When K+ output exceeds input, skeletal muscle releases intracellular fluid K+ to buffer the fall in extracellular fluid (ECF) K+. To investigate the mechanisms and muscle specificity of the K+ shift, rats were fed K+-deficient chow for 2–10 days, and two muscles at phenotypic extremes were studied: slow-twitch oxidative soleus and fast-twitch glycolytic white gastrocnemius (WG). After 2 days of low-K+ chow, plasma K+ concentration ([K+]) fell from 4.6 to 3.7 mM, and Na+-K+-ATPase α2 (not α1) protein levels in both muscles, measured by immunoblotting, decreased 36%. Cell [K+] decreased from 116 to 106 mM in soleus and insignificantly in WG, indicating that α2 can decrease before cell [K+]. After 5 days, there were further decreases in α2 (70%) and β2 (22%) in WG, not in soleus, whereas cell [K+] decreased and cell [Na+] increased by 10 mM in both muscles. By 10 days, plasma [K+] fell to 2.9 mM, with further decreases in WG α2 (94%) and β2 (70%); cell [K+] fell 19 mM in soleus and 24 mM in WG compared with the control, and cell [Na+] increased 9 mM in soleus and 15 mM in WG; total homogenate Na+-K+-ATPase activity decreased 19% in WG and insignificantly in soleus. Levels of α2, β1, and β2 mRNA were unchanged over 10 days. The ratios of α2 to α1 protein levels in both control muscles were found to be nearly 1 by using the relative changes in α-isoforms vs. β1- (soleus) or β2-isoforms (WG). We conclude that the patterns of regulation of Na+ pump isoforms in oxidative and glycolytic muscles during K+ deprivation mediated by posttranscriptional regulation of α2β1 and α2β2 are distinct and that decreases in α2-isoform pools can occur early enough in both muscles to account for the shift of K+ to the ECF.
- sodium-potassium-ATPase isoforms
- white gastrocnemius
extracellular fluid (ECF) and intracellular fluid (ICF) K+ levels are tightly controlled in mammals because their ratio is the principal determinant of cell membrane potentials. When whole body K+ output exceeds K+ input over time, the plasma K+ level falls, a condition known as hypokalemia (40). This condition can occur during loop diuretic treatment, which increases K+excretion, or if K+ intake is restricted, such as during prolonged fasting (40). In excitable tissues such as heart tissues, hypokalemia-induced disturbances in membrane potential can lead to life-threatening cardiac arrhythmias (10). K+ balance is maintained by the interplay of two key organ systems: the kidneys, which can secrete or actively reabsorb K+, and skeletal muscle, the major K+ reservoir, which can acutely control the transport of K+ between the ECF and the ICF (33, 40). Extracellular K+ loss is buffered by the transfer of muscle cell K+ to the ECF. This loss is likely mediated by the accompanying decrease in Na+ pump (Na+-K+-ATPase) levels, which would decrease active K+ transport from ECF to ICF (5,22, 36).
The Na+ pump is a ubiquitous integral membrane P-type ion pump that pumps Na+ out of the cell and K+ into the cell, a process driven by the hydrolysis of ATP (27). It is an αβ heteromer composed of a catalytically active α-subunit (M r ≈ 112,000) and a glycosylated β-subunit (M r ≈ 35,000), and recent evidence suggests that it is a tetramer (44). Multiple α- and β-subunit isoforms are expressed in a tissue-specific manner (13,29, 41, 42). In skeletal muscles, Na+ pump isoforms are expressed in a muscle fiber type-specific manner (19, 43). At the phenotypic fiber type extremes, slow-twitch oxidative soleus expresses α1-, α2-, and β1-isoforms as α1β1 and α2β1, whereas fast-twitch glycolytic white gastrocnemius (WG) expresses α1-, α2-, and β2-isoforms as α1β2 and α2β2 (43). Background information on the proportion of α2- to α1-type Na+ pumps in muscle is limited to two studies. For rat diaphragm membranes the fraction of α2 was estimated at between 30 (in hypothyroids) and 65% (in hyperthyroids) by a backdoor phosphorylation assay that measures pumps undergoing a reaction cycle (15). In rat red (oxidative) skeletal muscle subjected to subcellular fractionation on sucrose gradients, the ratio of α2 to α1 ranged from 1.6 (60% α2) in surface membranes to 7 (87% α2) in intracellular membranes. The ratio was calculated by using ouabain binding per milligram of protein to quantitate α2, and the immunoblot signal was scaled against the signal from a recombinant fragment of α1 per milligram of protein to measure α1 (23). In comparison, the proportion of α2 to total α mRNA in skeletal muscle has been reported as being between 70 and 80% (13,37). The ratio of total α2 to total α1 in control muscles, either oxidative or glycolytic, has not been previously determined.
