Isoform-specific regulation of the lactate transporters MCT1 and MCT4 by contractile activity

Arend Bonen1, Mio Tonouchi1, Dragana Miskovic1, Catherine Heddle3, John J. Heikkila2, and Andrew P. Halestrap3

Departments of 1 Kinesiology and 2 Biology, University of Waterloo, Waterloo, Ontario N2L 3G1, Canada; and 3 Department of Biochemistry, University of Bristol, Bristol BS8 1TD, United Kingdom


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

We examined the isoform-specific regulation of monocarboxylate transporter (MCT)1 and MCT4 expression by contractile activity in red and white tibialis anterior muscles. After 1 and 3 wk of chronic muscle stimulation (24 h/day), MCT1 protein expression was increased in the red muscles (+78%, P < 0.05). In the white muscles, MCT1 was increased after 1 wk (+191%) and then was decreased after 3 wk. In the red muscle, MCT1 mRNA accumulation was increased only after 3 wk (+21%; P < 0.05). In the white muscle, MCT1 mRNA was increased after 1 wk (+30%; P < 0.05) and 3 wk (+15%; P < 0.05). MCT4 protein was not altered in either the red or white muscles after 1 or 3 wk. MCT4 mRNA was transiently lowered (~15%) in both muscles in the 1st wk, but MCT4 mRNA levels were back to control levels after 3 wk. In conclusion, chronic contractile activity induces the expression of MCT1 but not MCT4. This increase in MCT1 alone was sufficient to increase lactate uptake from the circulation.

muscle fiber composition; lactate; perfusion; messenger ribonucleic acid


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

LACTIC ACID, the end-product of anaerobic glycolysis, is produced primarily by skeletal muscle. At physiological pH, lactic acid is almost completely dissociated because of its low pKa (3.86). Lactate can be oxidized in heart and an oxidative type of muscle fibers (36, 37), or it can be resynthesized to glycogen in fast-twitch muscle fibers (4, 5) or to glucose in liver and kidney (32). Thus there is a great flux of lactate out of and into a variety of cell types. There is now considerable evidence that lactate is transported across the plasma membrane of most cell types via a lactate proton cotransport system (for review see Refs. 22 and 43).

In the past few years, a family of monocarboxylate transporters (MCT) have been cloned. MCT1 is ubiquitously expressed in many tissues, including muscle and heart (2, 13, 18, 21, 31). MCT2 is expressed in hamster heart and muscle (12) but not in rat heart and muscle (19). MCT3 is expressed only in the retinal pigment epithelium (40, 49). More recently, an additional four MCTs have been identified (MCT4-MCT7, Ref. 44). Some of these MCTs are expressed in specific tissues; others are coexpressed in the same tissue (14, 44).

We have shown that MCT1 expression in muscle is highly correlated with the oxidative capacity of different types of muscles (r = 0.91), citrate synthase (r = 0.82), heart-type lactate dehydrogenase (H-LDH; r = 0.83), and with the rate of lactate uptake from the circulation (r = 0.90) (31). When the oxidative capacity of muscles is increased by chronic electrical stimulation (30) or exercise training (2), the expression of MCT1 is also increased.

We have recently reported that MCT1 and MCT4 are coexpressed in skeletal muscle (6, 47). Among all rat muscles, the slow-twitch oxidative soleus muscle expressed the least MCT4 and the most MCT1. Conversely, the fast-twitch glycolytic white tibialis anterior and white gastrocnemius muscles expressed the most MCT4, whereas MCT1 expression was very low. Fast-twitch oxidative glycolytic muscles, such as the red gastrocnemius and the red tibialis anterior, expressed large quantities of both MCT1 and MCT4 (6, 47). On the basis of these observations and others (2, 30, 31), it has been proposed that the expression of MCT1 is most closely associated with muscle characteristics favoring the uptake of lactate for oxidative disposal (30, 31), whereas MCT4 expression is related to the need for lactate extrusion (6, 47). Therefore, it is likely that the expression of MCT1 and MCT4 is regulated independently in skeletal muscle.

Low-frequency chronic electrical stimulation has been used to examine the effects of increased contractile activity on the expression of a number of proteins in muscle (25, 39), including transport proteins (3, 20, 23, 30). In this model of chronic muscle stimulation, the electrical impulses imposed on the motor nerve are designed to resemble those observed in oxidative types of muscles in vivo (15, 39). This type of stimulation rapidly increases the oxidative capacity of muscle in <7 days (29, 39). Moreover, given sufficient time, low-frequency chronic electrical stimulation can also transform fast-twitch fatigable muscle fibers into slower-twitching fatigue-resistant fibers (39). We have shown that it is also possible to stimulate simultaneously, via the peroneal nerve, both the red and the white muscle fiber compartments of the tibialis anterior (TA) muscle in the rat (20, 30). Thus this model of controlled, electrical pacing for varying periods of time provides a means to examine the contraction-regulated expression of proteins in red and white muscles, either in the absence or in the presence of changes in the metabolic phenotype of the muscles.

In the present studies, we have used chronic low-frequency electrical stimulation to examine the effects of increased contractile activity on the expression of MCT1 and MCT4 proteins and on the accumulation of MCT1 mRNA and MCT4 mRNA in the highly oxidative red tibialis anterior (RTA) and the highly glycolytic white tibialis anterior (WTA) muscles.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Animals. Male Sprague-Dawley rats weighing 280-350 g were used in all experiments. Animals were housed in an air-conditioned room on a 12:12-h light-dark cycle and were fed a diet of Purina Chow and water ad libitum. Ethical approval was obtained for this work from the Committee on Animal Care at the University of Waterloo.

