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First published online February 12, 2007
Journal of Experimental Biology 210, 825-835 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.02711
Getting the jump on skeletal muscle disuse atrophy: preservation of contractile performance in aestivating Cyclorana alboguttata (Günther 1867)
1 School of Integrative Biology, The University of Queensland, St Lucia,
Queensland 4072, Australia
2 Department of Physiology and Sport Science, Coventry University, James
Starley Building, Priory Street, Coventry, CV1 5FB, UK
* Author for correspondence (e-mail: c.franklin{at}uq.edu.au)
Accepted 9 January 2007
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Summary |
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Key words: aestivation, muscle disuse atrophy, fibre, morphology, contractile properties, locomotion, work loop, Cyclorana alboguttata
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Introduction |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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). An
increase in muscle activity builds and maintains muscle mass, while inactivity
results in tissue degeneration. In mammals, prolonged immobilisation or
unloading of skeletal muscle results in muscle `wasting' or muscle disuse
atrophy (Booth, 1982). Muscle
disuse atrophy is characterised by a reduction in muscle fibre cross-sectional
area due to the combined effects of increased catabolism and decreased
anabolism of skeletal muscle protein and an imbalance of muscle biochemistry
(Kandarian and Stevenson,
2002).
In experimental models of hindlimb immobilisation and unloading the effects
of muscle disuse atrophy on muscle morphology and muscle mechanics are
variable and depend upon muscle type and fibre type
(Ariano et al., 1973;
Booth and Kelso, 1973;
Edgerton et al., 2002;
Peter et al., 1972;
Trappe et al., 2004;
Witzmann et al., 1982). Most
muscles are composed of a heterogenous mixture of fibre types, although there
are exceptions. For example, the mammalian soleus muscle is almost completely
slow twitch (Ariano et al.,
1973).
During normal activity, oxidative muscles tend to be used frequently, for
extended periods of time and at low intensities. In comparison, fast-twitch
glycolytic muscles are used infrequently, for short periods and at high
intensity. When oxidative muscles become inactive the scope of the change in
the level of activity is much greater than that experienced by glycolytic,
fast-twitch muscles (Hudson and Franklin,
2002b). In general, slow-twitch (oxidative) fibres are more
vulnerable to muscle disuse atrophy than fast-twitch (glycolytic) fibres
(Edgerton et al., 2002;
Maier et al., 1973;
Nordstrom et al., 1995;
Tomanek and Lund, 1974).
Prolonged disuse alters muscle fibre type characteristics, with oxidative
fibres acquiring the structural, biochemical and mechanical properties of
glycolytic fibres (Anderson et al.,
1999; Diffee et al.,
1991). Generally, tetanic stress is reduced, maximum rate of
shortening is elevated, and fatigue resistance is slightly reduced or
unaffected (Booth and Seider,
1979a; Roy et al.,
1991; Roy et al.,
1996; Roy et al.,
2002). Commonly, the contractile rates of oxidative muscles are
increased after immobilisation leading to quicker twitch times that are more
similar to glycolytic muscles (Anderson et
al., 1999; Canon and Goubel,
1995; Witzmann et al.,
1982). Additionally, due to the comparatively greater numbers of
mitochondria contained within them, oxidative fibres produce comparatively
larger quantities of reactive oxygen species as a byproduct of normal cell
metabolism. Reactive oxygen species, or free radicals, are responsible for
stochastically damaging cell architecture, including proteins and lipids,
which leads to cellular apoptosis and hence muscle disuse atrophy
(Kondo et al., 1993a;
Kondo et al., 1993b;
Kondo et al., 1993c;
Kondo et al., 1994). During
disuse the dramatic downregulation of activity in the oxidative muscle may
trigger profound changes in the biochemical function of the muscle
mitochondria.