We previously reported that, when rats are fed a K+-deficient diet for 10 days, Na+ pump protein changes are isoform and muscle fiber type specific (43): in fast-twitch glycolytic WG, α2 and β2 protein levels decrease to only 6 and 30% of control, respectively, whereas in slow-twitch oxidative soleus α2 decreased to 45% of control and β1 did not decrease significantly. The α1 protein pool size for either muscle did not change. Despite the substantially greater decrease in α2 and β2 in WG than in soleus, the fall in whole-muscle tissue K+ level was ∼20% after 10 days of K+ restriction in both muscles. These findings provoked questions about the muscle-specific mechanisms of cell K+ loss. The first aim of this study was to learn whether the decrease in α2 abundance is linked to the loss of cell K+ by comparing the time courses of the two parameters at early time points in the response. The second aim was to measure both protein and mRNA levels of Na+ pump subunits over the time course of K+ depletion to understand the molecular mechanisms responsible for the decrease in expression in different muscle types. The third aim was to assess the ratio of α2- to α1-isoforms because it will affect the impact of α2-specific regulation. The present study demonstrates that changes in α2 protein levels occur early enough to account for the loss of cell K+ in both muscle types, that the decrease in α2 protein is not secondary to a decrease in α2 mRNA, and that the calculated percentage of α2-type pumps in control muscles ranges from 40% in soleus to 55% in WG.
MATERIALS AND METHODS
Animals and diets.
Male Sprague-Dawley rats, ∼8 wk of age (250–300 g), were placed on a K+-deficient diet (TD 88239; Harlan Teklad, Madison, WI) for 2, 5, or 10 days and were paired to controls fed a comparable diet with K+ restored (TD 88238; Harlan Teklad). Rats were anesthetized with 0.2 ml pentobarbital sodium/100 g body wt. Soleus and WG hindlimb muscles were removed, frozen in liquid nitrogen, and stored at −80°C pending analyses. Blood samples were taken from the abdominal aorta, and the serum was separated, frozen, and stored at −20°C pending analyses.
Serum and intracellular electrolytes.
Serum and muscle cell K+ and Na+ concentrations ([K+] and [Na+]) were measured by flame photometry. Muscles were thawed, blotted lightly to remove adherent fluid, and homogenized in 0.3 M trichloroacetic acid [TCA; 1:50 (wt/vol)] for 5 min with a Tissuemizer homogenizer and then centrifuged at 2,500 rpm for 20 min to remove cell debris. The K+ and Na+ contents of the muscle TCA extracts and serum samples were measured with an FLM 3 flame photometer (Radiometer, Copenhagen, Denmark), with lithium as the internal standard (21). The total muscle [K+] and [Na+] were corrected for the levels of these cations in the extracellular space.
The extracellular space of soleus and WG was determined as previously described (47). In brief,l-[3H]glucose was infused, via a cannula in a tail vein, at 0.2 μCi/min for 2 h and then rats were anesthetized with pentobarbital sodium. Soleus and WG muscles were removed from each hindlimb, immediately frozen in liquid nitrogen, and stored at −80°C. Blood samples were collected from the abdominal aorta. Muscles were homogenized in 0.3 M perchloric acid [PCA; 1:10 (wt/vol)] for 5 min with a Tissuemizer, then centrifuged at 10,000 g, 4°C, for 15 min. The supernatant was cleared of PCA with 1:4 trioctylamine-1,1,2-trichlorotrifluoroethane (Freon), 1:2 (vol/vol), and assayed forl-[3H]glucose by liquid scintillation counting. Plasma samples (20 μl) were directly assayed in scintillation fluid. Extracellular space was calculated as musclel-[3H]glucose content divided by plasmal-[3H]glucose concentration (47). Muscle ion levels, measured by flame photometry (recorded as μmol/g wet wt and converted to μmol/ml wet wt), were corrected for the level of ions in the extracellular space (μmol/μl). Extracellular space values atday 0were used for control ion corrections;day 10 extracellular space values were used for corrections at all low-K+time points (days 2, 5, and 10).
Na+-K+-ATPase α- and β-subunit immunoreactivities.
