles was higher than that of the turkey
leg muscles, but the difference between the chicken and turkey breast muscles was not
significant. The Tp of the IMCT of breast muscles was similar to the leg muscles in both
avian species. The To and Tp of the IMCT of the chicken and turkey breast muscles and
leg muscles were similar to or higher than those of the porcine muscles (Table 4).
The ΔH of the IMCT from the chicken and turkey muscles varied from 3.4 J/g to 4.8 J/
g. The ΔH of the IMCT from the chicken muscles was slightly higher than that of the
turkey muscles (p<0.05). When the ΔH values of the IMCT were again standardised with
the corresponding DSC sample moisture values, the difference between the chicken leg
muscles and the turkey leg muscles disappeared but stayed between the chicken breast
muscles and the turkey breast muscles. Only in the turkeys leg muscles had higher ΔH of
the IMCT and higher normalised ΔH values of the IMCT than the breast muscles (III,
Table 2).
5.4 Properties of meat and muscle fibres in pork and poultry
5.4.1 Meat quality traits
Colour (lightness L, redness a) values of the pork from population A were at the same
level as the colour (lightness L*, redness a*) values of the pork from population B (study
I, Table 1.). As the following colour measurements were done on scale L* a* b*, only
measurements done on this particular scale will be discussed to avoid confusion (studies
II-IV). The L* value of the porcine SM muscles varied from 45.3 to 54.4 (studies I, II and
IV). In the chicken breast muscles the L* value was 53.0 and in the turkey breast muscles
57
50.8 (III, Table 1). All the poultry muscles exhibited similar structure, so raw destructured
meat was not found in the poultry study.
The destructured muscles had higher drip loss than the normal muscles, 11.1 % and 2.4
% being the extremes of mean values for destructured and normal muscles, respectively
(II, p<0.001; IV, p<0.01). However, the difference in drip loss between the destructured
and the normal muscles was smaller in study IV than in study II. Drip losses of the
chicken and turkey breast muscles were only 1.7 % and 3.2 %, respectively.
5.4.2 Muscle fibre properties
Muscle fibre CSA in the chicken breast muscles was lower than in the turkey breast
muscles (III, Table 2, p<0.05).
The destructured SM muscles had a tendency to have longer sarcomeres than the
normal SM muscles (IV, Table 2, p<0.1). However, the fibre CSA in the destructured SM
muscles was similar to that in the normal SM muscles. Also capillaries/mm2,
capillaries/fibre and extracellular space in the destructured SM muscles were similar to
those in the normal SM muscles.
5.4.3 The relationships between and within the meat quality and the
connective tissue parameters
As paleness was one criterion of selection of the destructured and the normal samples
in studies II and IV, the lightness value (L*, II, Table 2; Appendix, Table A) was set as a
dependent value and other parameters were set as independent values on the first round of
analyses. After reducing the independent variables one by one it was found that in the
combined data of studies II and IV, the model with the pHu gave a good explanation of the
variation in L*, and the adjusted R2 was 0.545 (Figure A, Appendix, p<0.001).
When explaining the variation in the To and Tp of the IMCT, not the model with pHu,
but the DSC sample moisture was the best predictor. This was despite the positive
correlations between the pHu and the To of the IMCT and the pHu and the Tp of IMCT in
the study II with the data selected by the ten highest and ten lowest pHu values. For the To
of the IMCT explained by the DSC sample moisture the adjusted R2 was 0.133 (IV,
Figure 3, p<0.05), and for the corresponding Tp of the IMCT it was 0.263 (IV, Figure 4,
p<0.001). When the combined data from the studies II and IV were similarly analysed, the
DSC sample moisture did not explain the variation in the To and Tp of the IMCT: the
adjusted R2 was <0.1. This can be seen also in the Figures D and E (Appendix).
It was also investigated if any of the meat quality parameters (especially pHu or drip
loss) explained the variation in the DSC sample moisture (IV). However, this was not
found (IV).
When the combined data from the studies II and IV was analysed similarly, only the
pHu seemed to explain the variation in the DSC sample moisture (adjusted R 2 was 0.155),
58
but as can be seen in Figure C (Appendix), the relationship between the pH u and the DSC
sample moisture was poor.
Combined data for the SM muscles from the studies I and II revealed that collagen
solubility as the independent factor might explain the variation in the T o of the IMCT, as
the adjusted R2 from regression was 0.246 (p<0.001). However, from the Figure H
(Appendix) it can be concluded that the relationship is not very clear.
59
6 Discussion
6.1 Methodological and material aspects
6.1.1 Notes on sampling and materials
Few aspects considering materials and methods were considered relevant to be taken into
account. When studying animals produced for commercial use, it is difficult to determine
the exact age of the animals. They are farmed to grow to a certain weight, which despite
the achievements in animal breeding takes different times for different individuals. This is
an uncontrolled source of variation in the age-dependent properties of IMCT in the present
work, although a few days (poultry) or weeks (pigs) age difference would not be expected
to make a big difference. The DSC results from population A are not discussed in Table 4
because of the different equipment used in the analysis. The intercalibration in fact
showed that the results obtained for population A were different from those of population
B partly because of the equipment.
As the sampling spots in the SM muscles in the studies I and II were different it is a
potential source of difference in the collagen contents of the SM muscles in the studies I
and II. The sampling spot was changed due to practical reasons.
The combination of the two leg muscles in poultry (III) was necessary to make the
sample material sufficient for the analyses conducted. In addition, the chicken leg muscles
were very thin and small, it is thus, possible that some tendinous sheaths inside the QF
and ITL muscles were included in the collagen content and solubility analyses in larger
scale than in the IS muscles, although not meant to. However, to the author's opinion, this
was not the main reason for the results showing higher collagen content and solubility in
the dark coloured leg muscles than in the light coloured LD, SM and PS muscles.