Chronic electrical stimulation of rat muscles. Muscles were stimulated as previously described (20, 29, 30). Briefly, in anesthetized rats, stainless steel electrodes were sutured to underlying muscles on either side of the peroneal nerve. These electrodes were passed subcutaneously from the thigh and exteriorized at the back of the neck, where they were attached to a miniature electronic stimulator. After a wait of >= 5 days to allow animals to recover, stimulus pulses (10 Hz, 50 µs duration) were initiated. The peroneal nerve, which innervates the RTA and WTA muscles, was stimulated for 24 h/day for 7 or 21 days. In further experiments, muscles were stimulated for 0, 0.25, 1, 3, 6, or 24 h/day for 7 days. At the end of the stimulation periods, animals were euthanized, and muscles were separated into RTA and WTA compartments, as we have done previously (20, 30). Muscles were rapidly frozen and stored at -80°C until analyzed for MCT1 and MCT4 proteins and MCT1 and MCT4 mRNAs. We also determined the muscle fiber composition of control and stimulated muscles after 1 and 3 wk of chronic stimulation in separate groups of animals. Lactate uptake from the circulation by control and stimulated muscles was examined in the 1-wk-stimulated group only by use of the perfused rat hindlimb preparation (30).

Determinations of MCT1 and MCT4 proteins. Western blotting was used to detect MCT1 and MCT4 proteins by an enhanced chemiluminescence detection system (2-min exposures). These procedures have been described in detail in our previous studies (2, 6, 30, 31, 47). Densities of the protein bands were quantified by scanning the resultant films on a densitometer connected to a Macintosh LC computer with appropriate software.

Determinations of MCT1 mRNA and MCT4 mRNA. MCT1 mRNA and MCT4 mRNA were determined using Northern blotting procedures, as we have described previously (6). Briefly, total RNA was isolated from heart and muscle tissues using the guanidinium isothiocyanate/cesium chloride (GIT/CsCl) centrifugation method (11), with some modifications. The tissues were homogenized in 10 ml of 4 M GIT and layered on top of 3.3 ml of 5.7 M CsCl solution. The samples were centrifuged in an SW-41 Ti rotor (Beckman) at 154,000 g for 23 h. The RNA pellets were recovered and were purified by two precipitations in ethanol.

Northern blot analysis. Three micrograms of total RNA were used for electrophoresis on 1.2% formaldehyde agarose gels (45) and then transferred to positively charged nylon membrane (Boehringer Mannheim, Laval, QC, Canada). The Northern blots were ultraviolet cross-linked with a GS-Gene linker (Bio-Rad). Equal loading was confirmed with the 18S rRNA signal.

A 1.9-kb fragment containing the coding sequence of MCT1 cDNA was isolated from the full length (3.3-kb) MCT1 by digestion with EcoR I restriction enzyme from the full length (3.3-kb) MCT1 cDNA (18) and was subcloned into the EcoR I restriction enzyme site of pBluescript (KS). The orientation was checked by digestion with Hind III restriction endonuclease. Template DNA was linearized with Xba I restriction enzyme, and digoxigenin-11-UTP (DIG)-labeled MCT1 antisense riboprobe was generated by in vitro transcription with T3 RNA polymerase. MCT4 cDNA was originally subcloned into BamH I/Apa I restriction enzyme sites of pBluescript (47). DIG-labeled MCT4 antisense riboprobe was generated by digestion of the template DNA with Xba I restriction enzyme, and in vitro transcription was generated with T7 RNA polymerase.

The ingredients for RNA transcription included 1-2 µg of DNA template plus the NTP mix [2.5 mM CTP, 2.5 mM rGTP, 2.5 mM ATP, 1.625 mM UTP (Promega), and 0.875 mM DIG-11 UTP (Boehringer Mannheim)], 20 mM dithiothreitol (DTT; Promega), 1 U/1 µg template DNA of RNase inhibitor (Promega) and 1× RNA polymerase buffer [5× buffer: 400 mM Tris · HCl, pH 7.5; 60 mM MgCl2; and 20 mM spermidine-HCl (Promega)] maintained at room temperature. The appropriate RNA polymerase [T3 or T7 RNA polymerases (Boehringer Mannheim)] was added (>= 20 IU/1 µg of DNA template) and incubated for 2 h at 37°C. The DNA template was then digested for 10 min at 37°C with RNase-free DNase (1 IU/1 µg of DNA template; Promega). After precipitation in ethanol and centrifugation at 13,500 g for 15 min, the probe was resuspended in 10-20 ml DIG easy-hyb hybridization buffer (Boehringer Mannheim) or standard hybridization buffer with 50% formamide [5× SSC, 50% formamide, 0.1% sodium-lauroylsarcosine, 0.02% SDS, and 2% blocking reagent (Boehringer Mannheim)].

After prehybridization of the membrane for >= 4 h at 68°C, the prehybridization buffer was replaced with the same buffer containing a DIG-labeled antisense RNA probe, and the membrane was incubated with the probe overnight at 68°C. High-stringency washes and chemiluminescent detection were performed in accordance with the protocol supplied by the manufacturer (Boehringer Mannheim), and the membrane was exposed to Kodak BioMax film. After a 3-min exposure, the film was developed in Kodak developer and fixed in Kodak fixer.