These immunoreactivities were determined as previously described (43). In brief, skeletal muscle was homogenized 1:20 (wt/vol) in 5% sorbitol, 25 mM histidine-imidazole (pH 7.4), 0.5 mM EDTA disodium, and proteolytic enzyme inhibitors [0.5 mM phenylmethylsulfonyl fluoride (PMSF), 1 μg/ml leupeptin, and 1 mM 4-aminobenzamidine dichloride (pABAD)] with a Polytron homogenizer. To facilitate detection of the β-subunit, sugar residues were removed from β-subunits with PNGase F as previously described (43). A constant amount of homogenate protein (100 μg for α-subunit analysis, 50 μg for β-subunit analysis) was resolved by SDS-PAGE and blotted onto Immobilon-P membranes (Millipore, Bedford, MA). Blots were incubated overnight with one of the following antibodies: McB2, a monoclonal antibody specific for α2 (45), generously provided by K. Sweadner (Harvard Medical School); anti-β1 FP (1:500), a polyclonal antibody against β1 (43); SpET b2 (1:2,000), a polyclonal antibody against human β2 (14), generously provided by P. Martin-Vasallo (Universidad de La Laguna); and RNT β3 (1:2,000), a polyclonal antibody against rat β3 (4), provided by K. Sweadner. Blots probed with monoclonal McB2 were incubated for 2 h with rabbit anti-mouse IgG secondary antibody (Calbiochem, La Jolla, CA; 1:2,000). Antibody-antigen complexes labeled with 125I-protein A were visualized by autoradiography as described previously (28), and linearity was verified by assaying samples at multiple concentrations.
Na+ pump enzymatic activity.
Na+ pump activity in crude muscle homogenates was estimated by the K+-dependentp-nitrophenylphosphatase (K+-pNPPase) reaction (35), as recommended by Kjeldsen et al. (22) because levels of ouabain-sensitive Na+-K+-ATPase in skeletal muscle homogenate cannot be reliably determined, presumably because of the ouabain resistance of α1 in rats, the low abundance of Na+ pumps, and/or the high level of other ATPases in skeletal muscle. In brief, crude homogenates were freeze-thawed three times to permeabilize membranes, and 60 μg of homogenate protein were added to 500-μl sets of K+-pNPPase assay mixtures containing 100 mM KCl or 100 mM NaCl. The [Na+] carried over from the homogenization buffer was <0.5 mM. Activity is reported as micromoles of phosphate per milligram of protein per hour.
Na+-K+-ATPase α and β mRNA analysis.
Total RNA was isolated from rat skeletal muscle as described by Chomczynski and Sacchi (8) with Tri Reagent (Molecular Reasearch Center, Cincinnati, OH) according to the manufacturer’s protocol. RNA concentrations were determined by measuring the optical density at 260 nm (OD260), and purity was estimated by determining the OD260/OD280ratio. Total RNA was assayed by Northern analysis, as previously described (17, 32), on a NitroPure nitrocellulose transfer membrane (Micron Separations, Westborough, MA). Immobilized RNA was hybridized with isoform-specific restriction endonuclease fragments (∼300 bp) prepared from either α1, α2, or β1 clones as described by Orlowski and Lingrel (37) or from the rat β2 cDNA clone provided by P. Martin-Vasallo (30). The α- and β-cDNA probes were labeled to similar specific activities with [32P]dCTP probes by using a multiprimer DNA-labeling technique (12). Blots were washed three times in 2× SSC (0.3 M NaCl-0.03 M sodium citrate, pH 7.0) with 0.05% SDS at room temperature for 10 min each and then twice for 20 min each in 0.1× SSC with 0.1% SDS at 55°C. Autoradiograms of the blots were quantified by scanning densitometry.
Autoradiograms were quantitated by scanning with a GS670 imaging densitometer (Bio-Rad, Hercules, CA) and dedicated software. All data are expressed as means ± SE, normalized to the mean value at day 0, defined as 1. Significance was assessed by the two-tailed Student’st-test, and differences were considered significant at P < 0.05. The fractions of α1- and α2-isoforms in soleus and WG muscles were estimated as described inresults. Parameter identifications were performed with MLAB software (Civilized Software, Bethesda, MD), implemented on an IBM-compatible computer, which uses a Marquardt-Levenberg iterative least-squares algorithm. Again, data are reported as fractions of total pumps (defined as 1.0) ± SE.
Chemicals were reagent-grade, spectroquality, or electrophoresis purity reagents. SDS-PAGE reagents were from Bio-Rad. Leupeptin, PMSF, pABAD, and SDS-PAGE molecular weight standards were from Sigma Chemicals (St. Louis, MO). PNGase F (N-glycanase) was from Genzyme Corporation (Cambridge, MA).125I-protein A and [32P]dCTP were from ICN (Costa Mesa, CA).l-[3H]glucose was from DuPont NEN (Boston, MA).
