The authors declared no conflicts of interest.
Obese mice incur greater myofiber membrane disruption in response to mechanical load compared with lean mice†
Version of Record online: 16 MAR 2013
Copyright © 2013 The Obesity Society
Volume 21, Issue 1, pages 135–143, January 2013
How to Cite
Knoblauch, M. A., O'Connor, D. P. and Clarke, M. S.F. (2013), Obese mice incur greater myofiber membrane disruption in response to mechanical load compared with lean mice. Obesity, 21: 135–143. doi: 10.1002/oby.20253
See the online ICMJE Conflict of Interest Forms for this article.
- Issue online: 16 MAR 2013
- Version of Record online: 16 MAR 2013
- Manuscript Accepted: 31 MAY 2012
- Manuscript Received: 12 DEC 2011
Obesity is associated with modified transmembrane signaling events in skeletal muscle, such as insulin signaling and glucose transport. The underlying cause of these obesity-related effects on transmembrane signaling is still unknown. In general, the function of membrane proteins responsible for transmembrane signaling is modulated by the biochemical makeup of the membrane, such as lipid composition, in which they are embedded. Any obesity-related alterations in membrane composition would also be predicted to modify membrane biomechanical properties and membrane susceptibility to mechanical load-induced damage. The primary objective of this study was to investigate whether obesity influences myofiber membrane susceptibility to mechanical damage in skeletal muscle.
Design and Methods:
Myofiber membrane damage was compared between 12-week-old obese, hypercholesterolemic (B6.V Lepob/J) and isogenic, normocholesterolemic control (C57BL6/J) male mice following either normal cage activity or strenuous eccentric exercise (downhill running). Myofiber membrane damage was quantified in perfusion-fixed frozen sections of the gastrocnemius muscle via sarcoplasmic concentration of either albumin (cage activity experiment) or a fluorescent marker that had been injected immediately before activity (eccentric exercise experiment).
Obese mice exhibited evidence of increased myofiber membrane damage compared with lean mice after both normal cage activity and eccentric exercise indicating that myofiber membranes of obese mice are more susceptible to mechanical damage in general and that eccentric exercise exacerbates this effect.
These observations are consistent with the notion that obesity influences the biochemical and biomechanical properties of myofiber membranes.
Obesity is associated with several conditions such as excess body fat, hypercholesterolemia (1), and impaired insulin sensitivity (2), all of which can be attenuated through increased physical activity. It has been suggested that impaired insulin sensitivity may involve disruption of transmembrane signaling within skeletal muscle due to alterations in the lipid composition of muscle membranes induced by obesity (3). Such lipid alterations would likely also influence membrane biomechanical properties (4,5) including those of the sarcolemma/T-tubule and sarcoplasmic reticulum membrane systems of the myofiber. If true, such biomechanical alterations will be reflected by altered myofiber membrane susceptibility to mechanical load-induced damage in obese individuals.
Evidence to support the concept that obesity and associated dyslipidemia modify the biomechanical properties of cell membranes has been reported in a variety of cell types including skeletal myofibers. Both hypercholesterolemia (6) and consumption of a high-fat diet (7) have been shown to influence the biochemical makeup of cell membranes (8,9). For example, hypercholesterolemia resulting from elevated total circulating cholesterol levels (∼300 mg/dl) in humans has been shown to increase both membrane cholesterol content and mechanical shear damage in erythrocytes (10). In skeletal muscle, consumption of a high-fat diet, particularly those diets high in saturated fat, can increase the saturation of fatty acyl chains within the sarcolemma (9). As fatty acyl chain saturation increases, phospholipid packing density within the membrane is also increased (11). This serves to create a more rigid membrane (12,13), a biomechanical property that has previously been shown to increase susceptibility to mechanical load-induced membrane damage in tissue culture cells (14,15).
Whether such obesity-associated effects on cell membrane composition specifically influence myofiber membrane susceptibility to mechanical load-induced damage in vivo remain to be determined. At present, few studies exist which have evaluated myofiber membrane damage between lean and obese individuals. Salvadori et al. (16) reported that obese males exhibit significantly higher circulating levels of creatine kinase (CK), an indirect marker of muscle membrane damage, both before and after maximal exercise when compared with normal weight subjects. In addition, Paschalis et al. (17) showed a similar increase in circulating CK levels in overweight/obese women compared with lean women after identical exercise loading. However, the utility of total circulating CK as an indirect indicator of myofiber membrane damage is limited by interindividual differences in the time to peak circulating CK level after exercise and variations in time to clearance from the circulation (18,19).
