Apart from severe bone mineralization defects, alopecia, and other metabolic defects, vitamin D receptor knockout (vdr−/−) mice develop severe skeletal muscle atrophy.1 Vitamin D deficiency induces myopathy in humans as well as experimental animals,2 and vitamin D receptor (VDR) is shown to be expressed in skeletal muscles, albeit at low levels.3, 4 But the aetiology of skeletal muscle atrophy in the absence of VDR signalling is unknown. Several studies have shown that vitamin D signalling defects lead to metabolic impairments including insulin resistance. But the role of metabolic derangement in the origin of myopathies in the absence of vitamin D signalling is unexplored.
Protein homeostasis is the major determinant of skeletal muscle mass.5 Muscle protein synthesis increases in response to nutrients and growth factors while during the post-absorptive state, muscle catabolizes proteins and releases amino acids to the bloodstream.6 Muscle protein homeostasis is important for maintenance of organismal energy homeostasis. Skeletal muscles function as a major site of glucose disposal and fatty acid utilization. Furthermore, energy deprivation conditions lead to increased muscle proteolysis so that the amino acids released can be used for gluconeogenesis,7, 8 thus linking systemic energy metabolism to muscle mass. Conversely, changes in skeletal muscle metabolism affect the systemic energy metabolism as evidenced by the rewiring of whole body glucose metabolism upon disinhibition of muscle fatty acid oxidation.9
Multiple factors can induce an imbalance in these activities, often leading to skeletal muscle atrophy. For example, when the demand for amino acids is increased under conditions such as sepsis and burns, muscle protein degradation and amino acid release are up-regulated substantially, leading to skeletal muscle atrophy.10, 11 Alternatively, energy deprivation conditions such as starvation, cachexia, and type I diabetes also lead to skeletal muscle atrophy.7, 8 Here, we explored the role of metabolic impairments in the development of skeletal muscle atrophy in vdr−/− mice.
Some evidence from vitamin D-deficient rat models had shown increased proteolysis in skeletal muscles.12 Moreover, mTOR activity, which plays an important role in the skeletal muscle mass, is also reduced in this model,13 clearly suggesting the role of vitamin D signalling in maintaining protein homeostasis in skeletal muscles. But the molecular and metabolic defects that drive these changes in the absence of vitamin D signalling are unknown. Even though myocyte-specific VDR knockout mice exhibited sarcopenia suggesting that muscle intrinsic VDR signalling is important for maintenance of muscle strength and function during ageing,14 these mice as well as skeletal muscle-specific VDR knockout mice did not display the severe skeletal muscle pathogenesis observed in whole body VDR knockout, suggesting a role for systemic or local modulators in the onset of atrophy in vdr−/− mice.14, 15
In this study, we attempted the root cause analysis of skeletal muscle atrophy in vdr−/− mice. Indeed, this genetic model of vitamin D deficiency also exhibits dysregulation of muscle proteostasis characterized by increased proteolysis, decreased mTORC1 activity, and fasting protein synthesis. These proteostasis defects are associated with a systemic energy deprivation indicated by hypoglycaemia and hyperlactataemia. As a result, AMPK pathway and Foxo1 transcription factor were activated in skeletal muscles, leading to up-regulation of the atrophy markers. Further attempts to identify the cause for the energy deficiency and atrophy revealed dysregulation of glycogen synthase and glycogen phosphorylase activities resulting in increased glycogen storage levels in the skeletal muscles. Data presented here show that defective muscle and systemic energy metabolism defects caused by glycogen storage disorder in skeletal muscles drive severe muscle atrophy in vdr−/− mice.
Methods Animal maintenanceVitamin D receptor null mutant mice were obtained from Jackson Laboratories (Stock No. 006133, B6.129S4-Vdrtm1Mbd lJ; Bar Harbor, ME, USA) and bred at the Small Animal Facility at the National Institute of Immunology, New Delhi, India. Heterozygous mating pairs were set up, and genotyping was done for each litter. vdr−/− and vdr+/+ from the same litter were used as experimental and control groups. All mice were fed ad libitum with a commercially available rodent chow (Altromin International). Both male and female mice were used for the study as there was no difference in the parameters checked. Experiments were performed on mice at 3, 5, or 7 weeks of age as indicated in the text. Mice were housed in a sterile facility in individually ventilated cages (IVC).
