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ENVIRONMENTAL, EXERCISE AND RESPIRATORY PHYSIOLOGY
Faculty of Applied Health Sciences, Brock University, St. Catharines, Ontario, Canada
Submitted 15 September 2006 ; accepted in final form 12 January 2007
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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isolated extensor digitorum longus muscle; isolated soleus muscle; lactate metabolism; muscle energy charge; hyposmotic; hyperosmotic
The metabolic generation or disposal of osmotically active substances alters cellular osmolality by either introducing the products of catabolism into the internal milieu or by integrating monomers during anabolism and consequently influencing cell volume (8, 17). Although the intracellular pools of these substrates are large, the concentration in the blood is relatively low. In this way, metabolic processes influence the osmolality of the internal milieu through anabolism or catabolism and ultimately create osmotic gradients that result in equilibrating water transport (8).
To alleviate the strain caused by anisosmotic conditions, cells have developed regulatory volume increase (RVI) and decrease (RVD) mechanisms (14). For example, cells exposed to hyperosmotic medium initially shrink, but within minutes attempt to regain their original cell volume by accumulating electrolytes (22). Recent studies have shown that RVI and RVD may act as potent second messenger signals for cellular metabolism and gene expression (22). The incomplete return to the original cell volume by these mechanisms (RVI and RVD) is thought to play a role in the regulation of cell function. Recent studies in myotubes have demonstrated that alterations in myotube cell volume can alter both glycogen synthesis and glutamine transport (28, 31). This suggests that skeletal muscle may regulate its cell volume to assist in the control of metabolism. Various examples of this possible form of metabolic control have been indirectly observed. For example, studies in diseased patients have shown that a relationship exists between water content and nitrogen balance in humans, indicating that muscle water content may affect proteolysis (9, 19, 34). Additionally, whole body studies have examined the influence of changes in plasma volume (36, 40) and osmolarity (2); however, it remains unclear what influences these compartmental volume changes may have at the cellular level with regard to skeletal muscle cell volume and metabolism.
It has long been established that skeletal muscle fibers adjust their volume in response to alterations in the osmolality of the extracellular environment (3). There is limited evidence suggesting that changes in skeletal muscle cell volume affect contractile function in amphibians and possibly mammals (13, 33). However, what influences acute alterations in cell volume have on cellular function and metabolism have not been thoroughly investigated in intact mammalian skeletal muscle. Therefore, the purpose of the present investigation was to develop a resting in vitro organ bath-isolated mammalian skeletal muscle model that would be utilized to verify muscle viability in varying osmolalities and to investigate how these varying osmolalities affect resting cell volume and metabolism. It was hypothesized that a hyperosmotic extracellular environment would lead to a decrease in cell volume and macromolecular crowding, which would lead to alterations in the energy charge of the muscle [i.e., increased breakdown of phosphocreatine (PCr), decreased energy charge] as seen in experiments using hepatocytes, whereas a hyposmotic extracellular environment would lead to an increase in cell volume and an anabolic metabolic state (improved energy charge).
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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Experiments were conducted on two different cohorts of male Long-Evans rats (n = 40) (bred in the Brock University Animal Care Facility) with an overall mean mass of 135.3 ± 22.2 g (
3–6 wk old). The first cohort had a mean mass of 122.1 ± 10.9 g (n = 28), and the second cohort had a mean mass of 161.8 ± 13.0 g (n = 12). All experimental procedures and protocols were approved by the Brock University research subcommittee on animal care and conformed to all Canadian Council on Animal Care guidelines. Animals were housed in groups of four within the Brock University Animal Facility where they were maintained on a 12:12-h light-dark cycle at 22°C. The rats were fed a standard rodent diet (5012 rat diet; PMI Nutrition, Brentwood, MO) and had ad libitum access to food and water up until the experimental protocol. All experiments were conducted at the same time of day, to ensure minimal variability in the activity state of the animals.
Experimental Protocol
Two anisosmotic conditions were used along with a control condition to determine the effects of extracellular osmolality on resting intact skeletal muscle. The incubation medium (Sigma medium 199) was manipulated with either distilled water or mannitol to achieve an osmolality of 290 ± 10 mmol/kg (control), 400 ± 10 mmol/kg (hyperosmotic), or 190 ± 10 mmol/kg (hyposmotic). These manipulations did not vary in ionic content; the only difference across the three conditions was the amount of water that could be imported in or out of the cell. Pre- and postosmolalities of all incubation media were verified with a vapor pressure osmometer (VAPRO5520; Wescor, Logan, UT).