Epimysium was always trimmed from the muscles.
For the study IV, difficulties in predicting the incidence of the destructured meat led to
the uneven number of the destructured and normal samples as the destructured pork can be
detected only in the boning hall after boning.
6.1.2 Notes on the methodology
The SDS-PAGE analysis of the DSC sample material revealed that the IMCT samples for
DSC contained many proteins which was not surprising. However, the thermograms
obtained included only one major peak, which based on the information presented in the
literature review was the denaturation temperature derived from the shrinkage of collagen.
To the author's knowledge, proteins other than collagen in the DSC sample have not been
reported to shift the peak either to lower or higher temperature. Comparisons to other
researchers' results on the To or Tp of the IMCT were not appropriate as the instrument
60
calibration, heating rate and size of the samples would likely cause differences in the
results.
The difference in the DSC sample moisture between the IMCT of the destructured and
normal SM muscles was smaller in the study II than in the study IV (3.5 % units and 7.5
% units, respectively). However, the difference in pHu between the destructured and
normal SM muscles was larger in the study II than in study IV. If only the meat pHu effect
is considered, these two findings are contradictory: low pH is known to increase the
swelling of IMCT/collagen and a large difference in pHu should have produced a large
difference in DSC sample moisture in studies II and IV. Now a small difference in pHu
was accompanied with a large difference in DSC sample moisture between the
destructured and normal SM muscles and vice versa (II, IV). In the both studies II and IV
the DSC sample moisture was higher in the destructured SM muscles than in the normal
SM muscles and slightly higher than in the LD and IS muscles (Table 4). As all the DSC
analyses on the porcine IMCT were done by the same person and using the same
equipment in the studies I, II and IV, except for the analyses on population A in the study
I, it is very unlikely that the high DSC sample moisture content in the destructured SM
muscles was an artefact in the method. In addition, the samples were actually standardised
in pH, when the myofibrils were extracted as the pH of the buffer solution was always
5.75 and not adjusted for the muscle pHu. After the extraction and the DSC analysis the
DSC sample moisture was still higher in the IMCT of the destructured SM muscles than in
that of the normal SM muscles.
The DSC sample moisture of the IMCT was generally at lower level in the study II
than in the study IV. The lower level of the DSC sample moisture of the IMCT in the
study II than in the study IV might be explained by the procedure of sampling: the DSC
samples (whole meat) were frozen from the drip loss samples which had lost fluid as drip
in the study II but in the study IV the DSC samples were frozen immediately after
sampling, at 48 h post mortem. It is also possible that the meat had lost fluid already
before sampling as the SM muscles were brought for sampling in a 200 litre meat trolley.
All the poultry samples were very homogeneous material and none of the samples
exhibited exceptional structure in raw state within one species.
6.2 The amount of collagen in different porcine muscles
Collagen contents of three porcine muscles were determined in order to obtain a 'baseline'
for the study on the comparison of the destructured and normal SM muscles. When
comparing the results of the present study on collagen content to the collagen contents of
bovine muscles (Table 2), it could be seen that the collagen contents of porcine and
poultry muscles in the present study were lower than that of bovine muscles. Old
Aberdeen Angus cows could have collagen content of 21.9 mg/g in ST muscles (Hill,
1966) and according to Raes et al. (2003) LD muscles from commercial Limousin cattle
contained collagen 6.9 mg/g in comparison to the collagen content of the porcine LD
muscles in the population B (I), 2.5 mg/g. Only the collagen content of the porcine IS
61
muscles 4.7-5.0 mg/g exceeded the collagen content of bovine LD muscles reported by Li
et al. (2007).
Collagen content of the SM muscles ranged from 3.4 mg/g to 4.1 mg/g (I, II). The
highest collagen content in the SM muscles was found in study I in populations A and B.
In the destructured SM muscles a collagen content of 3.4 mg/g was found, which was
similar to that of the normal SM muscles (3.7 mg/g) (II, Table 3). These collagen contents
were in between the results of Wheeler et al. (2000) and Boutten et al. (2000) (Table 2)
but lower than that reported by Minvielle et al. (2001). Hill (1966) and Nakano and
Thompson (1980) have shown that in bovine muscles the collagen content is highest in the
neonates and very young animals and thereafter decreases due to the growth of
myofibrillar part of the muscle fibres. Although the collagen content of the destructured
and normal porcine SM muscles was similar, the role of collagen content in the
destructured meat could not be ruled out: During the past 40 years the animal breeding has
increased the growth rate of the farm animals, and it is possible that the gradually
increased growth rate has gradually decreased the strength of IMCT in meat, and, as
Swatland (1990) noted, muscle fibres outgrow their connective tissue sheath.
6.3 Thermal stability of intramuscular connective tissue in the
porcine muscles
6.3.1 Collagen solubility in the porcine muscles
In the study of Minvielle et al. (2001) collagen solubility of SM muscles was higher than
in the present study (15.8-17.8 % vs. 9.2-14.2 %). In the most obviously destructured SM
muscles (pHu on average 5.46) they reported higher collagen solubility than in the normal
SM muscles (pHu on average 5.74), (17.8% vs. 15.8%, p<0.05). They did not however,
find a difference between the normal SM muscles with low pH (pHu 5.48) and the most
obviously destructured SM muscles. As Laville et al. (2005) suspected that destructured
SM muscles were similar to PSE muscles; the findings of Minvielle et al. (2001) were in
agreement with the report of McClain et al. (1969) who found that collagen solubility of
epimysium was higher in PSE muscles than in normal muscles. However, collagen
solubility of the destructured SM muscles in the present study was similar to that of the
normal SM muscles (II, Table 3). This disagreed with the results of Minvielle et al. (2001)
but could be partly because of the classification system, as only two classes (destructured
and normal) were in the present study and Minvielle et al. (2001) ranked SM muscles into
four different classes according to the severity of the destructured meat.