Time course of change in extracellular [K+] during K+ deprivation.
In 8-wk-old rats placed on a K+-deficient diet, serum [K+] fell significantly from a control value of 4.6 ± 0.08 to 3.7 ± 0.09 mM by day 2 and continued to fall to 3.3 ± 0.2 and 2.9 ± 0.2 mM by days 5 and10, respectively (Fig.1). The fall in serum [K+] during the first 2 days (0.9 mM) is greater than or equal to the fall during the subsequent 8 days (0.8 mM), the time when renal and muscle adjustments to hypokalemia come into play (26, 48). Serum [Na+] did not change throughout the course of the study. The fall in serum [K+] is consistent with previous reports (22, 36) and demonstrates that K+ output exceeds input in this K+ deprivation model.
Time course of change in Na+-K+-ATPase isoform abundance vs. cell [K+] in soleus and WG muscles during K+ deprivation.
To determine whether changes in Na+ pump expression precede the decrease in cell [K+] and to assess the relative contributions of distinct muscles in the adaptive responses to hypokalemia, two muscles at phenotypic extremes were chosen for study: soleus, with 87% slow-twitch oxidative fibers and some fast-twitch glycolytic-oxidative fibers, and WG with 84% fast-twitch glycolytic fibers and some fast-twitch oxidative fibers (2,3).
The possibility that β3 was expressed in muscle and regulated during K+ deprivation was tested since β3 has been detected in skeletal muscle microsomes from both 7-day postnatal and adult rats (4). Immunoblot studies of both glycosylated and deglycosylated soleus and WG homogenates were conducted (not shown), and no β3 signal that shifted from a predicted glycosylated molecular mass of 55 kDa to a deglycosylated band at 35–38 kDa and/or that showed a positive signal at the appropriate molecular mass for deglycosylated β3 (35–38 kDa) was detected in adult soleus or WG. Adult rat testis homogenate, run on the same blots as a positive control, did yield glycosylated and deglycosylated signals for the β3-subunit at the appropriate molecular masses. We conclude that the β3-subunit, if present in adult rat skeletal muscle, is below the level of detection obtained with the currently available antibody on crude homogenate.
The relative expression of Na+-K+-ATPase α2- and β1-isoforms (soleus) and β2-isoforms (WG) after 0, 2, and 5 days of K+ deprivation was determined by immunoblotting. The previously reported immunoblot results for α2 and β after 10 days of K+ deprivation (43) are included in this analysis. The α1-subunit has been shown to be unchanged during 10 days of hypokalemia (5, 43). Typical autoradiograms of a subset of the samples are shown in Fig. 2. Control and K+-deprived samples at a given time point were run on the same gel and processed identically; samples for β detection were deglycosylated before analysis. The relative levels of expression, determined by scanning densitometry, are summarized in Fig. 3,A andB,top. In both muscles α2 protein expression decreased 36% as early asday 2. In soleus and WG, β1 and β2, respectively, were depressed significantly by 23% atday 5.
To compare the time course of change in Na+ pump subunits to that of cell [K+], muscle [K+] and [Na+] were measured after 0, 2, 5, and 10 days of K+deprivation. Extracellular space (expressed as μl/g wet wt) in the two muscles was measured withl-[3H]glucose, and that in soleus was found to be greater than that in WG (142 ± 6 vs. 72 ± 8 μl/g), as expected on the basis of the known difference in the respective levels of vascularization. K+ depletion for 10 days did not cause a significant change in extracellular space in either soleus (125 ± 11 μl/g) or WG (65 ± 10 μl/g). These findings agree with previous estimates of ECF space in muscle (1, 22). Intracellular [K+] and [Na+] were calculated as described in materials and methods. The changes in these cation concentrations in soleus and WG ICF are summarized in Fig. 3, A andB,bottom. For controls, ICF [K+] was higher, and ICF [Na+] was lower, in WG than in soleus, a difference which may reflect a lower level of muscle activity in fast-twitch glycolytic WG (thus less K+leak out and Na+ leak into the cell) compared to slow-twitch oxidative soleus (9, 20). After 2 days of K+ deprivation, soleus [K+] fell 9% (116 ± 3 to 106 ± 2 mM), and, although WG [K+] did not fall significantly (126 ± 3 to 120 ± 4 mM), WG [Na+] did increase (19.6 ± 1.2 to 23.2 ± 1.6 mM). From these results we conclude that the decrease in Na+ pump α2-isoform expression, 35–40% in both soleus and WG atday 2, occurs early enough in the time course of K+ deprivation to account for the changes in cell [K+]. Betweendays 2 and5, the two muscles responded quite differently: in soleus there were no further changes in α2, β1, or cell ion concentrations, whereas in WG there were significant further decrements in both α2 and β2 protein, which decreased 70 and 22%, respectively, and in cell [K+] which decreased ∼10% to 115.2 mM. Between days 5 and10 there were far greater decrements in Na+ pump expression in WG than in soleus: α2 and β2 decreased 94 and 70%, respectively, from control levels in WG, whereas in soleus α2 decreased 56% and β1 decreased no further. Despite these differences, the decrements in cell [K+] for the two muscles were not significantly different byday 10: [K+] fell 16 ± 2% in soleus and 19 ± 2.5% in WG. Because [Na+] increased as [K+] fell, the sum of cell [K+] and [Na+] was not significantly altered.