On each experimental day, the animals had ad libitum access to food and water before they were anesthetized (pentobarbital sodium; 55 mg/kg ip). Soleus and extensor digitorum longus (EDL) muscles were removed, ensuring a clean dissection from tendon to tendon. Upon full exposure of both tendons, sutures (4-0 silk) were tied in situ, and then the muscle was removed and placed immediately in an organ bath. Once all muscles of interest were removed, the animals were killed with an overdose of pentobarbital sodium.
Once the muscle was placed in a tissue bath, one silk-sutured end was attached to a force transducer (Grass Telefactor, West Warwick, RI) to monitor the resting tension, whereas the other sutured end was attached to a glass tissue support rod; this secured the muscle in the tissue bath (Radnoti Glass Technology, Monrovia, CA), which contained 15 ml of incubation medium maintained at a constant temperature of 30 ± 2°C . All muscles were placed and secured in the organ bath in under 45 s. Resting tension was adjusted to 1 g (6), and the tissue was continuously perfused with a gas mixture of 95% O2-5% CO2. The resting muscle was then incubated in the organ bath for 60 min.
Assessment of Fluid Shifts
After the 60-min incubation, EDL or soleus muscles were rapidly removed from the bath; visible connective tissue was removed, and then muscles were cut into two pieces, with one piece being snap frozen in liquid nitrogen for water weight analysis and the other piece being mounted for histochemical analysis (see below). These two procedures were completed in <20 s. One muscle (EDL and soleus) from each rodent from the first cohort was used to assess fluid shifts and mounted for histochemical analysis (n = 28 rats). Both muscles (2 EDL and 2 soleus) removed from the rodents in the second cohort were used to assess fluid shifts and for the determination of histochemical properties (n = 12 rats).
Histochemical analysis.
Briefly, the muscle was embedded horizontally in embedding medium (Cryomatrix, Pittsburgh, PA) on a piece of cork and plunged into isopentane (–155°C), cooled in liquid nitrogen. After a rapid freezing, the samples were stored at –80°C for later sectioning and analyses. Before they were sectioned, the muscles were reoriented to obtain a cross section of the muscle. Sectioning was completed with a cryotome (ThermoShandon, Runcorn, Chesire) optimally set at –22°C. A total of nine transverse serial sections 10 µm thick were obtained from each specimen for hematoxylin and eosin staining (to ensure proper orientation) and myosin ATPase or azure A ATPase staining for both type I and type II fibers (4). Myosin ATPase fiber types were determined by the differential staining resulting from preincubation at either pH 10.3 or 4.5. Because it was difficult to obtain useable type I muscle fibers with myosin ATPase staining, only type II fibers were obtained from the first nine muscle samples from each condition per muscle group (n = 27 rats, 1 EDL and 1 soleus per rat, 9 rats per group). A second cohort was required to acquire type I fibers for analyses. For the second cohort, the staining procedure was reevaluated and changed to an azure A ATPase stain. Azure A ATPase staining at pH 4.5 was used to determine both type I and type II skeletal muscle fibers (minimizing the amount of tissue required) from the last eight muscle samples from each condition for each muscle group (n = 12 rats, 2 EDL and 2 soleus per rat, 3 rats per group).
Stained muscle sections were examined under a bright-field microscope (Nikon Eclipse 80i, Chiyoda-ku, Tokyo). Digital images of the slides were captured with a digital camera (Retiga 1300, QImaging, Burnaby, BC, Canada) attached to the bright-field microscope. The images were used to determine the change in muscle cell size (myosin ATPase and azure A) due to the anisosmotic conditions. A total of five to nine fields of view were captured at a magnification of x40. Muscle cross-sectional areas of type I and II fibers (>100 dominant muscle fibers/sample and >45 nondominant muscle fibers/sample due to the differences in fiber type distribution between the soleus, predominately type I, and EDL, predominately type II) were determined with a computer and imaging software (SimplePCI 5.3; Compix, Cranberry Township, PA).
Muscle water content. The snap frozen muscle samples were weighed before and after lyophilization to determine wet-to-dry ratios. These procedures were only carried out on low humidity (<45%) days to ensure that environmental conditions would not affect muscle weights. The differences in weight were used to estimate muscle water content due to the different perturbations, and the relative water content of each sample was determined based on these differences. Relative water content was determined (assuming connective tissue was similar in all muscles) by use of the following formula: [(wet weight – dry weight) x wet weight–1] x 100.