When looking at the collagen solubility of the most studied porcine muscle, the LD
muscle, at the approximate commercial age of slaughter from the work of DeVol et al.
(1988) until the present study, collagen solubility varies from 10 % to 20 %: DeVol et al.
(1988) 10.6 %; Seideman et al. (1989) 9.6 %; Koohmaraie et al. (1991) 14.1-15.7 %;
Lebret et al. (1999) about 20 %; Wood et al. (1996) about 14.2 %; Correa et al. (2006)
62
13.4 % and study I, about 13.6 %. Oksbjerg et al. (2000) showed that the daily weight
gain of Danish pigs had increased from 670 g/day to 958 g/day during the twenty years of
selection for growth rate. As a percentage of the initial growth rate the increase is 43 %.
All muscles could be assumed to have increased their growth rate to some extent, although
the proportion of different muscles of the carcass weight changes during the growth of the
pigs. According to Lawrie (1998) the muscle proportions of carcass weight are in neonates
as follows: LL 2.6 %, LT 3.6 % and neck muscles 4.6 %. In 100 day old pigs the
corresponding proportions are for LL 3.9 %, for LT 5.4 % and for neck muscles 4.8 %.
Thus, the valuable loin part of the carcass grows more than the neck muscles. These
favour the thought that in the fast growing animals the fast growing muscles exhibit
weakened structure more easily than before because the animals slaughtered are younger
than before. This could be due to the change in the properties of the IMCT over a long
period of time, although in the present work, difference in collagen solubility was not
found between the destructured and normal meat.
Interestingly, as discussed above, the level of collagen solubility in porcine LD seems
to have moved from around 10 % to around 14 % and this change is 40 % of the initial
collagen solubility. It can be speculated, whether this shows in the quality of cooked meat,
but at least, the increases in pig growth rate and collagen solubility as percentages seem
very similar. From the report of Oksbjerg et al. (2000) and the proposed increase in
collagen solubility in LD muscles during the past decades (above) it could be concluded
that nowadays pigs are slaughtered at younger ages than before, and possibly therefore the
collagen solubility seems to have increased, as collagen is less mature than in commercial
pigs few decades ago. The possible breed and age induced differences in collagen
solubility were ignored in this comparison, as the age of the pigs was available either as
slaughter weight or as days. However, the method for collagen solubility was the same in
all the references cited above. From other porcine muscles, for example the SM muscles,
similar comparison was not possible as very limited data was available.
6.3.2 Thermal shrinkage temperatures of the porcine muscles in relation to
ultimate pH
As the destructured SM muscles in the present study exhibited the ten lowest pH u values,
the control muscles exhibited the ten highest pHu values (II) and Franck et al. (2000) and
Minvielle et al. (2001) concluded that the destructured SM muscles had lower pHu than
normal muscles, it was considered necessary to investigate the differences between the
SM muscles exhibiting the ten lowest pHu values and the ten highest pHu values (II). The
positive correlation between pHu and the To and Tp of the IMCT among the SM muscles
exhibiting the ten highest pHu values and the ten lowest pHu values indicated that a low
pHu was associated with the decrease in the shrinkage temperature of IMCT in the
destructured SM muscles (II). However, when the combined data of pHu and the Tp of the
IMCT from the studies II and IV were plotted, it could be seen that the relationship
between pHu and the Tp of the IMCT was poor (Figure F, Appendix).
63
Several studies have shown that low pH marinating of collagen alone or IMCT causes
a decrease in the To (Finch & Ledward, 1972; Horgan et al. 1991) and Tp of the
IMCT/collagen (Judge & Aberle, 1982; Horgan et al. 1991; Usha & Ramasami, 1999;
Aktas & Kaya 2001) and an increase in collagen solubility (Arganosa & Marriott, 1989;
Burke & Monahan, 2003). However, none of the mentioned studies were on porcine
IMCT. McClain et al. (1971) did analyse porcine IMCT and found that the IMCT from
porcine LD muscles had lower Tp than that from bovine or ovine LD muscles.
Unfortunately they did not report the pH of the muscles at the time of sampling. However,
marinating meat, at least in Finland, aims at pH of approximately 4.5, which is clearly
different from the pHu of meat (usually 5.5-5.8). The swelling effect of acidic marinade on
the connective tissue is likely detected more clearly as lowered To and Tp of the IMCT
than the swelling effect caused by the approximately 0.3 pH units lower pH u in the
destructured than normal meat. Therefore it is possible that the pHu of the meat might have
participated to the lowering of the To and Tp of the IMCT in destructured pork, but the
actual cause of the destructured pork could be somewhere else.
6.3.3 DSC sample moisture of the porcine intramuscular connective tissue
As the freezing of the DSC samples for the studies II and IV were at different time (72 h
post mortem and 48 h post mortem, respectively), it means that the samples of the study II
had likely proteolysed further than the samples for the study IV. Although Findlay and
Stanley (1984) and McClain et al. (1970) did not find a decrease in the Tp of the IMCT
with up to 8 d cold storage of beef, Judge and Aberle (1982) reported a decrease in the Tp
of the IMCT in beef with 7 d cold storage. In addition, according to Koohmaraie et al.
(1991) the proteolysis in pork occurs faster than in beef or lamb. As the destructured and
normal samples were handled with similar manner, it is, thus, possible that different or
faster type of proteolysis occurred in the destructured SM muscles in comparison to the
normal SM muscles. However, the activities of proteolytic enzymes were not investigated
in the present study.