Few studies have explored the effect of dormancy on muscle disuse atrophy
of skeletal muscle (for a review, see
Carey et al., 2003). Of the
available literature, the majority investigate mammalian models, but virtually
none have been conducted on amphibians. In general, the animals studied
exhibit some muscle disuse atrophy, though to a lesser degree than observed in
humans under similar conditions of immobility
(Harlow et al., 2001;
Tinker et al., 1998). For
example, hibernation for 6 months caused significant atrophy in skeletal
muscle from the golden-mantled ground squirrel (Spermophilus
lateralis) as measured by whole muscle mass
(Wickler et al., 1991b) and
whole muscle cross-sectional area, with observable concurrent, but
non-significant, declines in the protein content of these muscles
(Steffen et al., 1991).
Animals that experience regular bouts of dormancy, such as hibernators and
aestivators, exhibit a decline in metabolic and locomotor activity that is
correlated with seasonal fluctuations in temperature and resource availability
(Guppy and Withers, 1999).
During dormancy the animal is primarily reliant upon stored lipids, with
protein meeting additional energy demands
(Harlow et al., 2001;
Pakay et al., 2002;
Pedler et al., 1996;
Seymour, 1973;
Tinker et al., 1998;
van Beurden, 1980). Muscle
comprises the largest protein store in the body, therefore catabolism of
muscle tissue as an energy substrate during hibernation and aestivation may
contribute to muscle disuse atrophy
(Steffen et al., 1991;
Wickler et al., 1987;
Wickler et al., 1991b;
Yacoe, 1983). Additionally, it
has been demonstrated that protein synthesis is downregulated during dormancy,
which may further compound the atrophic effects of dormancy on skeletal muscle
(Frerichs et al., 1998;
Fuery et al., 1998;
Pakay et al., 2002). As such,
hibernators and aestivators provide a fascinating model for investigating
muscle atrophy associated with disuse (see review by
Hudson and Franklin,
2002b).
Amphibian aestivation usually occurs in response to unfavourable
environmental conditions (Kobelt and
Linsenmair, 1995), specifically a dry and warm (>10°C)
environment, and is characterised by the construction of a thermally stable
burrow (van Beurden, 1980),
the formation of a cocoon (Withers,
1995; Withers,
1998) and metabolic rate depression
(Flanigan et al., 1991;
Pinder et al., 1992). The
green-striped burrowing frog Cyclorana alboguttata (Günther,
1867) (Meyer et al., 1997) is
an amphibian aestivator that inhabits the arid and semi-arid zones of Eastern
Australia. This species undergoes aestivation for many months (6–9
months on average) between seasonal rain events
(Withers and Richards, 1995).
After 3 months of aestivation locomotor performance, wet muscle mass of
several hindlimb muscles and contractile properties of the gastrocnemius of
C. alboguttata remained unchanged from that of control animals,
indicating an absence of muscle disuse atrophy
(Hudson and Franklin, 2002a).
Long bone mechanical properties, capillary structure and gut function were
also preserved during aestivation in C. alboguttata
(Cramp and Franklin, 2005;
Hudson et al., 2004;
Hudson and Franklin,
2003).
It is clear that although C. alboguttata does not exhibit typical
atrophic responses, we are still unsure as to what exemplifies a typical
aestivator response to prolonged muscle disuse. The aim of the present study
was to examine the effect of prolonged aestivation on the morphology and
contractile performance of whole muscles from C. alboguttata. Unlike
previous studies, this project investigated the effect of aestivation over a
dramatically longer, more ecologically relevant, time-frame of 9 months. At
the time of publication, no other study had examined the effect of aestivation
or hibernation on any aspect of muscle physiology or biochemistry over such an
extended period. Though Hudson and Franklin's study
(Hudson and Franklin, 2002b)
demonstrated that C. alboguttata did not exhibit muscle disuse
atrophy after 3 months of aestivation, we felt that the results of that study
were not indicative of the true capabilities of this animal as a model
species.