In order to directly test the hypothesis that obesity results in altered myofiber membrane susceptibility to mechanical load-induced damage, we have used a mouse model of obesity in which to directly quantify the frequency and severity of myofiber membrane damage in response to activity-induced mechanical load using a combination of myofiber damage tracers and muscle histology (20,21,22). Since obese individuals appear to incur greater myofiber membrane damage both at low and high levels of physical activity compared with normal weight controls (16,17), we assessed myofiber membrane damage under mechanical loading levels associated with both normal cage activity and defined eccentric exercise during downhill running in obese and lean isogenic control animals. Membrane damage in those mice limited to cage activity was assessed using immunolocalization of sarcoplasmic albumin to reveal the degree of constitutive myofiber membrane damage elicited in response to normal cage activity occurring in the 24–48 h before tissue harvest. Separately, mice exposed to eccentric exercise were injected with a fluorescent myofiber membrane damage marker immediately before exercise allowing for direct assessment of myofiber membrane damage, which occurred subsequent to injection of the tracer (i.e., during eccentric exercise).
Methods and Procedures
Procedures were approved by the Institutional Animal Care and Use Committee before data collection. Unless noted, care and processing was identical for all mice. Ten-week-old obese, hypercholesterolemic (B6.V Lepob/J, Jackson Labs, Bar Harbour, ME) and isogenic lean, normocholesterolemic control mice (C57BL6/J; Jackson Labs) were acclimatized on-site for 2 weeks before the beginning of the experiment. Mice were housed without cage exercise equipment, and had access ad libitum to water and a group-specific diet of either standard chow for lean mice or high-fat chow (26% of calories from fat) (Labdiet 5K20; Purina Mills, St. Louis, MO) for obese mice. These group-specific diets were required in order to maintain the obese phenotype associated with the B6.V Lepob/J mice (e.g., increased body weight, elevated circulating levels of total cholesterol) since our pilot data indicated that switching obese animals to standard chow during the 2-week acclimatization period resulted in these variables returning to normal control levels. Two days before killing, a blood sample (∼50 µl) was obtained for spectrophotometric determination of total circulating cholesterol levels. Each animal's weight was determined using a digital scale on the day of each experiment.
Animals assigned to the cage activity condition (six lean, six obese) underwent whole body perfusion for subsequent determination of sarcoplasmic albumin via immunoperoxidase staining. Albumin is a naturally occurring molecule contained in the extracellular fluid of all tissues (21) and is normally impermanent to cell membranes due to its molecular weight (i.e., ∼67 kD) preventing its entry into the myofiber sarcoplasm unless it gains direct entry via mechanical load-induced myofiber membrane damage (21). As such, the presence of albumin in the sarcoplasm of a myofiber is indicative of sarcolemma/T-tubule damage that might have occurred any time in the previous 24–48 h period. This time frame is based on the period it takes for albumin to be cleared from the myofiber sarcoplasm via normal cellular activity.
Animals assigned to the exercise condition (five obese, five lean) were exercised by downhill running on a rodent treadmill after injection of an exogenous fluorescent myofiber damage marker using a modification of a previously described protocol (23). All obese animals exercised at a treadmill slope of −12°. After exercise, the downhill force (N) exerted parallel to the treadmill on the hind limbs of each obese animal was calculated according to its body weight and resolution of the vertical gravity force vector into perpendicular components. For lean animals, the slope of the treadmill during exercise was adjusted so that the downhill force (N) experienced by the hind limbs of each lean animal was identical to that of its pair-matched obese animal. The following formula was used to determine the slope of the treadmill used to exercise lean animals to account for body weight differences between lean and obese groups:
where: −0.208 = sin(−12°), OW = weight of obese (N), LW = weight of lean (N).