Metabolite analysis. After the lyophilized muscle was reweighed, it was powdered and any visible blood and connective tissue were removed and then acid extracted for measurement of muscle metabolites [ATP, PCr, creatine (Cr), and lactate]. A sample of extracellular medium was also acid extracted for the determination of extracellular lactate. Both muscle and medium metabolite content were determined by fluorometric techniques according to the procedures described by Harris et al. (15) and modified by Green et al. (12). Each sample was analyzed in triplicate during the same analytical session for each of the measured metabolites.
Statistical Analysis
A one-way ANOVA was conducted on the data set for each dependent variable within one muscle group. A two-way repeated-measures ANOVA was conducted to see whether differences existed in pre- and postextracellular osmolality values between both muscle groups. A two-way ANOVA was also used to determine whether water content between the two muscle groups was different as a result of the experimental manipulations. If statistical significance (P < 0.05) was observed, a Tukey's post hoc test was conducted to determine specific pair-wise differences. Experiments were done on two separate cohorts of rats, and the mean body weight of the two cohorts was found to be significantly different; therefore, a correlation was conducted between animal weight and the cross-sectional area of type I and type II fibers. Animal weight and the cross-sectional area of type II fibers were correlated in both muscle types; therefore, when we analyzed the cross-sectional area of type II fibers, animal weight was used as a covariate to eliminate the confounding effect of weight on fiber cross-sectional area. Analyses of covariance were performed on fiber cross-sectional areas for type II fibers in both muscle types. All statistical analyses were completed with SPSS 10.0 (Statistical Package for the Social Sciences, SPSS, Chicago, IL).
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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The mean weight for the rats was similar between all conditions (Table 1). Pre- and postincubation medium osmolalities did not differ in any of the experimental conditions within a given muscle group or experimental group (Table 2). Furthermore, when the pre- and postosmolality measurements of the extracellular media from the soleus and EDL were compared, there were no differences between the muscle groups.
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The observed distribution of skeletal muscle fiber types within the soleus was 83 ± 7% type I and 17 ± 7% type II fibers. Within the EDL, the observed distribution of the skeletal muscle fibers was 7 ± 4% type I and 93 ± 4% type II. Hyposmotic conditions resulted in an increase in water content compared with control conditions (P < 0.05) for both the EDL and soleus (Fig. 1). Hyperosmotic conditions resulted in a decrease in water content compared with control conditions (P < 0.05) for both the EDL and soleus (Fig. 1). The mean cross-sectional area of the soleus type I fibers placed in the hyposmotic condition was significantly greater than that for control (P < 0.01; Fig. 2). No other changes in type I fibers were observed in the remaining conditions. There were no changes in the cross-sectional area of the EDL type I fibers in any condition. A correlation was observed between animal weight and the cross-sectional area of type II fibers in both the EDL (r = 0.50, P < 0.05) and soleus (r = 0.56, P < 0.05). Therefore, weight was considered as a covariate in the cross-sectional area analyses of type II fibers. When weight was considered as a covariate, an increase in adjusted mean (variance from difference in animal weight removed) cross-sectional area of type II fibers was observed in the EDL when incubated in hyposmotic compared with control conditions (P < 0.05; Fig. 2). Alternatively, the EDL muscle placed in hyperosmotic conditions showed a decrease in adjusted mean cross-sectional fiber area of type II fibers compared with control (P < 0.05). The type II fibers in the hyposmotic soleus (P < 0.05; Fig. 2) showed results similar to the hyposmotic EDL (P < 0.05; Fig. 2). Although there was a change in the soleus hyperosmotic water content, there was no change in the soleus hyperosmotic mean fiber area of type II fibers.
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High-energy phosphagen compounds (ATP, PCr) were affected by the hyperosmolar condition (Table 3). EDL ATP content was lower in muscles incubated in the hyperosmotic medium (P < 0.01), whereas a trend was observed for ATP to be lower in the soleus when incubated in the hyperosmotic medium (P = 0.056) compared with control and hyposmotic (Table 2). Muscle PCr was also lower, whereas muscle Cr was higher in EDL (P < 0.05) and soleus (P < 0.05) during hyperosmotic conditions compared with both control and hyposmotic. Total Cr was similar among all of the experimental conditions for both the EDL and soleus. Moreover, no changes in muscle metabolites were observed between the hyposmotic and the control condition among either the EDL or soleus.