The observation that the decreases in α2 pools during the low-K+ diet are greater than the decreases in β1 or β2 pools at all time points is expected because a single β must form heteromers with not only α2 (which decreases) but also with α1 (which is invariant). In other words, the difference between the change in α2 and β (β1 in soleus, β2 in WG) will be a function of the ratio of expression of α1β to α2β heteromers in that muscle, as calculated in the next section.
Estimating the proportion of the Na+-K+-ATPase α2-isoform in soleus and WG.
It has been difficult to directly assess the ratio of α1- to α2-type Na+ pumps in muscle. Although absolute pool sizes of α2 vs. α1 protein subunits cannot be determined directly with antibody probes, muscle-specific changes in α2 vs. β1 or β2 abundance during K+ deprivation can be employed to calculate the ratios of α2 to α1. The calculations depend on three assumptions. First, α1 and α2 proteins are assumed to combine with only β1 in soleus and only β2 in WG in a 1:1 stoichiometry. Indeed, recent evidence suggests that Na+pumps may exist as tetramers with a 1:1 α-to-β stoichiometry in mammalian cells (44). Second, it is assumed that there are only negligible pools of uncomplexed α- or β-subunits, a difficult assumption to test in this system but supported by evidence that α- and β-subunits form complexes as αβ heteromers before leaving the endoplasmic reticulum and that increasing the synthesis of one subunit increases the stability of the other, implying that uncomplexed subunits are less stable (discussed in Ref. 25). Third, the abundance or pool size is a linear function of the autoradiographic signal, a fact established in previous investigations (6, 28, 46). It follows from these assumptions that the total number of β-subunits is equal to the total number of α-subunits (Eq. 1 ), the total number of α-subunits is equal to the sum of α1- and α2-subunits (Eq. 2 ), and thus that the total number of β pumps is equal to the sum of α1 and α2 pumps (Eq. 3 ) Equation 1 Equation 2 Equation 3The number of α- or β-subunits is equal to the measured immunoblot autoradiographic signal (S), expressed as fraction of control, times a constant (C) related to antibody efficiency (Eqs.4-4) Equation 4a Equation 4b Equation 4cThusEq. 3 becomes Dividing by Cβ Defining A as (Cα1/Cβ) and B as (Cα2/Cβ) Equation 5Because the α1 signal does not change during hypokalemia, Sα1 = 1 (control defined as 1) and Equation 6A and B are estimated by regression analysis, and relative proportions of α1 and α2 pumps are calculated for any time point from the immunoblot signals as follows Equation 7 Equation 8 The fractions of pumps that are α1 and α2 were calculated by simple linear regression (Eq. 6 ) and are summarized in Table1. Similar values were obtained by employing multiple regression analysis (Eq. 5 ) using α1 signals at either 0 or 10 days. In the control state, α2 is calculated to make up 0.4 ± 0.12 of total α in soleus (0.6 ± 0.12 α1) and 0.55 ± 0.07 of total α in WG (0.45 ± 0.07 α1). After 10 days of K+ deprivation, α1 is estimated at 0.76 ± 0.09 of the total pumps in soleus and 0.92 ± 0.09 of the total pumps in WG.
Because the calculated ratios of α1 and α2 depend on a linear regression analysis of measurements made at all four points in the time course, there is a finite error between the calculated and measured values of variability from one specific time point to the other. For example, the calculated amount of β2 atday 5(assumed to equal α1 + α2) should be 45% + 55%(0.3) = 62% of the control value, an underestimate of the 80% β2 measured by immunoblotting at this time point (Fig. 3), and the amount of β2 atday 10 should be 45% + 55%(0.08) = 50% of control, an overestimate of the 30% β2 measured by immunoblotting at day 10 (Fig. 3). However, this error does not change the prediction that there are nearly equivalent pools of α1 and α2 at the zero time point. Adding additional time points to the linear regression analysis would help to bring the calculated estimates closer to measurements of β.