Muscle fiber composition. Stimulated and control muscles were histochemically analyzed for fiber composition, as previously described (38). For this purpose a small portion of fresh excised muscle was embedded in Tissue-Tek OCT compound (Miles, Westhaven, CT) with isopentane-cooled tongs and was frozen in liquid nitrogen. Serial cross-sections (10 µm) were stained for myofibrillar ATPase (at pH 10.3), NADH diaphorase, and alpha -glycerophosphate dehydrogenase. Muscle fibers were classified according to the method of Peter et al. (38). Muscle fibers that stained high for NADH diaphorase and alpha -glycerophosphate dehydrogenase and intermediate-high for myofibrillar ATPase were designated as fast-twitch oxidative glycolytic fibers (FOG); muscle fibers that stained low for NADH diaphorase and high for both alpha -glycerophosphate dehydrogenase and myofibrillar ATPase were designated as fast-twitch glycolytic fibers (FG); muscle fibers that stained intermediate for NADH diaphorase and low for both alpha -glycerophosphate dehydrogenase and myofibrillar ATPase were designated as slow-twitch oxidative fibers (SO) (38).

Lactate uptake in perfused rat hindlimb muscles. Short-duration (5-min) hindlimb perfusion of rat muscles, designed to limit lactate oxidation to <10% of lactate taken up (30, 31), was used to determine lactate uptake from the circulation by control and chronically stimulated muscles. For these purposes we used a cell-free, gassed (95% O2-5% CO2) Krebs-Henseleit buffer containing 4% bovine serum albumin, pH 7.4, 10 mM glucose, and lactate (2 mM, 3 µCi [U-14C]lactate, 1 µCi [3H]sorbitol). A one-pass system was used, and therefore the venous outflow was discarded. Immediately after 5 min of perfusion, muscles were rapidly extracted and frozen in liquid nitrogen. These perfusion procedures have previously been described in detail in some of our recent studies in which we examined short-term (5-min) lactate uptake by perfused muscles (6, 30, 31).

Data analyses. Data analyses involved comparing paired muscles from the same animal (control vs. experimental muscle) during weeks 1 and 3. Significance was accepted at P < 0.05. All results are means ± SE. For graphing purposes only, the data from all the control muscles have been pooled.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

At the onset of the chronic stimulation period, after a 5-day recovery from surgery, all animals had surpassed their presurgery body weights (data not shown). Throughout the 1 and 3 wk of chronic muscle contraction, the animals continued to gain weight (data not shown).

Effects of chronic stimulation on muscle fiber composition. After 1 wk of chronic stimulation, no changes in muscle fiber composition were observed, except for a small change in the RTA. In this muscle, the FOG content was increased from 60 ± 2 to 75 ± 3% (P < 0.05), while concomitantly the FG fiber content was reduced from 26 ± 2 to 16 ± 3% (P < 0.05) (Fig. 1). The fiber composition of the WTA was not altered during this 7-day period (Fig. 1).


View larger version (21K):
[in this window]
[in a new window]
 
Fig. 1.   Muscle fiber composition of red (RTA, A) and white tibialis anterior (WTA, B) muscles after 1 and 3 wk of chronic stimulation; n = 6 and 5 muscles in each of weeks 1 and 3, respectively, control = 12 muscles. SO, slow-twitch oxidative fibers; FG, fast-twitch glycolytic fibers; FOG, fast-twitch oxidative glycolytic fibers. RTA: %FOG, control < week 1 week 3 (P < 0.05); %FG, week 1 > week 3 (P < 0.05); WTA: %FOG, week 1 week 3 (P < 0.05); %FG, week 1 > week 3 (P < 0.05).

In contrast, after 3 wk of chronic stimulation, the percentages of FG and FOG fibers were altered in both the RTA and WTA. In the RTA, the FG fibers were almost completely transformed into FOG fibers (Fig. 1). Similarly, in the 3-wk chronically stimulated WTA, the muscle was transformed from one consisting of 81 ± 1% FG fibers into one composed of 95 ± 1% FOG fibers (Fig. 1).

Effects of chronic stimulation on MCT1 mRNA and protein. In control muscles, the relative level of MCT1 mRNA in the RTA was 60% higher than in the WTA (P < 0.05, Fig. 2A). Chronic muscle stimulation increased MCT1 mRNA levels in both muscles (P < 0.05). However, there were differences in the pattern of MCT1 mRNA accumulations in the red and white muscles. In the RTA, 1 wk of chronic stimulation did not alter MCT1 mRNA (P > 0.05), whereas in the WTA there was a 30% increase in MCT1 mRNA after 1 wk of chronic stimulation. After 3 wk of stimulation, MCT1 mRNA levels in the RTA were increased by 21% (P < 0.05) and by 15% in the WTA (P < 0.05; Fig. 2A).


View larger version (27K):
[in this window]
[in a new window]
 
Fig. 2.   Monocarboxylate transporter 1 (MCT1) mRNA (A) and MCT4 mRNA (B) in control and 1- and 3-wk chronically stimulated RTA and WTA muscles; n = 6 muscles in control and 3 muscles in 1- and 3-wk stimulated muscles. MCT1 and MCT4 mRNA is expressed in arbitrary optical density (OD) units. aP < 0.05 control vs. 1 wk stimulated, control vs. 3 wk stimulated; bP < 0.05 control WTA vs. control RTA, 1-wk stimulated WTA vs. 1-wk stimulated RTA, 3-wk stimulated WTA vs. 3-wk stimulated RTA.