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DISCUSSION |
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Effectiveness of Water Movement
The osmolalities employed in this experiment caused fluid shifts to occur in both the hyposmotic and hyperosmotic conditions, as demonstrated by the increase and decrease in wet-to-dry weight ratios, respectively. In both muscle types, there also was an 3% increase and decrease in total muscle water content in each condition, respectively. However, this is lower than the 12% cell volume changes previously observed in hepatocytes (38). Although the changes in muscle water content were small, changes of <10% of the initial volume in hepatocytes have been shown to alter the metabolic environment (26, 38).
Another effect of the anisosmotic conditions was the altering of muscle cross-sectional area of both type I and II fibers. More specifically, EDL type II fibers were altered by hyposmotic and hyperosmotic conditions, whereas soleus type I and type II fibers were affected by the hyposmotic environment. This suggests that water movement was not only into the interstitial space but also directly into the skeletal muscle fibers' cytoplasm of some muscles and specific fiber types. The effect of osmotic strength on cross-sectional area and volume in resting amphibian muscle fibers has previously been established (3, 7). The changes in cross-sectional area observed in the EDL type II fibers (altered by hyposmotic and hyperosmotic) as well as the soleus type I and type II fibers (both were affected by hyposmotic) support the findings from the water content observations that the different osmotic conditions resulted in fluid shifting into and out of the skeletal muscle fibers.
Mechanisms that have been found in other tissues to regulate cellular volume and control fluid shifts are also present in the skeletal muscle sarcolemma. For example, the sodium potassium two-chloride cotransporter (NKCC) is activated on cell shrinkage, helping the cell to increase its volume back to its original state (i.e., RVI) (26). Although, both fast- and slow-twitch muscle phenotypes contain NKCC cotransporters, studies have revealed that their functional influence is greater in the slow-twitch phenotype (11). Also, the membrane protein aquaporin 4 (AQP4) has been observed in fast-twitch skeletal muscle only thus far (10). The present observations also suggest that type II fibers in the EDL may be influenced to a greater extent by fluid shifts caused by anisosmotic conditions than type I fibers. A key feature in their adaptability may be the difference in AQP4 content between the different fiber types (10). A possible explanation of why there were no visible changes in fiber size in the hyperosmotic soleus condition may be that the soleus is more reliant on the regulatory volume machinery for maintaining cell volume due to its relative lack of AQP4 content compared with the EDL (10). Despite the differences in AQP4 distribution, the RVI is faster and more complete in soleus (type I fibers) than in EDL (type II fibers) (37). Clearly, further work is required to determine the specific mechanisms underlying fluid shifts in mammalian skeletal muscle following exposure to anisosmotic conditions; however, on the basis of the present study, it is evident that fluid shifts can and do occur in this tissue.
Effect of Altering Osmolalities on Energy Charge
Both the soleus and EDL experienced a decrease in high-energy phosphagen compounds in the hyperosmotic condition, which has also been demonstrated in isolated fibers from amphibian muscle (20). One possible rationalization for this finding is that high osmolalities are detrimental to cellular function regardless of adequate oxygen perfusion. Past studies have shown that osmolalities in excess of 500 mmol/kg can induce apoptosis in some cell types (23, 30). However, our hyperosmotic condition (400 mmol/kg) more closely resembled osmolalities experienced during exercise (380 mmol/kg) (26) than the more extreme osmolalities that have been observed to induced apoptosis. Based on the present investigation, we can only speculate as to the specific mechanisms that resulted in the lower ATP content. However, two possible mechanisms that may have contributed to the decreased ATP content observed in the hyperosmotic condition are 1) macromolecular crowding (from excessive water loss) and its affect on the intracellular machinery responsible for maintaining energy (i.e., alterations on energy supply pathways) (5) and/or 2) an increase in the metabolic by-products (i.e., lactate and Cr) as a result of the increased activity of anaerobic energy pathways (glycolysis and PCr breakdown) that are affected by alterations in the energy balance of the cell (increased ADP and AMP) (29). Furthermore, the reductions in high-energy compounds (ATP and PCr) may be due to an increased demand on the system to maintain the energy requirements of the cell to sustain cell volume and the intracellular machinery. However, the decreased ATP concentration demonstrates that the energy pathways (glycolysis and PCr) were being challenged to supply the cell with enough energy to meet its requirements during an acute (60 min) exposure to hyperosmotic medium.