Na+ pump enzymatic activity.
Na+-K+-ATPase activity in muscle homogenates of soleus and WG was measured via the K+-pNPPase reaction as micromoles of Pi per milligram of protein per hour. Activities in soleus and WG were 0.034 ± 0.004 and 0.033 ± 0.002 μmol Pi ⋅ mg protein−1 ⋅ h−1, respectively (n = 6 for each). After 10 days of a K+-restricted diet, activity was 0.028 ± 0.002 μmol Pi ⋅ mg protein−1 ⋅ h−1in soleus, which was not significantly different from control, and 0.026 ± 0.002 μmol Pi ⋅ mg protein−1 ⋅ h−1(P < 0.05) in WG (n = 6). This 19% decrease in WG activity falls short of the 70% decrease predicted from the decrease in the β2 pool in this tissue. In comparison, in soleus there is only a 20% fall in β1, which comes close to the measured change in total activity. We conclude that enzymatic activity is not a direct function of abundance. For example, the change in cell Na+ that ensues during K+ deprivation may provoke a modification of the α1, in WG which persists through homogenization, that changes activity per pump.
Na+-K+-ATPase mRNA expression in soleus and WG muscles during K+ deprivation.
Considering the 35–40% decreases in α2 in both muscles after 2 days of K+ deprivation and the >90% decrease in WG α2 after 10 days, we tested the hypothesis that the response was driven by decreases in α2 mRNA in both muscles. Na+-K+-ATPase α2- and β-isoform mRNA levels in both muscles were measured after 0, 2, 5, and 10 days of K+deprivation. Northern blot results indicate no change in α2-, β1-, or β2-isoform mRNA expression in either muscle during 10 days of K+ deprivation (Fig.4, A andB). Although distinct bands at the appropriate sizes were observed by this analysis, the quality of the RNA was compromised by the time it takes to dissect the muscles. To circumvent this problem and to reexamine our previous report of a decrease in whole hindlimb α2 mRNA in rats deprived of K+ (5), we repeated the mRNA analysis in whole hindlimb muscle isolated as quickly as possible. The results confirmed the findings for the individual muscles: α2, β2 (Fig. 4 C), and β1 (not shown) mRNA levels in whole hindlimb did not change during 10 days of K+ deprivation.
Finally, we tested the possibility that there was a shift in β-isoform expression as hypokalemia progresses. Northern blots confirmed that even at day 10, when β2 protein had fallen 70%, WG did not express the β1 transcript. Similarly, β2 mRNA was not detected in soleus after K+restriction (data not shown).
The existence of Na+ pump isoforms suggests the potential for isoform-specific function, expression, and regulation (31). The need to shift K+ from intracellular stores to the ECF is satisfied by tissue-specific expression and regulation of the Na+-K+-ATPase α2-isoform in skeletal muscle. This report demonstrates that changes in the α2-, not the α1-, isoform occur early enough during K+ deprivation to drive the shift of cell K+ to the ECF, that there are distinct patterns of regulation in slow-twitch oxidative vs. fast-twitch glycolytic muscles, and that the regulation of α2 occurs by posttranscriptional regulation. As muscle cell [K+] falls during K+ deprivation, cell [Na+] reciprocally increases to as high as 37 mM in WG, which is a classical stimulus for increasing Na+ pump activity in most tissues (11). However, skeletal muscle has an altruistic response to elevated cell [Na+]: Na+ pump number is decreased, which leads to a further loss of K+ and gain in Na+ in order to contribute intracellular K+ to the K+-depleted ECF. Extracellular [Na+] does not change reciprocally with [K+] during this early phase of hypokalemia because the amount of Na+ in the ECF is a primary determinant of the ECF volume, a condition that keeps the concentration fairly constant.
In the first 2 days of dietary K+restriction there is a more rapid decrease in ECF [K+] (a 20% fall) than that over the subsequent 8 days (an additional 22% fall) (Fig.1). The reduced rate of ECF K+loss beyond day 2 is not unexpected because it takes ∼48 h for rats on K+-restricted diets to induce mechanisms to actively reabsorb K+ and maximally reduce urinary K+ excretion, mediated by the induction of renal collecting duct H+-K+-ATPase activity (26, 48).