Ten minutes before exercise, mice were injected intraperitoneally with 250 µl of sterile phosphate buffered saline (PBS) containing 400 mg/ml of tetra-methyl-rhodamine (TMR)-conjugated, lysine-fixable dextran (10,000 molecular weight) (Invitrogen, Grand Island, NY). This fluorescent tracer is normally impermanent to cell membranes due to its molecular weight, thereby preventing its entry into the myofiber sarcoplasm unless it gains direct entry via mechanical load-induced myofiber membrane damage (20). The presence of this fluorescent tracer within the sarcoplasm of a myofiber indicates that the sarcolemmal/T-tubule membrane of that particular myofiber has suffered a mechanical disruption some time after the tracer has been injected into the animal (i.e., during the exercise period). This is in contrast to the direct immunoperoxidase visualization of sarcoplasmic albumin (i.e., an endogenous tracer), which was used for the evaluation of constitutive sarcolemmal damage that had occurred in these animals due to normal cage activity during the previous 24–48 h.
Ten minutes after injection of the fluorescent marker, TMR-dextran was clearly visible as a dark pink coloration in the feet of the animal indicating systemic distribution of the tracer as has previously been observed using a similar molecular weight fluorescent dextran tracer (20). After this 10-min period during which time the mice were allowed to rest in their cage, animals subsequently performed 18 5-min exercise bouts separated by 2-min rest periods at a treadmill speed of 7 m/min. After completion of the exercise protocol, each mouse underwent whole body perfusion-fixation as described below. Two separate age-matched lean and obese animals were injected with TMR-dextran, returned to their cage for 10 min to allow the tracer to enter the circulation, and then immediately perfusion-fixed without exercise. These animals served as imaging controls to determine the basal level of myofiber damage, which occurred in muscle from animals administered the damage tracer, but not exposed to eccentric exercise.
Lean and obese animals from both the cage activity and eccentric exercise groups underwent whole body perfusion-fixation using intracardiac perfusion as described previously (20). In brief, mice were deeply anesthetized using 80 mg/kg of pentobarbital administered intraperitoneally followed by intracardiac perfusion at a perfusion rate of 6 ml/min using first 50 ml of warm 37 °C Ca+/Mg+-free PBS (pH 7.2) containing 0.1% (w/v) procaine followed immediately by 50 ml of Ca+/Mg+-free PBS (pH 7.2) containing 2% (w/v) formaldehyde. After 1 h, the gastrocnemius was dissected out and placed in fresh Ca+/Mg+-free PBS (pH 7.2) containing 2% (w/v) formaldehyde and postfixed by immersion for an additional 12 h. Next, the muscle tissue was sequentially immersed for 12 h each in Ca+/Mg+-free PBS (pH 7.2) containing 10% (w/v), 20%, and 30% sucrose followed by Ca+/Mg+-free PBS (pH 7.2) containing 30% sucrose and 50% (v/v) optimal cutting temperature (OCT) medium (Sakura Finetek, Torrance, CA). Finally, the muscle was placed in OCT for 3 h before freezing via slow immersion in liquid-nitrogen-cooled isopentane in preparation for frozen sectioning.
Sectioning was performed on a Shandon Cryotome SME operating at a −20 °C chamber temperature. Multiple 10 µ-thick frozen cross-sections of the belly of the gastrocnemius muscle were collected onto Superfrost Plus treated slides, with a minimum of 50 µ separating each section. Slides were stored at −80 °C until processed. For quantification of myofiber membrane damage during cage activity (by immunoperoxidase staining of sarcoplasmic albumin) or after eccentric exercise (using direct fluorescent visualization of sarcoplasmic TMR-dextran), a total of three complete cross-sections, each separated by at least 300 µ, of the belly of the left gastrocnemius muscle of each animal were used.