An alternative reason for the increase in the metabolites (lactate and Cr) is their possible influence on intracellular osmolality, thereby influencing the movement of water into the cell to alleviate the stress caused by the hyperosmotic perturbation. An increase in metabolic intermediates or an increase in the total numbers of metabolites within the cell would theoretically increase the osmotic charge within the cell, possibly in an attempt to offset the large osmotic gradient and promote cellular water retention. Clearly, further investigations on the mechanisms that lead to a decrease in these high-energy compounds in hyperosmotic medium are required.
Despite the alterations in metabolism with the hyperosmotic condition, no changes in metabolism were observed with the hyposmotic condition. We had originally hypothesized that that hyposmotic conditions would result in a more anabolic environment, perhaps resulting in an improved energy charge within the cells. Based on our present observations, we cannot conclude that the cellular environment was more anabolic, but it is clear that the energy charge is maintained with a hyposmotic extracellular environment. Future experiments will need to evaluate glycogen synthesis and protein metabolism to better understand any potential impact of cell volume on anabolic or catabolic states within skeletal muscle.
Anisosmotic Conditions and Lactate
The experimental manipulation resulted in an increase in muscle lactate in both soleus and EDL muscles incubated in the hyperosmotic medium. There was a 4-fold increase in tissue lactate in hyperosmotic EDL compared with control, whereas the hyperosmotic soleus only demonstrated a 2.5-fold increase compared with control. The accumulation of lactate may serve as a cell volume regulatory mechanism increasing intracellular osmolality, thereby influencing fluid to move into the muscle. Therefore, by creating the energy (glycolysis/glycogenolysis) required by transporters to maintain cell volume, intracellular osmolality could also be increased through the accumulation of lactate. However, what remains unclear is the specific mechanism(s) that resulted in the increased activity of the glycolytic machinery and the accumulation of lactate. Nonetheless, the increase in osmolality due to lactate may be a compensatory mechanism to influence water entry back into the cell as a result of water efflux caused by the hyperosmotic condition. The larger increase in lactate content in the EDL vs. the soleus is most likely due to the different metabolic characteristics of each muscle type.
Despite the differences in intracellular lactate in the experimental conditions, there were no observable increases in extracellular lactate in any of the anisosmotic conditions. Two explanations are possible: 1) the extracellular fluid lactate was below the level of detection of our assay, or 2) there was minimal movement of lactate out of the muscle via the monocarboxylate transporters. The extracellular lactate may have been beyond detection because of the volume of media in the organ bath compared with the size of the incubating muscle. However, recent experiments in our laboratory have found increased lactate accumulation in the organ bath media despite the volume discrepancies during skeletal muscle stimulation (unpublished observations). Therefore, it seems that the lack of detection of lactate in the extracellular medium was due to a minimal release of lactate from the muscle rather than the volume-to-size ratio being a factor. Why lactate accumulated and was not extruded during the hyperosmotic condition requires further research, but it is tempting to speculate that cellular lactate may have been regulated in this condition, by both increased production and decreased extrusion.
Also, it is worthy to note that, to the authors' knowledge, the reported increases in lactate, at rest, due to the hyperosmotic medium have not been reported thus far in in vitro resting mammalian skeletal muscle. This increase in lactate observed in resting muscle as a result of the hyperosmotic condition may have also contributed to an alteration in intracellular hydrogen ion concentration (not measured); however, acid-base status has not been extensively studied in regard to cell volume (22, 25). Although in the present study the pH status was not measured, it may have been altered as a result of the increased lactate observed in the hyperosmotic condition.
In summary, this study examined the effect of altering extracellular osmolality on fluid shifts and metabolism in resting rat skeletal muscle. Acute changes in extracellular osmolality lead to muscle and fiber type-specific changes in cell volume and induced fluid shifts into or out of resting skeletal muscle as observed through wet to dry weights. EDL type II fiber area increased with hyposmotic and decreased with hyperosmotic conditions, whereas soleus type I and type II fiber area increased in the hyposmotic environment. Furthermore, exposure of type I and II resting skeletal muscle to a hyperosmotic environment resulted in a reduction in the energy charge of the muscle and an accumulation of muscle lactate. Further work is required to determine the specific mechanisms underlying these observations.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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