In the control RTA, MCT1 protein concentrations were 2.3 times greater than in the control WTA (P < 0.05; Fig. 3A). After 1 wk of chronic stimulation, the MCT1 protein content in the RTA had increased to 178% of the contralateral control RTA (100%) (P < 0.05; Fig. 3A), and the MCT1 protein in the WTA content had increased to 291% of the control WTA (100%) (P < 0.05; Fig. 3A).


View larger version (25K):
[in this window]
[in a new window]
 
Fig. 3.   MCT1 (A) and MCT4 (B) proteins in control and 1- and 3-wk chronically stimulated RTA and WTA muscles; n = 10 muscles in control and 4-6 muscles in 1- and 3-wk stimulated muscles. aP < 0.05 control vs. 1-wk stimulated, control vs. 3-wk stimulated; bP < 0.05 control WTA vs. control RTA; cP < 0.05 3-wk stimulated WTA vs. 1-wk stimulated WTA.

Three weeks of chronic stimulation did not increase MCT1 protein further in the RTA (P > 0.05). In contrast, there was a decrease in MCT1 in the WTA between weeks 1 and 3 of stimulation (P < 0.05; Fig. 3A). But even then, the MCT1 protein was still 50% greater than in the control WTA (P < 0.05; Fig. 3A).

The increase in MCT1 protein in both muscles was dependent on the amount of contractile activity imposed on the muscle during a 7-day period (Fig. 4). Maximal increases in MCT1 occurred after 3 h/day of chronic stimulation for 7 days, with no further increase when the stimulation time was extended to 24 h/day.


View larger version (16K):
[in this window]
[in a new window]
 
Fig. 4.   Effects of varied amounts of chronic stimulation on MCT1 in RTA and WTA muscles. Muscles were stimulated for 0, 0.25, 1, 3, 6, and 24 h/day for 7 days; n = 3-6 muscles at each time point, except at t = 0, where n = 20.

Effects of chronic stimulation on MCT4 mRNA and protein. The abundance of MCT4 mRNA in the control red and white muscles did not differ (P > 0.05, Fig. 2B). After 1 wk of chronic stimulation, there was a small decrease (~15%) in MCT4 mRNA in the two muscles (P < 0.05; Fig. 2B). This reduction did not persist, because MCT4 RNA had returned to control levels after 3 wk of chronic contraction in both muscles (Fig. 2B).

Control RTA and control WTA expressed similar quantities of MCT4 protein (Fig. 3B; P > 0.05). After 1 and 3 wk of chronically increased muscle contractility, there were no changes in MCT4 protein content in either the RTA or WTA (P > 0.05; Fig. 3B).

Comparison of MCT protein and mRNA accumulation. The relative amounts of MCT protein and mRNA were compared in control and stimulated muscle. The ratio of MCT4 protein to MCT4 mRNA was similar in the control and the 1- and 3-wk-stimulated muscles (Table 1). On the other hand, the ratio of MCT1 protein to MCT1 mRNA increased sharply after 1 and 3 wk of chronic stimulation in the RTA, and after 1 wk in the WTA (Table 1). This occurred whether MCT1 mRNA accumulation was increased (i.e., RTA week 3, WTA weeks 1 and 3) or remained unchanged (i.e., RTA week 1). The decrease in the ratio of MCT1 protein to MCT1 mRNA in the 3-wk-stimulated WTA was due to reductions in the MCT1 protein, because the MCT1 mRNA levels in week 3 were similar to the level observed in week 1 (Fig. 2A).

                              
View this table:
[in this window]
[in a new window]
 
Table 1.   Ratio of MCT protein to MCT mRNA in control and 1- and 3-wk chronically stimulated muscles

Effects of chronic muscle stimulation on lactate uptake by perfused muscles. Lactate uptake from the circulation by perfused muscles was determined only in control and 1-wk chronically stimulated RTA and WTA, because this was when the optimal changes in MCT1 were observed in both red and white muscles. In the chronically stimulated RTA, lactate uptake was increased by +39% (P < 0.05), and in the chronically stimulated WTA it was increased by +120% (P < 0.05, Table 2).

                              
View this table:
[in this window]
[in a new window]
 
Table 2.   Lactate uptake by control and 1-wk chronically stimulated muscles


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

In this report, we demonstrate that muscle contractile activity regulates the expression of MCT1 and MCT4 in an isoform-specific manner. MCT1 protein was increased +78 and +191% in contracting red and white muscles, respectively, at 1 wk. Concomitantly, there were either no changes in MCT1 mRNA or only very modest accumulations in MCT1 mRNA (<= 30%). In contrast to MCT1, low-frequency chronic muscle stimulation activity did not affect the expression of MCT4 in either red or white muscles. Only a slight transient reduction (-15%) in MCT4 mRNA occurred in the red and white muscles after 1 wk of stimulation. Interestingly, the isoform-specific regulation of the MCT1 and MCT4 occurred independently of changes in muscle fiber composition. The contraction-induced increase in MCT1 alone was sufficient to increase lactate uptake from the circulation.

The 60% difference in MCT1 mRNA between control RTA and control WTA muscles is considerably less than the large difference in MCT1 protein content in these muscles (RTA 2.3 times greater than WTA). Such differences in the MCT1 mRNA and MCT1 protein have been observed previously (6, 30, 31, 47). Thus our current work suggests that skeletal muscle MCT1 protein expression is regulated largely by posttranscriptional mechanisms. Because motor unit firing patterns in red muscle are much greater than in white muscle (15, 33), it may well be that that this neural innervation pattern serves to regulate MCT1 expression via posttranscriptional mechanisms.