We hypothesised that aestivation of 9 months duration would result in changes to the fibre-type composition of the sartorius (relatively fast-twitch, glycolytic) and iliofibularis (relatively slow-twitch, oxidative) muscles. Specifically, the iliofibularis muscle would exhibit a compositional change from the atrophy-susceptible oxidative fibre type to the more atrophy-resistant glycolytic fibre type. If evident, we proposed these changes in fibre type would affect muscle contractile performance of aestivating frogs; the slow-twitch iliofibularis muscle would exhibit the contractile properties of a fast-twitch muscle, and the fast-twitch sartorius would become faster still. We measured the effect of aestivation on whole muscle cross-sectional area, muscle fibre type, fibre number and fibre cross-sectional area, and related the observed ultrastructural changes (if any) to muscle mechanical performance as measured by isometric twitch and tetanus kinetics, work-loop power output and work-loop fatigue resistance.
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Dissection
The skin of both hind limbs was removed to expose the muscle tissue. Slices
of approximately 5 mm thickness were dissected from the belly of the
right-side iliofibularis and left-side sartorius muscles with a razor blade.
The iliofibularis and sartorius were chosen to represent relatively slow and
fast skeletal muscles, respectively, based upon previous studies of anuran
muscle fibre type (Putnam and Bennett,
1983) and preliminary findings by the author. These fresh muscle
slices were placed into plastic moulds, mounted in Tissue-TekTM OCT
compound (ProSciTech, Townsville, Australia) and plunged into isopentane
(2-methylbutane), cooled to –150°C in liquid nitrogen, for
approximately 30 s. The frozen blocks were removed and wrapped in aluminium
foil to prevent desiccation, and stored at –80°C in an air-tight
container for histochemical analysis. The sartorius muscle from the right leg
and the iliofibularis from the left leg of control and treatment frogs were
removed, leaving a section of bone attached to the tendon at either end of
each muscle. These muscles were stored in refrigerated McKenzies frog Ringer
solution (111 mmol l–1 NaCl, 2.5 mmol l–1
KCl, 1.8 mmol l–1 CaCl2. 2H2O, 1 mmol
l–1 MgCl, 5 mmol l–1 Hepes, 10 mmol
l–1 glucose, adjusted to pH 7.4 with NaOH) until required for
isometric analysis.
Histochemistry
Whole muscle and individual fibre morphometrics were determined by
histochemical analysis. Frozen muscle blocks were mounted onto stainless steel
chucks with OCT compound, and placed in a cryostat chamber (HM505, Microm,
Walldorf, Germany) at –20°C. Frozen cross sections (10 µm
thickness) were cut and melted directly onto glass slides and air-dried for
approximately 3 min. The sections were returned to the cryostat chamber for
storage in an air-tight container prior to staining. Best results were
achieved for muscle that was sectioned and stained on the same day. Slides
were removed from the cryostat chamber and stained immediately for succinic
dehydrogenase (SDH) activity to differentiate between oxidative and glycolytic
muscle fibres: 50 mmol l–1 phosphate buffer, 80 mmol
l–1 sodium succinate, 1 mg ml–1 Nitro blue
tetrazolium, adjusted to pH 7.6 with NaOH
(Rowlerson and Spurway, 1988).
Sections were incubated in the staining medium in air-tight, light-proof
containers for at least 90 min. Each slide was washed three times in distilled
water, air dried and mounted with Crystal/MountTM (Biomeda Corp.,
Burlingame, CA, USA). Each section was photographed at a 250x
magnification with a digital camera (DFC280, Leica, Wetzlar, Germany) mounted
to a compound light microscope (BH2, Olympus, LeHigh Valley, PA, USA). Digital
images were analysed with SigmaScanTM (SPSS Inc., Chicago, IL, USA) to
determine muscle cross-sectional area, fibre area, fibre number and fibre
density of each whole muscle section.