Assuming that different type or faster proteolysis occurred in the destructured than in
the normal SM muscles (II), the speculation on the proteolysed molecules should take the
non-collagenous molecules into account. Although the formation of the destructured meat
according to other researchers (Le Roy et al., 2001; Franck et al., 2000; Minvielle et al.,
2001; Laville et al., 2005) seems to follow the traditional steps of formation of PSE
condition, the main protein of connective tissue, collagen, may or may not be affected,
measured as collagen solubility, in the destructured meat (II; Minvielle et al, 2001).
However, McClain and Pearson (1969) suspected that either the amount or the properties
of the ground substance of epimysium in PSE meat was altered in comparison to normal
meat. According to Hamm (1972) the pH at which the water holding capacity of meat is at
its lowest, is 5.0, it would seem logical that the myofibrils of the samples with the low pHu
in the study II had released more water to the ECM than the myofibrils of the high pH u
samples.
64
Miles et al. (2005) showed that the state of hydration mainly determines the Tp of
collagen, but in the study IV the Tp of the IMCT of the destructured SM muscles and the
normal SM muscles were similar although the DSC sample moisture was higher in the
destructured SM muscles than in the normal SM muscles. Therefore, a favourable thought
is that the moisture content of the IMCT that according to the SDS-PAGE analysis (II)
contained also other than insoluble proteins had increased in the destructured porcine SM
muscles because of the change of the properties of the IMCT early post mortem.
6.4 Properties of the intramuscular connective tissue in poultry
6.4.1 The amount of collagen in the poultry muscles
The chicken breast muscles had collagen content of 2.5 mg/g which seemed similar to the
collagen content of the porcine LD muscles (Table 4). However, when comparing the
collagen contents of the breast muscles from other chicken breeds (Table 2), it was found
that the collagen contents of the chicken and turkey breast muscles in the present study
were among the lowest ones. Only Nakamura et al. (2004a) reported lower collagen
contents in chicken breast muscles than those in the present study. As Coro et al. (2000)
did not find a change in collagen content in laying hens after 42 days of age and
Nakamura et al. (2004b) reported highest collagen contents in breast muscles from Red
Cornish x New Hampshire cockerels at the age of two weeks (14 days) and five weeks (35
days), the differences between studies were most likely due to the differences between the
breeds.
The higher collagen content of the poultry leg muscles in comparison to the breast
muscles was an expected result. Although the distribution of different muscle fibre types
was not measured in study III, it is known that the breast muscles have thicker muscle
fibres than the leg muscles (Ono et al., 1993; Papinaho et al., 1996). In addition, Oshima
et al. (2009) showed that despite the differences in collagen content and muscle fibre
diameter the structure of endomysium was similar in different muscles from differently
bred pigs. They also reported that the perimysium was least developed in a white muscle
(LT). This supports the thought that in poultry, there is less collagen per volume in breast
muscles than in the leg muscles. As the general appearance of chicken breast meat is very
soft and turkey meat can be peeled in strips, it is possible, but not proven here that the low
collagen content predisposes chicken breast meat to soft and turkey breast meat to
disintegrating structure. If this is true, the phenomenon has slowly developed during the
last decades of intensive breeding of poultry. A comparison between a commercial breed
and a less selected breed might provide information on that in the future.
65
6.4.2 Collagen solubility of the poultry muscles
As discussed in the literature review (page 40), the meat producing animals are
slaughtered well before puberty. This is the most probable explanation for the very high
collagen solubility in the chicken muscles obtained in the present study, as the chickens
are the youngest, not only in days, but also in their maturity, at the time of slaughter when
compared to pigs and turkeys. Even turkeys are younger in physiological age than pigs at
the time of slaughter. Therefore the collagen solubility of the turkey muscles might have
been expected to be higher than that of the pig muscles but in the present study collagen
solubility of the turkey muscles was at the same level with the porcine muscles (Figure
12).
6.4.3 Collagen solubility with animal age and anatomical location
The pigs, turkeys and chickens in the present study were of commercial age so the age of
the animals could vary several days especially in pigs. However, as the carcass weights
were recorded in the studies I and III together with collagen solubility, it was possible to
plot the collagen solubility against the carcass weight (Figure 12). The Figure 12 shows
similarities with the results of Hill (1966), Nakano and Thompson (1980) and Cross et al.
(1984) (Figure 13).
Figure 12. Collagen solubility in chicken breast muscles (n=12, study III), turkey breast
muscles (n=12, study III) and porcine SM muscles (n=27, study I) in relation to carcass
weight.
The proportions of the body change during growth. In newborns the head is big and
well developed relative to the rest of the body, because the brain co-ordinates the body
66
activities. Also, the gut is already well developed in newborns as it is responsible for
digesting food for energy source for the body functions and growth. After birth the wave
of growth spreads from head backwards along the trunk. Secondary waves of growth start
from low in the limbs and move up to meet the first wave in loin. For pigs this means that
in the body of a piglet the lower part of the legs are relatively big, consisting mainly of
bone. During growth, muscles of back and thighs develop so that upper part of the legs
become round and the loin region develops (Hammond et al., 1971). However, Charles
(1986) showed that in cattle the proportion of the carcass muscle weight decreased in the
hind quarter muscles and increased in the fore quarter muscles between the age of 9 and
15 months.
Figure 13. Collagen solubility according to different authors in different bovine muscles.