Immunoperoxidase staining for sarcoplasmic albumin
All slides underwent batch-processing in Coplin staining jars at room temperature unless otherwise noted using a modification of a previously reported protocol (21). Frozen sections were washed twice in Dulbecco's PBS (D-PBS) containing 1 mmol/l Ca2+ and 1 mmol/l Mg2+ (pH 7.2) followed by a 5-min incubation in D-PBS containing 0.1% (v/v) Triton-X 100. Sections were then washed over 5 min using two changes of wash buffer comprised of D-PBS (pH 7.2) containing 0.05% (v/v) Triton-X 100. Next, sections underwent a 30-min blocking for endogenous peroxidase activity using D-PBS containing 2.0% (w/v) sodium azide and 0.3% (v/v) H202 followed by a 5-min incubation in wash buffer and a 1-h blocking step for nonspecific protein binding using wash buffer containing 4% heat-inactivated goat serum. After washing with two changes of wash buffer over a period of 5 min, sections were incubated for 2 h with primary antibody solution consisting of 20 µg/ml of horse radish peroxidase conjugated goat polyclonal to mouse serum albumin (#19195; Abcam, Cambridge, UK) in wash buffer containing 1% (v/v) heat-inactivated goat serum at 37 °C in a humidified chamber. Sections were then washed five times over 30 min with wash buffer and twice with 100 mmol/l Tris buffer (pH 7.0) before incubating at room temperature in the dark for 5 min in diaminobenzidine chromagen/substrate (Invitrogen #88-2014). Slides then underwent five washes in fresh D-PBS before mounting in Fluoromount-G mounting medium and sealing the cover-slip to the slide with clear nail polish. Negative control slides were also prepared using identical processing except that they were not incubated with primary antibody.
Fluorescence slide preparation (exercise group)
Upon removal from frozen storage, sections from exercised mice were rehydrated in D-PBS (pH 7.2) for 5 min in dark before mounting in Fluoromount-G and sealing the cover-slip to the slide with clear nail polish.
Sections were imaged using an Olympus BX-70 epifluorescent microscope with attached DP72 camera operated using CellSens camera control software (Olympus America, Center Valley, PA). Collected images captured all myofibers within the central region of the gastrocnemius (Figure 1), which resulted in an average of 9,760 myofiber cross-sectional profiles analyzed per lean mouse and 9,529 myofiber cross-sectional profiles per obese mouse. All images of immunoperoxidase stained sections were captured by light microscopy under identical photographic conditions using 10× magnification, 0.24 aperture, 1-ms camera exposure, and an ISO sensitivity of 200. Fluorescence images were captured under identical photographic conditions using a TRITC filter set (Chroma Technology, Bellows Falls, VT) at 20× magnification, 90 ms exposure, 0.24 aperture, and ISO sensitivity of 200. Images were saved in .jpg format for subsequent image analysis.
Myofiber membrane damage in the gastrocnemius of lean and obese mice was quantified through image analysis by determining the sarcoplasmic staining intensity of albumin under cage activity conditions or the fluorescent signal intensity of sarcoplasmic TMR-dextran after eccentric exercise. Images were imported into ImageJ v. 1.44 (NIH) software and converted to an 8-bit grayscale. This creates individual pixel intensities from 0–255 in that a pixel value of zero (0) represents absence of light (i.e., “black”) and 255 is fully bleached (i.e., “white”). For immunoperoxidase staining, a greater staining intensity corresponds to a darker gray scale value whereas for fluorescence imaging, higher levels of TMR-dextran signal correspond to a higher grey scale value (i.e., brighter fluorescence).
Mean pixel intensities per myofiber were determined by averaging individual pixel intensities within a user-defined area located within the myofiber sarcoplasm. Using ImageJ software v1.44p (NIH) in conjunction with the Time Series Analyzer (version 2.0; NIH, http://rsb.info.nih.gov/ij/) plug-in, a 40 × 40 pixel oval was placed centrally within each myofiber. Areas exhibiting intramyofiber tears or areas of the muscle exhibiting folds or processing artifacts were not included within the frame of the analysis oval. Mean pixel intensities for all myofibers within the area of interest per animal were then binned according to their mean gray scale intensity (0–255), generating a myofiber gray scale frequency distribution per animal. Because myofiber count varied per animal, the myofiber gray scale frequency distribution was normalized by total myofibers per animal, generating a percent of total myofibers at each gray scale value per animal. These binned percentages were then averaged across six lean or six obese for the cage activity experiment and five lean and five obese for the exercise experiment, creating an aggregate “damage profile” for each group.
For the defined exercise experiment, determination of damaged vs. undamaged myofibers was performed by identifying the 90th percentile of the profile in the unexercised lean mouse that received fluorescent dextran (i.e., lean imaging control) to account for myofiber damage, which may have occurred during the 10 min of cage activity before the beginning of the eccentric exercise protocol. All myofibers having a fluorescence intensity above this 90th percentile were considered to have suffered myofiber membrane damage as previously described (20,21).