Isometric analysis
Isometric analysis was used to determine the twitch and tetanus kinetics of
the isolated hindlimb muscles. The bone at either end of the muscle
preparation was clamped via crocodile clips to a strain gauge (UF1,
Pioden Controls Ltd, Canterbury, UK) at one end, and a motor arm (V201, Ling
Dynamics Systems, Royston, UK) attached to an LVDT (Linear Variable
Displacement Transformer, DFG 5.0, Solartron Metrology, Bognor Regis, UK) at
the other. The LVDT was used to monitor the length changes delivered to the
muscle preparation. The whole of the muscle, tendon and bone preparation was
then maintained within the organ bath at 25±0.5°C in circulating,
oxygenated McKenzies frog Ringer solution. The preparation was held at
constant length and stimulated via parallel platinum electrodes to
generate a series of twitches. Stimulus amplitude, pulse width and muscle
length were adjusted to determine the stimulation parameters and muscle length
corresponding to maximal isometric twitch force. Time to peak twitch and time
from peak twitch to half relaxation were measured via a Powerlab 4SP
(AD Instruments, Colorado Springs, CO, USA) and viewed with Chart 5.0 software
(AD Instruments). An isometric tetanic force response was elicited by
subjecting the muscle to a 300 ms train of stimulation. Stimulation frequency
(130–150 Hz for iliofibularis; 140–170 Hz for sartorius) was
altered to determine maximal tetanic force. Time to half peak tetanic force
and time from last stimulus to half tetanic force relaxation were measured. A
rest period of 5 min was allowed between each tetanic response.
Work loop analysis
The work-loop technique was used to determine the power output of muscles
during cyclical length changes (Josephson,
1993). Unlike fixed-length isometric studies and fixed-load
isotonic studies of muscle performance, the work-loop technique allows
measures of muscle power output under length and activation changes that are
generally more indicative of in vivo contractile performance
(Caiozzo, 2002;
James et al., 1996;
Josephson, 1993). Each muscle
preparation was subjected to a set of four sinusoidal length changes
symmetrical around the length found to generate maximal twitch force. The
muscle was stimulated using the stimulation amplitude and frequency found to
yield maximal isometric force. Electrical stimulation and length changes were
controlled via a data acquisition board and a custom-designed program
developed with TestPoint software (Measurement Computing, Norton, MA, USA).
Muscle force was plotted against muscle length for each cycle to generate a
work loop, the area of which equated to the net work produced by the muscle
during the cycle of length change
(Josephson, 1993). The net
work produced was multiplied by the frequency of length-change cycles to
calculate net power output. The total strain of length-change cycles was
maintained at 0.10 throughout all experiments, which equated to ± 5% of
resting muscle length. The cycle frequency of length change was altered up and
down within the range 1–9 Hz for iliofibularis and 2–10 Hz for
sartorius to generate power output-cycle frequency curves. During these length
change cycles the muscle was usually subjected to phasic stimulation (i.e.
active work-loop cycle) but sometimes these length changes were performed
without stimulation (passive work-loop cycle) to monitor the net work done on
the muscle during the length change cycle. Every 5 min the muscle was
subjected to a further set of four work-loop cycles with stimulation duration
and stimulation phase parameters being altered until maximum net work was
achieved at each cycle frequency. A set of control sinusoidal length change
and stimulation parameters were imposed on the muscle every 3–4 sets of
work loops to monitor variation in the muscle's ability to produce
power/force. Any variation in power was found to be due to a matching change
in ability to produce force. Therefore, the power produced by each
preparation, prior to the fatigue run, was corrected to the control run that
yielded the highest power output, assuming that alterations in power
generating ability were linear over time. All muscles still produced over 80%
of maximal control run power prior to the fatigue run. On completion of the
power output-cycle frequency curve each muscle was subjected to a fatigue run
consisting of 30 work-loop cycles at the lowest cycle frequency that had
earlier been found to produce 80% of maximal power output.
At the end of the isometric and work-loop experiments, the bones and
tendons were removed and each muscle was blotted on absorbent paper to remove
excess Ringer solution. Wet muscle mass was determined to the nearest 0.0001 g
using an electronic balance (BP211D, Sartorius). Mean muscle cross-sectional
area was calculated from muscle length and mass assuming a density of 1060 kg
m–3 (Mendez and Keys,
1960). Maximum isometric muscle stress was then calculated as
maximum tetanic force divided by mean cross-sectional area (kN
m–2). Normalised muscle power output was calculated as power
output divided by wet muscle mass (W kg–1). Each muscle was
dried for 48 h in an oven at 37°C, moved to room temperature for 1 h and
then reweighed to calculate dry muscle mass (to the nearest 0.0001 g) and
water content (% wet mass).