It could be speculated that the state of growth in different porcine muscles is likely to
be different when measured for the same animal at a given time. For collagen solubility
this would mean that different muscles contain different proportions of newly synthesised
collagen, which according to Bailey and Light (1989) is more soluble than mature
collagen. When considering only this, the results from the studies I would indicate that the
growth of the pigs at the commercial slaughter age would be most intensive in shoulder as
the IS muscles in pigs had the highest collagen solubilities. According to Mahon (1999),
in poultry, leg muscles grow fast initially, but then the breast muscles rapidly grow and
mature actually later than the leg muscles. Still the poultry leg muscles had slightly higher
collagen solubility than the breast muscles. The hindquarters of the pig carcasses and
breast muscles of chickens and turkeys were well developed. As the muscles, that do not
produce destructured (pigs, turkeys) or soft (chicken) meat, had higher collagen solubility
than the muscles that produce destructured or soft meat, it seemed that high collagen
67
solubility could not be directly linked to the destructuration in meat. However, as it seems
that collagen solubility in porcine muscles has gradually increased (page 62) and the
collagen solubility of chicken muscles is already high, it is possible that the increased
collagen solubility might predispose the cooked meat products to the disintegrating
appearance.
The amount of insoluble collagen holding the structure of cooked meat is very low in
chicken and turkey PS muscles (1.7 and 2.0 mg/g, respectively, calculations made from
the Table 4) in comparison to the leg muscles (2.7 and 3.3 mg/g, respectively). Also the
porcine LD and SM muscles have less insoluble collagen than the IS muscles (2.4 and 3.6
mg/g vs. 4.0 mg/g). It could be speculated whether the amount of insoluble collagen in
porcine SM and LD muscles and in poultry PS muscles is sufficient to hold the cooked
meat cohesiveness.
Zimmerman et al. (1993) showed that training prevented the accumulation of mature
collagen cross-links (HP) in dark coloured rat muscles. As the IS muscles of the pigs and
the leg muscles of the chickens and turkeys are dark(er) coloured muscles (than the LD,
SM and PS muscles) and work more constantly than the LD, SM and PS muscles, it seems
likely that the collagen solubilities found in the present study were more connected to
anatomical location and work load of the muscles than directly to the destructured meat.
However, it should be remembered that the relationship between collagen solubility and
collagen cross-link content in muscles is not clear as discussed in the literature review (p.
39).
6.4.4 Thermal shrinkage temperatures of intramuscular connective tissue in
the poultry muscles in relation to ultimate pH
In contrast to porcine samples, the only correlation, a positive one, between the Tp of the
IMCT and pHu was in the chicken breast muscles (Appendix, Figure G). However, the pHu
range in the muscles of different animal species differed: In the study III, the mean pHu of
the chicken and turkey breast muscles was 5.86-5.96 and the mean pHu of the leg muscles
was 6.32-6.62 whereas the mean pHu of the porcine SM muscles varied from 5.42 to 5.57
(Table 4) and the mean pHu of the IS muscles was 5.94-6.25 (I). As according to Puolanne
& Ruusunen (1981) the amount of bound water in pig skin did not change dramatically at
the pH range 6-8, the To and Tp of the IMCT, which according to Miles et al. (2005) would
be dependent on the state of hydration of collagen, should have not been expected to
change either in the study III. This was also supported by the finding that none of the
breast and leg muscles of the chickens and turkeys exhibited exceptional structure when
inspected during sampling (III).
The variation of meat pHu seems to be greater in poultry than in red meat species (for
example Koohmaraie et al., 1991, Wilkins et al., 2000). Also in the present work the
variation in pHu was greater in poultry meat than in pork. As discussed in study III,
different breeds and fasting time of the birds are potential sources of variation. One factor
could be also the duration and extent of the stress the birds experience during catching,
transportation and slaughter. However, for study III, these were not monitored.
68
6.4.5 DSC sample moisture and enthalpy of thermal shrinkage in poultry
The DSC sample moisture in the chicken and turkey samples (III) was not exceptional in
comparison to the porcine samples. Interestingly, in the turkey breast muscles the DSC
sample moisture correlated negatively (p<0.01) with the To but not with the Tp of the
IMCT. In the chicken leg muscles in turn, the DSC sample moisture correlated negatively
with the To and also with the Tp of the IMCT.
Interestingly, the enthalpy (ΔH) of thermal shrinkage (the energy needed to transform
the collagen from a crystalline to an amorphous structure) of the IMCT was higher in the
chicken muscles than in turkey muscles (III). Assuming that the difference in the ΔH
between chickens and turkeys was due to the cross-links of IMCT or collagen, collagen
solubility might have correlated with the ΔH, but this was not found. It proved that the
energy was spent on the transformation discussed above. The numbers showed that the
chickens had slightly more intramuscular collagen than the turkeys although the difference
was not significant. This favours the thought that the relative proportions of the collagen
cross-links were different in the chickens and turkeys, which would be logical as the
turkeys were more mature than the chickens when slaughtered.
Interestingly, the ΔH of the poultry dark coloured leg muscles did not differ from that
of the breast muscles. In contrast, the porcine dark coloured IS muscles had clearly higher
ΔH than the light coloured LD and SM muscles (I). The different proportions of crosslinks in collagen could be due to the functional differences of the dark and light coloured
muscles. However, these were not investigated.
The enthalpies of denaturation of IMCT, either J/g or J/g d.m., in the studies I-IV were
clearly lower than those obtained by Miles et al. (1995), Usha and Ramasami (1999) and
Miles et al. (2005) from rat tail tendon. This was not surprising: according to Bailey and
Light (1989) the collagen content of tendon is 95 % per dry weight and that of muscle 2 %
per dry weight. Of course, most of the myofibrillar proteins were extracted before DSC
analyses in the present work, and, undoubtly the samples contained more than 2 %
collagen per dry weight, but as was shown in study II (Figure 2) the samples contained
also other proteins.