Two-sample Kolmogorov–Smirnov tests were used to compare the damage profile distributions between lean and obese mice in both the cage activity and exercise groups, with statistical significance set at P < 0.05. To evaluate whether the degree of load experienced by the hind limbs of the animals during downhill running was related to the frequency of myofiber damage, we plotted the proportion of damaged myofibers (i.e., those myofibers above the 90th percentile) per animal within each group (lean and obese) against downhill force placed upon the muscle.
Mean body weight (g) and total circulating cholesterol levels (mg/dl) per group are reported in Table 1 indicating both significantly higher body weight and total circulating cholesterol levels in obese compared with lean mice at time of tissue collection.
Effect of obesity on myofiber damage during normal cage activity
Figure 2 depicts representative light microscopy images of damaged myofibers labeled with sarcoplasmic albumin in cross-sections of the gastrocnemius muscles from lean and obese animals under normal cage activity conditions. Negative control sections from both lean (Figure 2a) and obese (Figure 2b) mice showed no significant staining. Sarcoplasmic staining of albumin is evident in the myofibers of both lean (Figure 2c) and obese (Figure 2d) mice, with those myofibers suffering the greatest amount of membrane damage having the highest intensity of sarcoplasmic albumin staining.
Comparison of the myofiber damage profiles obtained from lean and obese mice under normal cage activity conditions indicated that obese mice had significantly more (D = 23.73, P < 0.001) damaged myofibers than observed in lean animals (Figure 3). Average albumin staining intensity within myofibers of the gastrocnemius muscle of lean animals was 178.3 (±27.9) compared with 162.3 (±22.8) in obese animals (lower gray scale values representing higher staining intensity). These data also indicate that during normal cage activity the myofiber membrane damage inflicted in obese animals is more severe than in lean animals, as evidenced by the higher frequency of myofibers exhibiting more intense sarcoplasmic albumin staining (i.e., lower gray scale values) in obese compared with lean animals (Figure 3).
Effect of obesity on myofiber damage during eccentric exercise
Figure 4 depicts representative fluorescent images of damaged myofibers labeled with sarcoplasmic TMR-dextran in cross-sections of the gastrocnemius muscles from lean and obese animals. Low levels of sarcoplasmic TMR-dextran signal are evident in the sarcoplasm of both lean (Figure 4a) and obese (Figure 4b) mice exposed to the fluorescent damage marker for 10 min and then perfusion-fixed without eccentric exercise. These results indicate that some myofiber membrane disruption occurs even in the 10 min period of cage activity during which these animals were exposed to TMR-dextran before perfusion-fixation, an observation consistent with our findings in cage activity animals using sarcoplasmic albumin as a myofiber membrane damage marker (Figures 2 and 3). However, a much higher frequency of wounded myofibers exhibiting a higher sarcoplasmic TMR-dextran fluorescent intensity than detected in unexercised animals (Figures 4a,b) is observed in both lean (Figure 4c) and obese (Figure 4d) animals, which underwent eccentric exercise. These data indicate that the TMR-dextran tracer revealed the low levels of myofiber membrane damage, which occurred during the 10 min of cage activity before exercise and the much more frequent and severe myofiber membrane damage, which occurred during the eccentric exercise protocol.
The frequency of myofibers having specific fluorescent TMR-dextran signal intensities (gray scale) in each animal was plotted to generate average myofiber damage profiles in both lean and obese animals exposed to eccentric exercise (Figure 5). A signal intensity (gray scale) value of 17, the 90th percentile in an unexercised lean animal exposed to TMR-dextran (Figure 4a), was defined as the threshold to identify damaged myofibers (Figure 5a).
Using this threshold gray scale value, the myofiber damage profile from an obese mouse exposed to TMR-dextran for 10 min without eccentric exercise and immediately perfusion-fixed was significantly different (D = 30.0, P < 0.001) from the unexercised lean mouse (Figure 5c). Whereas 10% of myofibers (with an average sarcoplasmic fluorescent signal of 24.0) were deemed to have suffered myofiber damage in the unexercised lean animal exposed to TMR-dextran for 10 min, the same 10-min exposure to TMR-dextran in the unexercised obese animal resulted in 38% of myofibers (with an average sarcoplasmic fluorescent signal of 26.5) being identified as damaged. These data indicate that the incidence of myofiber damage in obese animals even during this short period of cage activity (i.e., 10 min exposure to TMR-dextran before perfusion fixation) is significantly greater than that observed in lean animals. These data parallel those obtained using sarcoplasmic albumin staining as a marker of myofiber membrane damage in lean and obese animals under normal cage activity conditions (Figure 3).