Statistical analysis
Frog length (mm) was not correlated with any of the parameters recorded
(Pearson correlation; P>0.05), therefore data sets were not
normalised to body size (SVL) prior to analysis. Dispersion
measurements are given as standard error. In all cases P=0.05 was
accepted for statistical significance.
Muscle morphometrics
Frog sizes and fibre size distributions were compared using a rank sum
t-test, while muscle morphological data was analysed by two-way ANOVA
using SigmaStatTM. Where differences were detected they were localised by
a Holm–Sidak multiple comparison test.
Muscle mechanics
In most cases control and experimental results were compared using
independent sample t-tests with correction for unequal variances
where appropriate. Control to treatment comparison of power output-cycle
frequency data was performed using ANCOVA, with treatment as the factor and
cycle frequency as the covariate. Control to treatment comparison of relative
power output data during the fatigue tests was performed using ANCOVA, with
treatment as the factor and cumulative power as the covariate.
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Muscle morphology
There was no significant difference in the total number of fibres present
in the muscle cross-sections, between either muscle type
(F1,19=2.863, P=0.110) or treatment group
(F1,19=2.557, P=0.129)
(Fig. 2A). Overall, there was
no significant difference in fibre density between muscle type
(F1,19=4.485, P=0.06) or treatment group
(F1,19=0.882, P=0.362)
(Fig. 2B), though there was a
significant interaction between muscle type and treatment group
(F1,19=11.53, P=0.004). A Holm–Sidak
multiple comparison test revealed that control sartorius fibre density was
significantly greater than 9-month aestivator sartorius fibre density
(P=0.007) and control iliofibularis density (P=0.001).
Control sartorius density was not more dense than 9-month aestivator
iliofibularis density, but only arbitrarily (P=0.050). This suggests
that, overall, sartorius muscle fibres were more densely packed within the
muscle. There was no significant difference in the relative proportions of
muscle fibre types (oxidative versus glycolytic) between muscle type
(F1,7=0.000175, P=0.992) or treatment group
(F1,7=0.000175, P=0.992)
(Table 1). Oxidative fibres
accounted for greater than 63% of the total number of fibres within each
muscle, with the control iliofibularis possessing the greatest proportion of
oxidative fibres (76%). Glycolytic fibres accounted for approximately 24% of
the total number of fibres within each muscle, the greatest proportion of
which was within the 9-month treatment sartorius muscle (37%).
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Fibre morphology
There was a significant difference in the CSA of individual fibres between
fibre type (F1,41=48.54, P<0.001) and
treatment group (F3,41=17.15, P<0.001). Within
the iliofibularis muscle, the CSA of oxidative fibres from 9-month aestivators
were significantly smaller than the control oxidative fibres
(P=0.041), and 9-month aestivator glycolytic fibres were
significantly smaller than control glycolytic fibres (P<0.001)
(Fig. 3A). There was no
significant difference between treatment groups for fibre CSA of the sartorius
muscle (P=0.221), though there was a significant difference between
fibre types: glycolytic fibres were significantly larger than oxidative fibres
in both controls (P=0.002) and 9-month aestivators (P=0.037)
(Fig. 3B). There was no
significant difference in the relative size distribution of fibres between
treatment group or muscle type (P>0.05), except for the
iliofibularis muscle. In this case, 9-month aestivator oxidative fibres were
skewed left, with significantly more fibres occurring in the smallest size
classes (P=0.045) (Fig.
4A).
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Work loop performance
Nine-months aestivation did not affect the peak power output produced by
either the iliofibularis or sartorius muscles
(Table 2). Power output-cycle
frequency relationships did not differ between control and treatment
iliofibularis muscles (Fig. 5A;
P=0.29). Aestivation caused a significant shift in the sartorius
muscle power output-cycle frequency curve
(Fig. 5B; P=0.004),
with treatment muscles producing lower power output at higher cycle
frequencies. Control sartorius muscles produced higher maximal power output at
a higher cycle frequency than the iliofibularis muscle, with the 9-month
aestivator sartorius muscle producing a power output-cycle frequency curve
similar to that of the iliofibularis. Aestivation tended to reduce fatigue
resistance in both the iliofibularis and the sartorius muscles, as evidenced
by a more rapid initial decline in 9-month aestivator muscle power output
during bouts of repeated work loop cycles
(Fig. 6A,B; P=0.53 and
P=0.09, respectively).