In our experimental studies, it is seen that a new neural innervation pattern, imposed onto the muscle by chronic electrical stimulation, upregulates MCT1. This appears to occur largely via posttranscriptional mechanisms. For example, in chronically stimulated white muscle, there was only a small increase in MCT1 mRNA (+30%), whereas MCT1 protein concentrations were increased far more (+191%). And similarly, in stimulated red TA muscle, MCT1 protein was increased (+78%) without any changes in MCT1 mRNA. Others have shown a stable level of MCT1 protein while MCT1 mRNA was being reduced in the developing rat brain (26). It has also been observed that the MCT1 protein-to-MCT1 mRNA ratios differed greatly among different tissues. For example, much lower levels of MCT1 mRNA are present in rat testes than in rat heart, yet both express the same relative amount of protein (19). Thus our studies and others (19, 26) indicate that posttranscriptional mechanisms regulate the expression of MCT1 in muscle and a number of other tissues. Similarly, posttranscriptional mechanisms also appear to regulate the expression of MCT2 (19). It has been proposed that pools of MCT1 mRNA may be in messenger ribonucleoprotein, thus facilitating rapid translation when more protein is required. The very long 3' untranslated region of MCT1 (1.6 kb) (13, 18) may be involved in the translational repression. With chronic muscle stimulation, translational capacity and efficiency are enhanced, as evidenced by the early (2-3 days) increase (~2-fold) in monosomes, polysomes, and ribosomal RNA (1, 24, 46).

The lack of any changes in MCT4 after either 1 or 3 wk of muscle contraction is in marked contrast to the responses of MCT1 in the same muscles. Yet, it is clear that MCT4 is abundantly expressed in skeletal muscle (present study and Ref. 47). Our data indicate that MCT4 is expressed in both FG and FOG types of skeletal muscle fibers. Only in slow-twitch muscles (soleus) are MCT4 protein and mRNA very low (6). Presumably, MCT4 is expressed only in the few FOG fibers that are present in this muscle. Recently, it was shown that MCT4 protein was reduced when muscle activity was completely eliminated by denervation (47). These observations, along with those in the present study, suggest that at least some level of activity is required to maintain MCT4 levels in muscle, but that chronically increasing muscle activity further is not necessary to maintain these MCT4 levels. In contrast, MCT1 is seen to be very sensitive to changes in muscle activity. With denervation this MCT1 protein content is also reduced (47); however, when muscle activity was increased (present study and Refs. 2 and 30), MCT1 was increased. Indeed, we have shown a good relationship between the quantity of muscle contractile activity and MCT1, although maximal increases in MCT1 are evident with 3 h of chronic stimulation for 7 days (see Fig. 4).

We have previously observed that chronic muscle stimulation increased lactate flux by muscles (29, 30). We proposed that this could be accounted for by an increase in MCT1 (30), but this explanation was undermined by the discovery that MCT4 was also expressed in muscle (47). However, the present studies demonstrate clearly that the increase in lactate uptake in chronically stimulated muscle is attributable to changes in MCT1 alone, because MCT4 was not altered. This is consistent with our previous observations that MCT1 content in muscles appears to be be highly correlated with the uptake of lactate from the circulation (r = 0.90) (30, 31).

A number of studies in recent years have shown that increased muscle contractile activity induced by chronic muscle stimulation regulates the expression of the glucose transporter GLUT-4 (17, 20, 23, 48) and the fatty acid transporter fatty acid translocase (3). All of these transporters, as well as MCT1, are highly correlated with the oxidative capacity of different muscles (20, 30, 31, 34, 35). This facilitates a greater rate of uptake of glucose, fatty acid, and lactate by oxidative muscles (20, 30, 31, 34, 35), where these substrates can be readily oxidized in these muscles (3, 7, 37). To facilitate the oxidative disposal of lactate, MCT1 proteins are perhaps concentrated at the surface of the muscle clustered near mitochondria. In heart, we observed that the most dense MCT1 labeling occurred in T tubules that were in close proximity to mitochondria (21). More recently, it has been shown that MCT1 is located on the plasma membrane (6, 8) as well as on subsarcolemmal and intermyofibrillar mitochondria (8). These locations would seem consistent with the idea that MCT1 is most closely associated with the removal of lactate from the circulation, as well as oxidation of lactate produced by other muscle cells.

Several lines of evidence suggest that the expression of MCT4 is related to the need to extrude lactate from muscle cells. In the present study and in a previous report (47), we showed that MCT4 expression is largely confined to FOG and FG fibers, which are each capable of producing copious quantities of lactate. Highly oxidative tissues such as the rat heart do not express MCT4, and the highly oxidative soleus muscle, which is comprised almost entirely of slow-twitch oxidative fibers, expressed very little MCT4 (47). It also appears that MCT4 is not associated with the subsarcolemmal and intermyofibrillar mitochondria (8). Also, we have shown that there is an intracellular pool of MCT4 (6), which may perhaps be translocated to the plasma membrane to assist with the removal of lactate when lactate concentrations are very high (e.g., intense exercise). Collectively, these data link MCT4 expression to the capacity for anaerobic glycolysis in muscle tissue, from which it is necessary to extrude lactate.