The responses in soleus and WG are indistinguishable during the initial 2 days of K+ restriction (Fig. 3): both muscles lose over 36% of the initial α2-subunits, and changes in muscle [K+] and [Na+] are similar. Kjeldsen et al. (22) reported that after 3 days of K+ deprivation ouabain binding to whole soleus muscle (a measure of active α2-type pumps at the plasma membrane) decreased 15%. That the reduction in α2 Na+ pump pool sizes in soleus is greater than the decrease in surface ouabain binding sites suggests that there is some ouabain binding by α1 in soleus or that the decrease in α2 pumps may include both surface and nonsurface pools. Even though α2 levels in both muscles fall 36%, intracellular [K+] is only beginning to fall in soleus and does not fall significantly in WG, evidence that the decrease in α2 precedes, and likely accounts for, the decrease in muscle K+ stores. The finding also indicates that a loss of 36% of the muscles’ Na+ pumps is not associated with a rapid change in cell K+ stores. That is, the remaining α2 and invariant α1 pumps are capable of nearly matching active K+ influx to passive K+ outflux, so that cell K+ stores change slowly and progressively during the 10 days of the K+-restricted diet. In previous studies, losses in total muscle K+levels as high as 11% in whole gastrocnemius muscles after 3 days of K+-deficient fodder have been reported (22).
There is a provocative divergence in the responses of soleus and WG to K+ deprivation after 2 days (Fig.3). In soleus, neither total α2 Na+ pump abundance nor cell K+ level decreases further betweendays 2-5, but both decrease between days 5 and10. Thus changes in soleus [K+] mimic the changes in soleus α2 immunoreactivity, evidence for a causal link between the two. The lack of change in cell [K+] or [Na+] betweendays 2 and5 of K+ restriction in soleus may be influenced by the fact that slow-twitch oxidative soleus muscle is more sensitive to changes in intracellular [Na+] than fast-twitch muscles (11), that is, the increases in muscle [Na+] might stimulate Na+ pumps and retard the loss of K+ more in slow-twitch oxidative soleus than in fast-twitch glycolytic WG. In WG, total α2 Na+ pump abundance falls dramatically throughout the 10 days of K+ restriction. Byday 5, the fall in WG is twofold greater than that in soleus, and by day 10 only 6% of the total α2 pools remain in WG. Despite this large difference between the muscles, they both lose about the same percentage of cell K+. One hypothesis is that both muscles lose the same fraction of Na+ pumps expressed in the plasma membrane, with soleus storing pumps in endosomal pools during K+ restriction.
What is the magnitude of the physiological impact of the shift of K+ from the muscle ICF to the ECF? ECF [K+] would fall precipitously without the skeletal muscle adjustment because the amount of K+ in the ECF is small. In fact, the amount of K+ shifted from ICF to ECF is more than seven times the amount of K+ contained in the ECF of a control animal, a result calculated as follows (assumptions from Ref. 40). If we assume that ECF is 20% of the body weight, then a 280-g rat would have an ECF of 0.056 liters. Because ECF [K+] is 4.5 mM (Fig.1), the control ECF contains ∼0.25 mmol K+. Assuming that ICF is 40% of the body weight, the same rat would have an ICF of 0.112 liters. Because muscle contains ∼80% of the ICF (0.090 liters) and muscle ICF [K+] is 120 mM (Fig. 3), muscle ICF contains 10.8 mmol of K+, roughly 40 times more than that in the ECF pool. After 10 days of the low-K+ diet, muscle loses an average of 17% of the intracellular stores (Fig. 3), equivalent to 1.84 mmol, which is more than seven times the amount contained in the ECF at day 0 (0.25 mM). In other words, the extracellular K+ has been replaced seven times over with K+ from the muscle stores after 10 days of a K+-restricted diet, evidence that this regulatory adjustment is critical.
The calculated ratio of α2 to α1 of near 1:1 in both muscles is not what one would predict from the relative RNA levels in control muscles. RNA ratios can be estimated directly with isoform-specific cDNA probes of similar lengths and labeled to similar32P specific activities. mRNA ratios of α2 to α1 in skeletal muscle have been reported to be between 2.5 (0.7 α2 and 0.3 α1) (13) and 4 (0.8 α2 and 0.2 α1) (37). However, there is no a priori reason to assume that mRNA ratios are good predictors of α protein ratios because other factors, from isoform-specific translatability to competition for heteromer formation with β1 or β2, will influence the protein ratio. Our estimate of 50% α2 in soleus agrees with what was observed by backdoor phosphorylation (15), assuming that the fraction of α2 in the euthyroid diaphragm is midway between that observed in the hypothyroid (30%) and hyperthyroid (65%) diaphragms. The percentage of α2 in soleus, calculated by Lavoie et al. (23) at between 60% in the surface membrane and 87% in intracellular membranes, is higher than our estimate. This is not unexpected because the subcellular membrane fractions assayed are expected to be enriched in α2, and it has been established that the subcellular distribution of α2 is different from that of α1 in red muscle (18). In comparison, in this study we calculated the α2-to-α1 ratio in total homogenate, which contains all the cell membranes, thus averaging the ratio in membranes enriched in α2 with those enriched in α1.