6.4.6 The possible role of non-collagenous molecules of intramuscular
connective tissue in the DSC sample moisture and weakened structure
The DSC samples included also other proteins than collagen (II, Figure 2) and it is known
that proteoglycans, especially the GAGs that are bound to the core protein to form the
complex of proteoglycan, provide the hydrated space for other molecules also in
connective tissue (Ruoslahti, 1988). According to Bailey and Light (1989) the role of
hyaluronate, the largest glycosaminoglycan, is to hold water in the tissue. This might also
be in association with the state of hydration of collagen.
Bailey and Light (1989) also pointed out that the shrinkage temperature of tendon
collagen was reported to be linearly dependent on the chondroitin sulphate content of
connective tissue. It is, thus, possible for the destructured SM muscles, that during the
69
formation of the destructured meat the properties of proteoglycans changed so that the
water content in the IMCT of the destructured SM muscles became higher than in the
IMCT of normal SM muscles. As Usha and Ramasami (1999) concluded that extensive
hydration could lead to volume changes in matrix and initiate rupturing, it is possible, that
the change in the properties of proteoglycans changed the hydration of the IMCT so much
that it caused the destructured appearance which was seen as the increased DSC sample
moisture and the decreased To and Tp of the IMCT. In fact, Ignatieva et al. (2004) showed
that the type II collagen in cartilage was stabilised from denaturation by proteoglycans.
They concluded that at least in cartilage the proteoglycans act as thermostabilisers of
collagen fibrils.
6.5 Muscle fibre properties in pork and poultry
6.5.1 Muscle fibre CSA
The main purpose of studying the muscle fibre properties (III, IV) was to see if large
muscle fibres had lowered thermal stability of the IMCT and were more prone to the
destructured appearance. As already reported by Swatland (1990) raw turkey breast meat
has a destructured appearance and cooked turkey breast meat disintegrates, and chicken
breast meat has been found to be generally very soft (Allen et al., 1998; Makkonen, 2003)
and can even be mushy, the relationship between the muscle fibre CSA, the thermal
stability of IMCT was studied both in pork and poultry.
According to Ruusunen and Puolanne (1997) the fibre CSA of LD muscles was 4.2 x
3
10 μm2 and the fibre CSA of M. adductor (ADD muscle) was 4.8 x 103 μm2. In the studies
of Essen-Gustavsson et al. (1992), Cerisuelo et al. (2007) and Sorensen et al. (1996) the
fibre CSA of LD muscles varied from 3.6 x 103 μm2 to 7.5 x 103 μm2 depending on the pig
breed, halothane genotype of the pigs and porcine growth hormone administration to the
pigs. Ruusunen and Puolanne (1997, 2004) and Sorensen et al. (1996) measured the fibre
CSA after myosin ATPase staining whereas Essen-Gustavsson et al. (1992) measured the
fibre CSA after amylase-PAS staining and Cerisuelo et al. (2007) recorded the fibre CSA
straight from the frozen section.
The reports of destructured pork have concentrated on the SM muscles and to a lesser
extent on the LD muscles (Franck et al. 2002). In the study of Ruusunen and Puolanne
(2004) the decreasing order of muscles in fibre CSA was SM (5.7 x x 103 μm2) > M.
gluteus superficialis (GS, 5.3 x 103 μm2) > LD (5.3 x 103 μm2) > IS (5.1 x 103 μm2) >
MAS (2.8 x 103 μm2), thus, the SM muscles had the thickest fibres of the studied muscles.
Swatland also (1990) pointed out that the muscle fibres were thicker in the destructured
turkey breast meat than in the normal turkey breast meat. It is therefore possible, that the
thickness of the muscle fibres in the SM muscles could predispose the meat to the
destructured appearance, but this was not proven in the present study as the
destructuration was detected around the muscle fibre bundles.
70
Many different diseases have been reported in poultry muscles: poultry leg weakness
has been associated with focal degenerative changes in the muscles of turkeys. The
heritable muscular dystrophy caused by abnormality of single autosomal recessive gene
was described in the 1960's. Yet there is a degenerative myopathy causing green colour
and dehydrated wood-like texture of meat. This occurs only in deep pectoral muscles
(Sosnicki et al. 1991b), which is why it does not fit in the picture of destructured turkey
meat and soft chicken meat, as also the superficial breast muscle is destructured/soft.
Focal myopathy (for example Mahon, 1999) would be a candidate for the defect, but no
exceptional muscle fibres (necrosis, degenerative changes) were found when determining
the muscle fibre CSA. A stress induced myopathy (Mitchell, 1999) could be a candidate
for the destructured and softened structure of turkey and chicken meat, respectively, but
this was not studied. However, none of the poultry muscle samples for the present study
exhibited exceptional structure or colour although the breast muscle fibre CSA in turkeys
varied from 3.7 x 103 μm2 to 6.5 x 103 μm2 and in chickens from 2.9 x 103 μm2 to 4.3 x 103
μm2.
6.5.2 Could the destructured appearance of meat be associated with
weakness of single muscle fibres?
It has been shown that in cooked muscles the fracture initiates in perimysium, when meat
is pulled perpendicular to the muscle fibre direction (Purslow, 1985). For disintegrating
cooked porcine or turkey meat products this has not been proven, but the strip cut by knife
(Figure 1A) could not be pulled in normal raw meat, as the strip broke, thus a perimysial
defect in the destructured meat seems likely. Lewis et al. (1991) also showed that the
perimysium from meat, that had been cold stored (14 days at 1 °C) to soften the meat
structure, had lower breaking strength than the perimysium from untreated meat.