It is evident that MCT1 expression attains a maximum after a critical amount of contractile activity has been attained. Stimulation of 3 h/day for 7 days was sufficient to induce maximal MCT1 increments in both RTA (+89%) and WTA (+115%). Extending the stimulation period up to 24 h/day for 7 days did not produce a further increase. In the red muscle, stimulations of 24 h/day for 21 days also failed to increase MCT1 further. However, in the white muscle, this prolonged 21-day stimulation period reduced MCT1 protein somewhat from that observed after 7 days. It is not clear why this occurred, except to note that the activity of some enzymes is also reduced after reaching a peak during many weeks of chronic stimulation of skeletal muscles (10, 16, 28). This phenomenon appears to be a function of the stimulation frequency, because this decline in enzyme activity occurs at stimulations of 10 but not 2.5 Hz (28). Because there is an extensive remodeling of the muscle fiber composition between weeks 1 and 3 in red and white muscles at 10 Hz (present study and Ref. 28) but not at 2.5 Hz (28), it has been proposed that this muscle remodeling may interfere, in an unknown manner, with maintaining the levels of some enzymes (9, 28) and perhaps a transport protein such as MCT1. This explanation is supported by the fact that the decrease in MCT1 in our studies occurred only in the white muscle, in which the muscle fiber changes were far greater than in the red muscle.

Both MCT1 and MCT4 have been observed in type I and type II fibers in human muscle (42). MCT4 is more predominant in type II fibers than in the type I fibers. MCT1 was comparable in both of these muscle fiber types (42). After 8 wk of intense training, both MCT isoforms were increased (MCT1 +76%; MCT4 +32%) (41). However, this training model is not directly comparable to the chronic stimulation model in the present studies. First, the amount of contractile activity in chronically stimulated muscles (128 h in 7 days) was far greater than the amount of contractile activity encountered by training in humans in the study by Pilegaard et al. (41) (calculated to be a total of 10 h over a 56-day period). In addition, the quality of the contractile activity in the present study and that of Pilegaard et al. also differed greatly. With chronic stimulation, the contractile activity pattern is designed to be similar to that in a slow-twitch muscle (39), whereas in the training study of Pilegaard et al., the training procedure (1-legged extensions) was undoubtedly in excess of the maximal aerobic capacity of the muscle performing the exercise. The high lactate concentrations in the plasma in that study (41) suggest that the training was anaerobic in nature, whereas that was not the case with chronic contractile stimulation. It may well be that intense anaerobic exercise can increase both MCT1 and MCT4 proteins in muscle, as was observed by Pilegaard et al., whereas aerobic training increases only MCT1 protein, as we observed.

In summary, these studies have shown that chronically increased muscle activity fails to alter the MCT4 protein or mRNA in either red or white muscle, whereas the content of MCT1 is greatly increased in both types of muscles in either the presence or the absence of muscle fiber remodeling. It appears that MCT1 expression is regulated primarily by posttranscriptional mechanisms, because there were substantial increases in protein levels but only modest changes in mRNA levels. The overexpression of the MCT1 protein alone is sufficient to increase lactate uptake from the circulation.


    ACKNOWLEDGEMENTS

These studies were funded by grants to A. Bonen from the the Heart and Stroke Foundation of Ontario and the Natural Sciences and Engineering Research Council of Canada, and to A. P. Halestrap by the British Heart Foundation, The Wellcome Trust, and the Medical Research Council (UK).


    FOOTNOTES

Address for reprint requests and other correspondence: A. Bonen, Dept. of Kinesiology, Univ. of Waterloo, Waterloo, Ontario N2L 3G1, Canada (E-mail: abonen{at}healthy.uwaterloo.ca).

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.

Received 11 January 2000; accepted in final form 6 July 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Annex, BH, Kraus WE, Dohm GL, and Williams RS. Mitochondrial biogenesis in striated muscles: rapid induction of citrate synthase mRNA by nerve stimulation. Am J Physiol Cell Physiol 260: C266-C270, 1991[Abstract/Free Full Text].

2.   Baker, SK, McCullagh KJA, and Bonen A. Training intensity-dependent and tissue-specific increases in lactate uptake and MCT-1 in heart and muscle. J Appl Physiol 84: 987-994, 1998[Abstract/Free Full Text].

3.   Bonen, A, Dyck DJ, Ibrahimi A, and Abumrad NA. Muscle contractile activity increases fatty acid metabolism and transport and FAT/CD36. Am J Physiol Endocrinol Metab 276: E642-E649, 1999[Abstract/Free Full Text].

4.   Bonen, A, and Homonko D. Effects of exercise and glycogen depletion on glyconeogenesis in muscle. J Appl Physiol 76: 1753-1758, 1994[Abstract/Free Full Text].

5.   Bonen, A, McDermott JC, and Tan MH. Glycogenesis and glyconeogenesis in skeletal muscle: effects of pH and hormones. Am J Physiol Endocrinol Metab 258: E693-E700, 1990[Abstract/Free Full Text].

6.   Bonen, A, Miskovic D, Tonouchi M, Lemieux K, Wilson MC, Marette A, and Halestrap AP. Abundance and subcellular distribution of MCT1 and MCT4 in heart and fast-twitch skeletal muscles. Am J Physiol Endocrinol Metab 278: E1067-E1077, 2000[Abstract/Free Full Text].

7.   Bonen, A, Tan MH, and Watson-Wright W. Effects of exercise on insulin binding and glucose metabolism in muscle. Can J Physiol Pharmacol 62: 1500-1504, 1984[ISI][Medline].