Among the five exercised lean mice, 41.7% (±19%) of the myofibers of the central portion of the gastrocnemius muscle were identified as having suffered damage, with an average sarcoplasmic fluorescence signal intensity of 26.8 (±1.51) (Figure 5a). The damage profile from the five exercised obese mice shows that 75.8% (±9%) of the myofibers of the central portion of the gastrocnemius muscle were identified as having suffered damage, with an average sarcoplasmic fluorescence signal intensity of 30.8 (±5.0) that exhibited a significantly different (D = 24.79, P < 0.0001) damage profile compared with exercised lean mice (Figure 5b). These data indicate that despite performing eccentric exercise activity under identical load, obese mice incur significantly greater myofiber membrane damage after exercise than lean mice.
Comparison of the shape of the damage profiles obtained from unexercised animals using TMR-dextran as the damage marker indicates that there is a distinct population of wounded myofibers observed in the unexercised obese animal that does not appear in the unexercised lean animal (Figure 5c). When all four damage distributions (lean unexercised, obese unexercised, lean exercised, and obese exercised) are plotted on a scale restricted to the damaged fiber range, it can clearly be seen that in the obese animals this distinct population of damaged myofibers at a gray scale value of ∼25 increases in frequency after eccentric exercise (Figure 5d), whereas a second more severely damaged myofiber population at a gray scale value of ∼32 also appears (Figure 5d). In contrast, eccentric exercise in lean animals induces the appearance of a wounded population of myofibers similar to the damage profile observed in the unexercised obese animal (Figure 5d).
Effect of loading force on myofiber membrane damage in exercised mice
To determine whether myofiber membrane damage was dose dependent relative to the load placed upon each mouse during eccentric exercise, we plotted the percent of damaged myofibers per mouse against the force vector load in Newtons (N) per animal (Figure 6). Load showed a higher correlation with percent of myofibers damaged in the lean (R2 = 0.75) than in the obese mice (R2 = 0.24). This indicates that factors other than the load imposed on the myofiber membranes likely influenced the increased severity of myofiber membrane damage, which occurred during eccentric exercise.
The purpose of this study was to investigate whether obesity is associated with an increased susceptibility to mechanical load-induced myofiber membrane damage in skeletal muscle. Increased susceptibility to myofiber membrane damage may be reflective of obesity-induced changes in the biochemical/biomechanical properties of myofiber membranes as has previously been reported for other cell types in obese individuals. Previous work has shown that obese humans exhibit higher circulating levels of CK both at rest and after strenuous exercise (16), indirectly indicating increased myofiber membrane disruption. Our data obtained in a mouse model of obesity at the tissue level using direct histological examination of individual myofibers exposed to loading at normal cage activity levels or defined eccentric exercise loads provide direct evidence that myofibers in obese muscle are more susceptible to mechanical load-induced damage. This finding in obese mice supports the concept that the increased circulating level of CK observed after exercise in obese individuals is most likely due to a similar increased susceptibility to mechanical load-induced myofiber membrane damage in obese individuals.
The frequency and severity of myofiber membrane disruption in our obese animals were significantly higher than in lean animals both at loading levels associated with normal cage activity and during eccentric exercise damage. In the cage activity animals, we evaluated the degree of constitutive myofiber membrane damage using sarcoplasmic albumin as a tracer because it is an endogenous protein normally present within the extracellular fluid, which when present in the myofiber sarcoplasm is an indicator of myofiber wound damage. Since it takes ∼24–48 h to clear albumin from the myofiber sarcoplasm, sarcoplasmic albumin signal detected in skeletal myofibers is an “aggregate” measure of the amount of damage that has occurred in the 24–48 h period before tissue fixation and harvest. Using this approach, our data indicate that constitutive myofiber membrane damage is more frequent and more severe than observed in control animals under normal cage activity levels. In order to temporally resolve any differences in the level of constitutive myofiber membrane damage that occurs in all mice under normal cage activity from myofiber membrane damage specifically occurring in response to eccentric exercise, we used a exogenous membrane damage tracer (TMR-dextran) injected into the animal immediately before exercise.