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Discussion |
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Muscle morphology
The sartorius and iliofibularis muscles of the 9-month aestivators
maintained dry muscle mass and whole muscle cross-sectional area at control
levels (Fig. 1A,B). Both
sartorius and iliofibularis act to abduct the femur and flex the knee
(Duellman and Trueb, 1994),
therefore these are not the main extensor muscles used to power jumping. As
such, they might represent skeletal muscles less likely to be protected from
atrophy during a challenge such as aestivation.
The absence of change at the whole muscle level is in stark contrast to the
effects of immobilisation or dormancy previously reported in other animals
(Adams et al., 2003;
Baldwin and Haddad, 2001;
Booth, 1977;
Booth and Seider, 1979b).
Similar significant reductions in muscle mass were observed in guinea pig
muscle after immobilisation for 28 days
(Maier et al., 1976).
Hibernation caused significant atrophy in the muscles of golden-mantled ground
squirrels Spermophilus lateralis
(Wickler et al., 1991a), big
brown bats Eptesiscus fuscus
(Yacoe, 1983) and hamsters
(Wickler et al., 1987).
However, aestivation for 3 and 6 months in C. alboguttata has
previously been shown to have no effect on the wet muscle masses of locomotory
muscles (Hudson and Franklin,
2002a; Hudson et al.,
2006).
There were no significant changes to total fibre number between control and
9-month aestivator groups for either muscle type in C. alboguttata
(Fig. 2A), which rules out loss
of fibres (hypoplasia) during aestivation. Fibre density declined
significantly after aestivation in the sartorius muscle only
(Fig. 2B). Combined with the
maintenance of sartorius dry muscle mass and fibre number, it is possible that
this reduction in fibre density reflects an increase in sartorius muscle
length during aestivation. Aestivating C. alboguttata adopt a
characteristic water-conserving posture
(Withers and Richards, 1995),
and it is possible that this behaviour might confer a postural advantage to
selected hindlimb muscles, such as the sartorius, by maintaining the muscle in
a lengthened position. It has been demonstrated in previous studies that
fixing muscle in the lengthened position during immobilisation can delay the
onset of muscle disuse atrophy (Booth,
1977; Goldspink,
1977). Our results are unlike previous studies in which the
cross-sectional areas of muscle fibres and numbers of fibres have
significantly decreased after immobilisation and dormancy. Immobilisation
caused significant reductions in the diameter of guinea pig soleus muscle
fibres after 21 days of immobilisation
(Tomanek and Lund, 1974), the
cross-sectional area of cat tibialis muscle
(Nordstrom et al., 1995) and
fibre number in rat soleus muscle (Booth
and Kelso, 1973). Hibernation caused a significant reduction in
the cross-sectional areas of fibres from the soleus and extensor digitorum
longus muscles of hibernating golden-mantled ground squirrels Spermophilus
lateralis (Steffen et al.,
1991) though not in hibernating black bears Ursus
americanus (Tinker et al.,
1998).
The most dramatic morphological change observed in this study was the
significant reduction in iliofibularis muscle fibre cross-sectional area
(Fig. 3A). Based on the results
of previous studies (e.g. Booth and Kelso,
1973; Maier et al.,
1976; Tomanek and Lund,
1974) we had predicted that the predominantly oxidative
iliofibularis muscle would be more susceptible to muscle disuse atrophy than
the glycolytic sartorius muscle. During normal activity, oxidative muscles
tend to be used frequently, for extended periods of time and at low
intensities. In comparison, fast-twitch glycolytic muscles are used
infrequently, for short periods and at high intensity. When oxidative muscles
become inactive the scope of the change in the level of activity is much
greater than that experienced by glycolytic, fast-twitch muscles
(Hudson and Franklin, 2002b).