8.   Brooks, GA, Brown MA, Butz CE, Sicurello JP, and Dubouchaud H. Cardiac and skeletal muscle mitochondria have a monocarboxylate transporter MCT1. J Appl Physiol 87: 1713-1718, 1999[Abstract/Free Full Text].

9.   Brown, WE, Salmmons S, and Whalen RG. The sequential replacement of myosin subunit isoforms during muscle type transformation induced by long term electrical stimulation. J Biol Chem 258: 14686-14692, 1983[Abstract/Free Full Text].

10.   Chi, MM, Hintz CS, Henriksson J, Salmons S, Hellendahl RP, Park JL, Nemeth PM, and Lowry OH. Chronic stimulation of mammalian muscle: enzyme changes in individual fibers. Am J Physiol Cell Physiol 251: C633-C642, 1986[Abstract/Free Full Text].

11.   Chirgwin, J, Przbyla A, MacDonald R, and Rutter W. Isolation of biologically active ribonucleic acid from source enriched in ribonuclease. Biochemistry 18: 5294-5299, 1979[ISI][Medline].

12.   Garcia, CK, Brown MS, Pathak RK, and Goldstein JL. cDNA cloning of MCT2, a second monocarboxylate transporter expressed in different cells than MCT1. J Biol Chem 270: 1843-1849, 1995[Abstract/Free Full Text].

13.   Garcia, CK, Goldstein JL, Pathak RK, Anderson GW, and Brown MS. Molecular characterization of a membrane transporter for lactate, pyruvate, and other monocarboxylates: implications for the Cori cycle. Cell 76: 865-873, 1994[ISI][Medline].

14.   Halestrap, AP, and Price NT. The proton-linked monocarboxylate transporter family: structure, function and regulation. Biochem J 343: 281-299, 1999[ISI][Medline].

15.   Hennig, R, and Lomo T. Firing patterns of motor units in normal rats. Nature 314: 164-166, 1985[ISI][Medline].

16.   Henriksson, JM, Chi MY, Hintz S, Young DA, Kaiser KK, Salmons S, and Lowry OH. Chronic stimulation of mammalian muscle: changes in enzymes of six metabolic pathways. Am J Physiol Cell Physiol 251: C614-C632, 1986[Abstract/Free Full Text].

17.   Hofmann, S, and Pette D. Low frequency stimulation of rat fast-twitch muscle enhances the expression of hexokinase II and both the translocation and expression of glucose transporter 4 (GLUT-4). Eur J Biochem 219: 307-315, 1994[Abstract].

18.   Jackson, VN, Price NT, and Halestrap AP. cDNA cloning of MCT1, a monocarboxylate transporter from rat skeletal muscle. Biochim Biophys Acta 1238: 193-196, 1995[ISI][Medline].

19.   Jackson, VN, Price NT, and Halestrap AP. Cloning of the monocarboxylate transporter isoform MCT2 from rat testis provides evidence that the expression is species specific and may involve posttranscriptional regulation. Biochem J 324: 447-453, 1997[ISI][Medline].

20.   Johannsson, E, McCullagh KJA, Han XX, Fernando PK, Jensen J, Dahl HA, and Bonen A. Effect of overexpressing GLUT-1 and GLUT-4 on insulin- and contraction-stimulated glucose transport in muscle. Am J Physiol Endocrinol Metab 271: E547-E555, 1996[Abstract/Free Full Text].

21.   Johannsson, E, Nagelhus EA, McCullagh KJA, Sejersted OM, Blackstad TW, Bonen A, and Ottersen OP. Cellular and subcellular expression of the monocarboxylate transporter MCT1 in rat heart. A high resolution immunogold analysis. Circ Res 80: 400-407, 1997[ISI][Medline].

22.   Juel, C. Lactate-proton cotransport in skeletal muscle. Physiol Rev 77: 1-37, 1997[Abstract/Free Full Text].

23.   Kong, X, Manchester J, Salmons S, and Lawrence JC. Glucose transporters in single skeletal muscle fibres. J Biol Chem 269: 12963-12967, 1994[Abstract/Free Full Text].

24.   Kraus, WE, Bernard TS, and Williams RS. Interactions between sustained contractile activity and beta -adrenergic receptors in regulation of gene expression in skeletal muscles. Am J Physiol Cell Physiol 256: C506-C514, 1989[Abstract/Free Full Text].

25.   Kraus, WE, Torgan CE, and Taylor DA. Skeletal muscle adaptation to chronic low frequency stimulation. In: Reviews in Exercise and Sport Sciences, edited by Holloszy JO., 1994, p. 313-360.

26.   Leino, RL, Gerhart DZ, and Drewes LR. Monocarboxylate transporter (MCT1) abundance in brains of suckling and adult rats: a quantitative electron microscopic immunogold study. Dev Brain Res 113: 47-54, 1999[ISI][Medline].

27.   Luiken, JJFP, Turcotte LP, and Bonen A. Protein-mediated palmitate uptake and expression of fatty acid transport proteins in heart giant vesicles. J Lipid Res 40: 1007-1016, 1999[Abstract/Free Full Text].

28.   Mayne, CN, Sutherland H, Jarvis JC, Gilroy SJ, Craven AJ, and Salmons S. Induction of a fast-oxidative phenotype by chronic muscle stimulation: histochemical and metabolic studies. Am J Physiol Cell Physiol 270: C313-C320, 1996[Abstract/Free Full Text].