One advantage of using an animal model to study the incidence of myofiber membrane wounding is the ability to introduce a membrane damage marker, such as TMR-dextran, into the circulation of the animal allowing the detection of myofiber membrane disruption in a temporal fashion. Unlike sarcoplasmic albumin staining, which reflects constitutive myofiber membrane disruption that occurred in the previous 24–48 h period, sarcoplasmic TMR-dextran signal detects myofiber membrane damage, which has occurred only during the time period in which the fluorescent damage marker has been present in the tissues. Furthermore, the use of a 10 kD-sized TMR-dextran wound marker in our study allows for increased sensitivity in detecting myofiber membrane damage than can be achieved using alternate larger (i.e., ∼68 kD) tracers such as an Evans Blue Dye albumin complex (24). This increased sensitivity is based on the ability of lower molecular weight membrane damage tracers (e.g., 10 kD TMR-dextran) to enter the myofiber sarcoplasm via smaller myofiber membrane tears or disruptions than higher molecular weight membrane damage tracers (e.g., 68 kD Evans Blue Dye albumin complex).
Our data provides direct histological evidence that these wounding events occur both at a higher frequency and are more severe in the myofibers of obese as compared with lean animals even under loading levels associated with normal cage activity. For example, the incidence and severity of myofiber membrane damage detected in unexercised obese compared with unexercised lean muscle after an identical 10-min exposure to TMR-dextran is significantly greater in the obese animal (Figure 5c). A similar increase in susceptibility to myofiber membrane damage was detected using sarcoplasmic albumin staining (reflecting damage that occurred in the previous 24–48 h) in obese compared with lean animals maintained under normal cage activity (Figure 3). These data indicate that obesity in this animal model increases the susceptibility of myofiber membranes to mechanically induced damage at low levels of mechanical loading experienced during cage activity.
Also of interest was our finding that high levels of mechanical loading experienced during eccentric exercise appears to affect the myofiber membrane damage response in two distinct populations of myofibers as detected using the temporal damage marker, TMR-dextran. When lean animals are exercised, this distinct population of wounded myofibers appears in the damage profile (Figure 5d). This damaged myofiber population appears to be pre-existing in unexercised obese muscle (Figure 5d), corresponding to myofiber membrane wounding, which occurs in obese muscle at normal cage activity levels. After eccentric exercise in obese animals, the frequency of this pre-existing population of wounded myofibers is increased whereas an additional distinct population of wounded myofibers with more severe myofiber membrane damage appears in the obese animals (Figure 5d). Our data suggests that different populations of myofibers within the gastrocnemius muscle may have different levels of susceptibility to mechanical load-induced myofiber membrane damage, and that obesity exacerbates this susceptibility.
Implications of obesity's effect on myofiber membrane disruption
The hypertrophic cascade associated with strength training in skeletal muscle may be facilitated by myofiber membrane disruption. Satellite cells, the muscle-specific stem cell, lie immediately adjacent to the sarcolemma (25) and are thought to be activated in response to mechanical loading activity (26,27,28). Satellite cell activation is observed after a single strenuous exercise bout (26), with the highest frequency of activated satellite cells being observed within muscles containing the highest proportion of damaged myofibers (29). The muscle growth factor, fibroblast growth factor, is also efficiently released from damaged myofibers (20) and has previously been shown to cause satellite cell activation (30).
In contrast, evidence suggests that diet-induced obesity (31) evokes an atrophy response rather than a hypertrophic response. The underlying basis of myofiber atrophy is the loss of contractile protein from the sarcomere and/or loss of complete sarcomeres from the myofiber. In healthy resting muscle tissue, extracellular calcium concentrations are ∼10,000 times higher than sarcoplasmic levels (32). The intact sarcolemma serves as a “barrier” to maintain this concentration gradient between the extracellular fluid and sarcoplasmic compartment of the myofiber. Failure to maintain low levels of sarcoplasmic calcium, such as occurs after repeated myofiber membrane disruption, results in many detrimental effects within the myofiber including disruption of excitation-contraction coupling and force production (33) as well as the activation of calcium-dependent proteolytic enzymes such as calpain (34), which triggers selective degradation of contractile and structural proteins within skeletal muscle (34). One consequence of repeated and severe myofiber membrane disruption as observed in our obese animals in response to both normal cage activity and eccentric exercise may be to initiate protein catabolism within the myofiber by activation of calcium-dependent proteolysis of contractile protein. Since repeated myofiber membrane damage results in continued influx of calcium to the myofiber sarcoplasm, we postulate that the myofiber atrophy response previously observed in obese animals (31) and evident in our particular model (preliminary data derived from 100 randomly sampled myofibers in the central region of the gastrocnemius from each of six lean and obese unexercised animals indicated that myofiber cross-sectional area in obese muscle was 985.5 ± 360.2 µm2 as compared with 1465 ± 668.4 µm2 in lean muscle) can be attributed to the increased susceptibility of myofibers in obese muscle to mechanical load-induced myofiber membrane damage.