Additionally, due to the comparatively greater numbers of mitochondria
contained within them, oxidative fibres produce comparatively larger
quantities of reactive oxygen species as a byproduct of normal cell
metabolism. Reactive oxygen species, or free radicals, are responsible for
stochastically damaging cell architecture, including proteins and lipids,
which leads to cellular apoptosis and hence muscle disuse atrophy
(Kondo et al., 1993a;
Kondo et al., 1993b;
Kondo et al., 1993c;
Kondo et al., 1994). The
dramatic downregulation of activity in the oxidative muscle may trigger
profound changes in the biochemical function of the muscle mitochondria. As
such, it is possible that the susceptibility of oxidative muscles to muscle
disuse atrophy is due to changes in mitochondrial density or architecture
rather than compositional fibre changes. In other words, the ultimate
consequences of muscle disuse atrophy may be more important at the whole
muscle level rather than at the individual fibre level.
The reduction in cross-sectional area of the iliofibularis muscle fibres, but not sartorius muscle fibres, supports our initial hypothesis; however, the results of the histochemical analysis show that the iliofibularis muscle of C. alboguttata is not a homogenously oxidative muscle (Table 1). If oxidative fibres are more susceptible to muscle disuse atrophy than glycolytic fibres, we would anticipate that the oxidative fibres would have been preferentially affected over the glycolytic fibres. In actuality, the significant reduction in fibre cross-sectional area in the iliofibularis muscle of 9-month aestivators occurred in both oxidative and glycolytic fibre types (Fig. 3A).
The enzyme histochemistry component of this study does not support the hypothesis that prolonged aestivation induces the transition of fibre types from characteristically (histologically) slow, oxidative to fast, glycolytic. The relative proportions of the two fibre types, measured as a percentage of total number of fibres present in the muscle cross-section, was consistent between controls and 9-month aestivators (Table 1). Furthermore, the size-distribution of fibres, as measured by cross-sectional area, remained consistent between controls and 9-month aestivators (Fig. 4A–D). There was a trend for larger iliofibularis glycolytic fibres (>10 000 µm2) to occur more frequently after 9 months of aestivation; however, it was not significant (Fig. 4B).
Muscle mechanics
Previous research has demonstrated that prolonged aestivation has no effect
on skeletal motor nerve terminals or membrane potentials in C.
alboguttata (Hudson et al.,
2005); however, the effect on muscle mechanics was more complex.
Twitch activation, twitch relaxation and tetanus relaxation times were
significantly longer in 9-month aestivator iliofibularis muscle, but there
were no significant changes in sartorius muscle
(Table 2). The findings for the
faster twitch sartorius muscle concur with previous work on 3-month aestivator
C. alboguttata gastrocnemius muscle
(Hudson and Franklin, 2002a),
suggesting that the isometric kinetics of relatively fast muscles are
unaffected by aestivation. The apparent slowing of isometric kinetics of the
relatively slower twitch iliofibularis muscle was contrary to the findings of
most previous studies (Maier et al.,
1973; Nordstrom et al.,
1995) and the reverse of our hypothesis. However, the slowing of
isometric kinetics in our study was consistent with other findings (Rourke,
2004), which showed a non-significant tendency for the expression of myosin
heavy chain protein isoforms to shift towards slower isoforms in hibernating
ground squirrels. The observed fast-to-slow transition in squirrels may be due
to shivering thermogenesis during periodic arousal from hibernation; however,
this is clearly not the case for an amphibian aestivator.
There were no significant differences in isometric stress between controls
and 9-month aestivators for either sartorius or iliofibularis muscles
(Table 2). This is similar to
the results found previously in 3-month aestivator C. alboguttata
gastrocnemius muscle (Hudson and Franklin,
2002a). 12–16 weeks of aquatic hibernation in Rana
temporaria has also been previously shown not to affect sartorius muscle
isometric stress (West et al.,
2006), therefore skeletal muscle stress appears to be conserved in
the natural models of anuran dormancy studied so far.