29.   McCullagh, KJA, Juel C, O'Brien M, and Bonen A. Chronic muscle stimulation increases lactate transport in rat skeletal muscle. Mol Cell Biochem 156: 51-57, 1996[ISI][Medline].

30.   McCullagh, KJA, Poole RC, Halestrap AP, Tipton KF, O'Brien M, and Bonen A. Chronic electrical stimulation increases MCT1 and lactate uptake in red and white skeletal muscle. Am J Physiol Endocrinol Metab 273: E239-E246, 1997[Abstract/Free Full Text].

31.   McCullagh, KJA, Poole RC, Halestrap AP, O'Brien M, and Bonen A. Role of the lactate transporter (MCT1) in skeletal muscles. Am J Physiol Endocrinol Metab 271: E143-E150, 1996[Abstract/Free Full Text].

32.   McDermott, JC, and Bonen A. Glyconeogenic and oxidative lactate utilization in skeletal muscle. Can J Physiol Pharmacol 70: 142-149, 1992[ISI][Medline].

33.   McDermott, JC, Elder GCB, and Bonen A. Non-exercising muscle metabolism during exercise. Pflügers Arch 418: 301-307, 1991[ISI][Medline].

34.   Megeney, LA, Michel RN, Boudreau CS, Fernando PK, Prasad M, Tan MH, and Bonen A. Regulation of muscle glucose transport and GLUT-4 by nerve-derived factors and activity-related processes. Am J Physiol Regulatory Integrative Comp Physiol 269: R1148-R1153, 1995[Abstract/Free Full Text].

35.   Megeney, LA, Neufer PD, Dohm GL, Tan MH, Blewett CA, Elder GCB, and Bonen A. Effects of muscle activity and fiber composition on glucose transport and GLUT-4. Am J Physiol Endocrinol Metab 264: E583-E593, 1993[Abstract/Free Full Text].

36.   Pagliassotti, MJ, and Donovan CJ. Influence of cell heterogeneity on skeletal muscle lactate kinetics. Am J Physiol Endocrinol Metab 258: E625-E634, 1990[Abstract/Free Full Text].

37.   Pagliassotti, MJ, and Donovan CJ. Role of cell type in net lactate removal by skeletal muscle. Am J Physiol Endocrinol Metab 258: E635-E642, 1990[Abstract/Free Full Text].

38.   Peter, JB, Barnard RJ, Edgerton VR, Gillespie CA, and Stempel KE. Metabolic profiles of three fibre types of skeletal muscle in guinea pigs and rabbits. Biochemistry 11: 2627-2633, 1972[ISI][Medline].

39.   Pette, D, and Vrbova G. Adaptation of skeletal muscle fibers to chronic electrical stimulation. Rev Physiol Biochem Pharmacol 120: 116-202, 1992.

40.   Philp, NJ, Yoon H, and Grollman EF. Monocarboxylate transporter MCT1 is located in the apical membrane and MCT3 in the basal membrane of rat RPE. Am J Physiol Regulatory Integrative Comp Physiol 274: R1824-R1828, 1998[Abstract/Free Full Text].

41.   Pilegaard, H, Domino K, Noland T, Juel C, Hellsten Y, Halestrap AP, and Bangsbo J. Effect of high-intensity exercise training on lactate/H+ transport capacity in human skeletal muscle. Am J Physiol Endocrinol Metab 276: E255-E261, 1999[Abstract/Free Full Text].

42.   Pilegaard, H, Terzis G, Halestrap A, and Juel C. Distribution of the lactate/H+ transporter isoforms MCT1 and MCT4 in human skeletal muscle. Am J Physiol Endocrinol Metab 276: E843-E848, 1999[Abstract/Free Full Text].

43.   Poole, RC, and Halestrap AP. Transport of lactate and other monocarboxylates across mammalian plasma membranes. Am J Physiol Cell Physiol 264: C761-C782, 1993[Abstract/Free Full Text].

44.   Price, NT, Jackson VN, and Halestrap AP. Cloning and sequencing of four new mammalian monocarboxylate transporter (MCT) homologues confirms the existence of a transporter family with an ancient past. Biochem J 329: 321-328, 1998[ISI][Medline].

45.   Sambrook, J, Fritisch EF, and Maniatis T. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory, 1989.

46.   Seedorf, U, Leberer E, Kirschbaum BJ, and Pette D. Neural control of gene expression in muscle. Effects of chronic stimulation on lactate dehydrogenase isozymes and citrate synthase. Biochem J 239: 115-120, 1986[ISI][Medline].

47.   Wilson, MC, Jackson VN, Hedle C, Price NT, Pilegaard H, Juel C, Bonen A, Montgomery I, Hutter OF, and Halestrap AP. Lactic acid efflux from white skeletal muscle is catalyzed by the monocarboxylate transporter MCT3. J Biol Chem 273: 15920-15926, 1998[Abstract/Free Full Text].

48.   Yaskelpis, BB, III, Castle AL, Farrar RP, and Ivy JL. Contraction-induced intracellular signals and their relationship to muscle GLUT-4 concentration. Am J Physiol Endocrinol Metab 272: E118-E125, 1997[Abstract/Free Full Text].

49.   Yoon, H, Fanelli A, Grollman EF, and Philp NJ. Identification of a unique monocarboxylate transporter (MCT3) in retinal pigment epithelium. Biochem Biophys Res Comm 234: 90-94, 1997[ISI][Medline].


Am J Physiol Endocrinol Metab 279(5):E1131-E1138
0193-1849/00 $5.00 Copyright © 2000 the American Physiological Society