At the single muscle fibre level, Christensen et al. (2004) showed that cold storage (10
days at 2 °C) reduced the tensile strength of bovine muscle fibres. For porcine LD
muscles, it has been shown that the type IIB muscle fibres are weaker per cross sectional
area than the type I muscle fibres (Christensen et al., 2006). According to Ruusunen and
Puolanne (2004) the SM muscles contained almost 90 % (in % area) of the IIB type
muscle fibres. From these a possible conclusion could be that the destructured appearance
could be seen also at single muscle fibre level as it is known that proteolysis post mortem
occurs faster in pork than in beef (Koohmaraie et al., 1991) and that the destructured
appearance is usually detected in the boning hall at least 24 h post mortem.
From the study of Oksbjerg et al. (2000) it could be seen that fast growing (FG) pigs
had almost as thick fibres in LD muscles as the slow growing (SG) pigs (fibre CSA 5.9 x
103 μm2 vs. 4.9 x 103 μm2, respectively), although the FG pigs needed 25 days less time to
grow from 40 kg to 98 kg than the SG pigs when slaughtered approximately at the same
weight. Oksbjerg et al. (2000) suspected that the younger age of the FG pigs at slaughter
in comparison to the SG pigs caused the slight softening of the IMCT when interpreted
from Warner-Bratzler shear force measurements. This conclusion would support the
thought that nowadays the thick fibred muscles of young fast growing pigs are prone to
71
the destructured appearance and that also the connective tissue is affected in the
destructured meat.
Indeed, Swatland (1990) reported that the muscle fibres from disintegrating turkey
meat were almost twice as thick as the muscle fibres from normal muscles. However, as
the destructured meat investigated in the present study was detected at perimysial level
(Figure 1A) and the muscle fibre properties were similar in the destructured and normal
SM muscles, the weakness of the single fibres in the SM muscles could not be directly
associated with the destructured raw SM muscles.
Lawson (2004) found that in porcine LD muscles with high drip loss integrin (a protein
attaching the membranes to the actin cytoskeleton, discussed in the literature review p.
31), more specifically the β1-subunit of it, could be seen to be degraded as early as 3 hours
post mortem which was before the rigor mortis took place in the muscle. In Lawson's
(2004) opinion however, high early post mortem temperature did not cause rapid
degradation of integrins although high temperature together with low pH early post
mortem have been associated with high drip loss. Therefore the integrins and other linkage
proteins between the endomysium and the muscle fibre might be of interest in association
with the destructured meat with high drip loss.
Lawson (2004) also concluded that the rapid pH fall alone without integrin degradation
could not directly lead to drip channel formation, but according to her hypothesis, early
calpain mediated integrin degradation and drip channel formation favoured the early and
high drip loss formation. Lawson's (2004) hypothesis was supported by the findings of
Zhang et al (2006) who reported that a high amount of detectable intact integrin was
associated with low drip loss and with low purge loss. However, Straadt et al. (2008) did
not find a correlation between the intensity of intact integrin in Western blot figure and
drip loss. For the destructured pork the role of integrins would mean that something
actually happens to the integrins at the single muscle fibre level during formation of the
destructured appearance in raw meat. This still remains unknown.
6.5.3 Sarcomere length in destructured SM muscles
Sarcomeres were very short in both the destructured and normal SM muscles (IV).
However, as there was a tendency of the destructured SM muscles to have longer
sarcomeres (IV, Table 2, p<0.10) than the normal muscles, it is possible that the
destructured SM muscles had experienced accelerated glycogenolysis. As the destructured
SM muscles have been concluded to be of PSE origin (Le Roy et al., 2001; Franck et al.,
2000; Minvielle et al., 2001; Laville et al., 2005) the accelereated glycogenolysis would
fit in the theory. In the present study it was not feasible to obtain the early post mortem pH
values as the destructured meat is seen in deep SM muscles not earlier than in the boning
hall. However, Honikel and Kim (1985) concluded that long sarcomeres in PSE muscles
are caused by the early decrease of the ability of sarcomeres to contract due to the
decrease of the activity of myosin ATPase.
72
6.6 Concluding remarks
The animals farmed for meat production are younger than before as a result of intensive
breeding for fast growth. This may have led to the situation that fast growing animals,
although stress resistant genotypes and handled humanely, are more prone to muscular
defects. From PSE paleness and exudation are easy to measure, but softness is still the
least studied part. The data from study II point to the direction that the collagen content or
collagen solubility are not different between destructured and normal structured pork.
However, the general level of collagen content is so low and the animals slaughtered are
so young that the low amount of immature collagen might predispose the meat to
structural defect, although meat from a bit more mature animals with slightly higher
collagen content would not exhibit destructuration after similar slaughter conditions. High
collagen solubility does not seem to be the key, because the muscles that exhibit
destructuration have lower collagen solubility than the muscles that do not exhibit
destructuration. However, the amount of insoluble collagen holding the structure also in
cooked meat procucts is generally very low in the muscles exhibiting destructuration. The
data from study IV in turn, show that the muscle fibres in destructured and normal
structured pork are equally large in fibre CSA. However, it can not be a coincidence that
the destructuration is reported most in SM and LD muscles (light coloured muscles) in
which the glycolytic fibres are dominant and the muscle fibres are among the largest ones
in the pig body. In the literature it has been shown that perimysial connective tissue is
least developed in light coloured muscles. In addition, the defective areas are deep in the
carcass and chill very late after slaughter. Thus, for pork, destructuration seems a
connective tissue defect caused by a local PSE type condition.
Poultry breast muscles have low collagen content, and they consist of only glycolytic
muscle fibres. Chicken muscles have also high collagen solubility. Similarly to pigs,
softness in chicken and destructuration in turkey occur in the muscles bred to grow fast
and big. As according to the author’s opinion all the chicken breast meat provided
nowadays is soft and most of the turkey breast meat is destructured, the baseline type data
about the current situation of the IMCT of poultry were obtained.