There were no aestivation-related changes to power output-cycle frequency
curves in the slow-twitch iliofibularis of C. alboguttata, but the
fast-twitch sartorius became slower after 9 months of aestivation
(Fig. 5B), which is contrary to
the results obtained from previous studies
(Diffee et al., 1991) and the
reverse of our hypothesis. This result may have been indicative of fibre-type
remodelling of the sartorius, though there was no evidence to support this
(Table 1). In contrast,
sartorius muscle isovelocity power output remained stable after 12–16
weeks of aquatic hibernation in Rana temporaria, suggesting that the
muscle mechanical properties of over-wintering frogs remain stable. 9 months
of aestivation had no significant effect on fatigue resistance in either the
iliofibularis or sartorius muscles (Fig.
6A,B), which was similar to the findings in the gastrocnemius
muscle from 3-month aestivator C. alboguttata
(Hudson and Franklin,
2002a).
Within the present study, the effect of aestivation on isometric kinetics
of skeletal muscle partially contradicts the findings from work-loop
assessment of mechanical properties. However, previous studies have
demonstrated that isometric kinetics and force–velocity properties alone
are poor predictors of work-loop shape
(Caiozzo, 2002;
James et al., 1996). Work-loop
shape is also affected by force enhancement, shortening deactivation and
passive properties of muscle (Josephson,
1993). Therefore, the work-loop technique can give a more
functionally relevant estimate of muscular performance capabilities by
assessing performance under dynamic conditions
(Josephson, 1993).
The apparent slowing of both muscle types, as evidenced by the isometric
contractile properties data, indicated that prolonged aestivation did not
induce a transition from slow to fast twitch in the iliofibularis and
sartorius muscles. As such, prolonged aestivation may have an effect on muscle
physiology or biochemistry that indirectly changes the isometric properties of
the muscle tissue without affecting fatigue resistance. Changes to muscle
mitochondrial structure, function or density may be one possible explanation,
though it has been demonstrated that mitochondrial density is not correlated
with contraction speed and fatigability in individual fibres
(Lipska et al., 1998). A
decline in the energetic stores of the individual muscles, rather than fibre
type changes, may be responsible for this outcome; however, levels of muscle
metabolites, such as phosphorylated creatine and ATP, are known to remain
stable in the muscles of cold-submerged hibernating R. temporaria
(West et al., 2006), though it
should be noted that the quiescent period of this anuran species is
substantially shorter than that experienced by C. alboguttata.
Furthermore, it has been postulated that one of the mechanisms acting to
protect animals from muscle disuse atrophy during dormancy is low body
temperature, in conjunction with lowered metabolic rate
(Hudson and Franklin, 2002b).
In this way, C. alboguttata is truly remarkable as it is the only
animal model ever investigated that does not exhibit any muscle disuse atrophy
during a temperature-independent dormancy.
Concluding remarks
The present study demonstrated that 9 months of aestivation had little
effect on muscle morphology, histochemistry and dynamic mechanical performance
in C. alboguttata. There was no skeletal muscle atrophy of the type
normally associated with either experimental limb disuse or hibernation. Whole
muscle mass, muscle cross-sectional area, fibre number and proportions of
fibre types remained unchanged after prolonged disuse, and relatively minor
morphological changes to fibre cross-sectional area and density were
observed.
Overall, prolonged aestivation had little effect on the isometric properties of the skeletal muscle of C. alboguttata, with moderate, though significant, changes occurring in the slow-twitch iliofibularis muscle. Though the results of the isometric analysis suggested that the muscles were becoming slower after 9 months of aestivation, the results of the work-loop analysis clearly demonstrate that the contractile properties of aestivating C. alboguttata are maintained at control levels. Despite the fact that significant changes are occurring at the muscle structural level, overall muscle performance, and hence whole animal performance, is preserved. Further investigation is required to elucidate the mechanisms that prevent skeletal muscle disuse atrophy during aestivation.
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Acknowledgments |
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