Obesity-associated conditions influencing membrane structure
We have provided direct evidence that obesity affects the biomechanical properties of myofiber membranes that results in an increase in susceptibility to mechanical load-induced myofiber membrane damage. Such alterations in membrane biomechanics are related to the biochemical composition of the membrane (35,36). Two conditions associated with obesity present in our obese mice, but not our lean mice that may influence membrane composition and function are hypercholesterolemia and consumption of a high-fat diet.
Hypercholesterolemia has been shown to increase membrane cholesterol content within cells exposed to the plasma within the vasculature including erythrocytes, endothelial, smooth muscle, and macrophages (8). Cells normally self-regulate membrane cholesterol within a physiologic range that provides optimal levels of membrane fluidity necessary for membrane function (37). Further increases in membrane cholesterol create a higher packing density of membrane-associated molecules, resulting in a membrane exhibiting decreased fluidity, increased rigidity, and consequent increased susceptibility to mechanical damage (15). The obese mice in the current study exhibited circulating total cholesterol levels nearly three times that of lean mice. This may have resulted in a concomitant increase in uptake and delivery of cholesterol to myofiber membranes as occurs within cells exposed to the plasma during hypercholesterolemia (38). Such an increase in cholesterol content within myofiber membranes could certainly influence membrane biomechanical properties, leading to an increase in susceptibility to mechanical load-induced myofiber membrane damage as has been reported in other cell types (10,15).
Consumption of a high-fat diet similar to that fed to the obese mice (26% of calories from fat) can influence myofiber membrane lipid composition (9). In particular, consumption of a diet high in saturated fat can increase saturation of myofiber membrane fatty acyl chains (9). Similar to membrane cholesterol content, membrane phospholipid acyl chain saturation levels can influence membrane biomechanics (39). Membrane domains high in phospholipids containing saturated acyl chains exhibit a higher phospholipid packing density than domains comprised of unsaturated acyl chains (11) which, like increased membrane cholesterol, can increase membrane packing density resulting in increased membrane rigidity (4). Obese mice in the current study were fed a diet high in saturated fat, which may have contributed to a shift toward a higher proportion of membrane phospholipids containing saturated acyl chains within the membrane systems of their myofibers. Since a higher level of saturated phospholipids within the sarcolemma has previously been shown to increase sarcolemmal membrane rigidity (40), an increase in membrane rigidity due to higher membrane cholesterol content and/or phospholipid packing density may explain the increase in myofiber membrane susceptibility to mechanical damage observed in our obese animals.
In conclusion, we have provided direct histological evidence that obesity increases the susceptibility of skeletal myofiber membranes to mechanical load-induced damage during both normal activity and strenuous exercise. This increased susceptibility to myofiber membrane damage reflects that obesity results in detrimental effects upon the biomechanical and/or biochemical properties of myofiber membranes. In addition, increased myofiber membrane disruption noted in obese mice may lead to unique skeletal muscle adaptation over time, such as obesity-induced myofiber atrophy. Furthermore, such membrane alterations may also provide insight into the altered skeletal muscle transmembrane signaling events, such as insulin signaling, commonly observed in obese individuals. Future work should focus on investigation into the individual factors of obesity, which could influence myofiber membrane lipid composition, such as hypercholesterolemia or saturated fat consumption, which in turn could explain the increased susceptibility to myofiber membrane damage observed in obese mice.
This study was funded in part through a doctoral Graduate Research Grant received by MK through the National Strength and Conditioning Association.
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