However, the final explanation to why the strip of muscle can be pulled by hand in the
raw destructured meat but not in the normal meat has not been presented. It was not
possible either in the present study. The association between destructured meat and pale,
soft, exudative meat and the role of IMCT in both phenomena remain still problematic.
73
7 Conclusions
Collagen content of light coloured muscles (M. longissimus dorsi, LD muscle and M.
semimembranosus, SM muscle in pigs, M. pectoralis superficialis PS muscle in poultry) is
lower than that of dark coloured muscles (M. infraspinatus, IS muscle in pigs,
combinations of M. quadriceps femoris and M. iliotibialis lateralis, leg muscles in
poultry). As the light coloured muscles may exhibit structural defects, it is possible that
the connective tissue system is not sufficient to hold the meat structure.
Collagen solubility is higher in dark coloured muscles than in light coloured muscles,
thus it can not be directly associated with the destructured raw meat or disintegrating
cooked meat. However, the amount of insoluble collagen is still lower in light coloured
muscles than in dark coloured muscles which might be associated with the structural
defects in cooked meat. Collagen solubility is especially high in chicken muscles, which
might predispose the cooked chicken meat to mushy structure.
In pigs, the onset and peak temperature of thermal shrinkage (To and Tp) of
intramuscular connective tissue (IMCT) in dark coloured muscles are higher than those of
light coloured muscles. Similar difference does not exist between the To and Tp of IMCT in
dark and light coloured poultry muscles. This may be explained by the higher range of
meat ultimate pH (pHu) in poultry muscles than in porcine muscles and supports the
thought that low pHu predisposes the meat to destructuration.
The muscle fibre properties are not associated with the properties of IMCT in pigs and
poultry. Destructured meat is similar to normal meat in collagen content and solubility,
muscle fibre cross sectional area, capillary density, sarcomere length and extracellular
space. However, destructured meat has lower pHu and higher drip loss than normal meat.
The IMCT of destructured meat, with low pHu, has lower To and Tp than normal meat with
high pHu. Thus, in addition to pale colour, which was a criterion of selection, and drip
loss, the low To and Tp indicate destructuration of meat.
The To and Tp of poultry IMCT were similar to those of porcine IMCT. Thus, the
animal species and animal maturity does not seem to affect the thermal shrinkage
temperature of IMCT.
74
8 Suggestions for further studies
A comparison of the content of non-collagenous molecules in the IMCT of the
destructured and normal SM muscles might be the easiest procedure to continue the
research of the present topic, as the DSC results of the present indicated that something
changed in the IMCT of the destructured meat during cold storage. Another approach
could be the investigation of enzymatic activity towards the IMCT.
As it seems that the destructured appearance is a defect in perimysium a detailed,
possibly a 3D-microscopic study following the changes in the properties of the
perimysium (SM muscle) during the first hours and days post mortem could provide
additional information on the mechanisms associated with the development of the
destructuration found in raw meat. The detection of a change in non-collagenous
molecules and enzymatic activity in the destructured SM muscles in comparison to normal
muscles could provide preliminary information for the 3D-microscopy as only few types
of molecules can be labelled per section to be examined for example with the confocal
laser scanning microscopy.
Lawson (2004) showed that m-calpain co-localised with the β1 subunit of integrin on
the cell surface and μ-calpain was spread throughout the muscle cells. It might be
therefore of interest to study whether changes in the activities of calpains can be
associated with the formation of the destructured meat. However, one should remember,
that even though a lot of information on integrins and calpains are available, modelling the
in vitro to in vivo and eventually to post mortem conditions should be done carefully.
All these approaches would be most informative if the destructured meat could be
detected during the early hours post mortem and followed until, for example, 4 days post
mortem. However, the early detection of the destructured meat seems possible only if it is
assumed to be closely related to the PSE phenomenon, as it is known that excess stress of
pigs prior to slaughter induces PSE in pork and these pigs could be also selected prior to
slaughter. Causing stress to pigs would require permission from the ethical committee of
animal testing.
75
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Appendix A. Figures of combined data
Figure A. Relationship between the pHu and L* (72 h post mortem) value in the combined
data of studies II and IV, when both destructured and normal muscles are included.
Figure B. The relationship between pHu and drip loss when a combined data from studies
II and IV are presented.
90
Figure C. Relationship between the pHu and DSC sample moisture in the combined data of
studies II and IV, when both destructured and normal muscles are included.
Figure D. Relationship between To (the onset temperature of thermal shrinkage of
intramuscular connective tissue) and DSC sample moisture in the combined data of
studies II and IV, when both destructured and normal muscles are included.
91
Figure E. Relationship between Tp (the peak temperature of thermal shrinkage of
intramuscular connective tissue) and DSC sample moisture in the combined data of
studies II and IV, when both destructured and normal muscles are included.
Figure F. Relationship between Tp and pHu in studies II and IV.
92
Figure G. Relationship between Tp and pHu in study III, when only breast muscles of
chickens and turkeys are included.
Figure H. Relationship between collagen solubility and To (the onset temperature of
thermal shrinkage of intramuscular connective tissue) in the combined data on SM
muscles in studies I (baseline) and II (destructured and normal).
93
Table A. The Colour readings (mean ± s.d.) of the destructured and control muscles at 96
h post mortem in study IV.
Destructured (n=9)
Control (n=25)
L* 96 h
53.2±2.1
50.5±2.1
a* 96 h
10.6±1.5
9.4±1.4
b* 96 h
9.9±0.8
8.2±1.6
94