Azuma et al. (5) reported that, in rats maintained on a low-K+ diet for 14 days, hindlimb α2 mRNA decreased 35% (α1 and β1 were unchanged) and that α2 protein decreased by 82%, suggesting that changes in α2 mRNA alone could not account for the changes in α2 protein (5). Because the whole hindlimb in rats is a composite of different muscles, it was conceivable that the mRNA changes were muscle specific. We have previously determined that during the transition from euthyroid to hypothyroid states, changes in α2 mRNA levels in mixed gastrocnemius muscle predicted the changes in α2 protein levels (both decreased 45%) (6). However, unlike what was found for regulation by thyroid status, mRNA levels in either soleus, WG, or whole hindlimb measured in this study did not change during 10 days of K+ deprivation (Fig. 4). Thus we do not confirm the observations in our previous study (5) and hypothesize that α2 mRNA levels decrease only after a K+ deprivation of more prolonged duration. Alternatively, it is possible that the K+-deficient diet used in the present study is better matched to the control diet than in the previous report, in which the weights of the K+-deprived rats were significantly lower than the controls after 14 days of the low-K+ diet. We conclude that the decreases in Na+ pump expression during K+ deprivation can be explained by either a decrease in α- and β-transcript translatability and/or increased protein degradation. A rapid increase in protein degradation in muscle is not without precedent. The ubiquitin-proteasome pathway plays a role in increasing the rate of muscle protein degradation during denervation, fasting, or insulinopenia (34). Plasma membrane proteins are also degraded by internalization to endosomal pools and routing to lysosomes. The α2β1-type heteromers are known to shuttle between endosomal pools and the plasma membrane with insulin stimulation in oxidative muscles such as soleus but not in glycolytic muscles (24). These findings suggest the hypothesis that when ECF [K+] falls the rate of Na+ pump internalization increases in both muscle types, that a portion of the internalized pumps are stored in endosomal pools in the soleus and are available for return to the plasma membrane with K+restoration or insulin stimulation, and that pumps are routed for degradation in the lysosomes in both muscles. Such a pattern would account for the smaller decrease in α2 in soleus than in WG, along with the similar rates of loss of K+ from both muscle types (Fig.3).
How tissues like muscle or kidney sense the fall in extracellular [K+] and then effect tissue-specific changes such as the decrease in muscle Na+-K+-ATPase α2 levels remains unclear. One theory is that there are K+ sensors in the gut, portal circulation, and/or liver that respond to local changes in extracellular [K+], secondary to enteric changes (39). This suggestion complements the recent identification of the calcium-sensing receptor (CaR) (7). Indeed, Quarles et al. (38) have theorized that CaRs may represent one member of a family of “cation-sensing” cell surface receptors. Further, Hevener et al. (16) localized glucosensors to the portal vein. When these results are taken together, it is tempting to speculate that both glucose and K+ sensors in the hepatic portal vein or liver may respond to dietary intake, stimulating the release of humoral factors (such as insulin when glucose and K+ levels are elevated) that alter muscle and kidney K+ handling by regulating transporter levels.
In conclusion, our results demonstrate a temporal relationship between the decreases in α2 Na+ pump pools and a coincident or subsequent decrease in muscle [K+] during K+ deprivation in both slow-twitch oxidative and fast-twitch glycolytic muscles. The study establishes there are muscle type-specific mechanisms in place to effect the shift of K+ from ICF to ECF and that in both muscles the changes are independent of changes in mRNA levels.
This work was supported by National Science Foundation Grant IBN 9S13958 to A. A. McDonough and J. N. Youn and by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-34316 to A. A. McDonough.
Address for reprint requests and other correspondence: A. A. McDonough, Dept. of Physiology and Biophysics, Univ. of Southern California School of Medicine, 1333 San Pablo St., Los Angeles, CA 90033.
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- Copyright © 1999 the American Physiological Society