RNA extraction and quantitative RT–PCRQuad muscles were dissected and snap-frozen in liquid nitrogen and homogenized in RNAiso plus Reagent (TAKARA, 1 mL/100 mg tissue) to isolate total muscle RNA as per the manufacturer's recommendations. Total RNA was quantified with a Nanodrop 8000 Spectrophotometer (Thermo Scientific, Wilmington), and a ratio of 2 for the absorbance of 260 to 280 nm was used to determine the RNA quality. First-strand cDNA was synthesized from total RNA using the PrimeScript First-Strand cDNA Synthesis Kit with PrimeScript Reverse Transcriptase according to the manufacturer's protocols (TAKARA). Quantitative RT–PCR was performed using the QuantStudio-6 and -7 Flex Real-Time PCR Systems (Thermo Fisher Scientific) with SYBR® Premix Ex Taq (Tli RNase H Plus) by TAKARA. Each sample was amplified in triplicates using primers specific to Murf1, Fbxo32, GBE1, glycogenin, Glut1, Glut4, UCP2, UCP3, and glyceraldehyde 3-phosphate dehydrogenase (GAPDH). The expression levels of each transcript were normalized to the housekeeping gene GAPDH. The primer sequences used are available upon request.
Western blotting and antibodiesProtein extracts from the hindlimb muscles of wild-type (WT) and VDR−/− mice were obtained by homogenizing muscles in lysis buffer (50 mM Tris–HCl pH 7.5, 150 mM NaCl, 5 mM EDTA, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS) containing protease/phosphatase inhibitor cocktail (Cell Signaling Technology). The proteins were resolved by SDS-PAGE (8%, 10%, 12%, and 15%) and transferred onto nitrocellulose membrane (BioRad and Himedia). The membranes were incubated with primary antibodies followed by incubation with HRP-conjugated anti-mouse or anti-rabbit secondary antibodies and visualized using Supersignal West Pico Chemiluminescent Substrate (Thermo Scientific, Rockford, IL, USA). Blots were probed with antibodies against the gene of interest.
Glucose tolerance and serum insulin estimationTo examine alterations in glucose tolerance and insulin secretion, WT and homozygous VDR mutant mice were deprived of food in their normal cage environment for a period of 7 h. During that period, the mice had free access to water. Glucose (2 mg/kg body weight) was administered at time 0 by intravenous injection in saline. Blood glucose levels at 0, 10, 30, 60, and 120 min were determined in whole blood from the tip of the tail using the Accu-Chek Active Glucometer. Serum samples for measurement of insulin levels were obtained from the retro orbital venous plexus at baseline and 30 min after an intraperitoneal glucose challenge.
In vivo estimation of muscle protein synthesisFor SUnSET assay, mice were fasted for 7 h and then given an intraperitoneal injection of 0.040 μmol/g puromycin dissolved in 100 μL of PBS. At exactly 30 min after injection, tibialis anterior (TA) muscle was collected and frozen in liquid N2 for western blot analysis as described in a previous section. A mouse IgG2a monoclonal anti-puromycin antibody (clone 12D10, 1:5000) was used to detect puromycin incorporation. To check the postprandial protein synthesis, 200 μL of food dissolved in water was given via oral gavage. Puromycin was injected 30 min after feeding, and TA muscle was harvested 30 min post-puromycin injection.
In vivo estimation of protein ubiquitinationMice at 7 weeks were injected intraperitoneally with a proteasomal inhibitor, MG132 (2.5 mg/kg/day in 20% DMSO daily) for 2 days. Mice were euthanized, hindlimb muscles were harvested and lysed, and western blot analysis was performed as described in a previous section. To detect total protein ubiquitination, anti-ubiquitin antibody (Cloud-Clone Corp.) was used. To detect the K48-linked ubiquitination, anti-K48 ubiquitin antibody (Cell Signaling Technology) was used.
ATP estimation in musclesATP estimation was done using the kit from Thermo Scientific. A total of 100 mg of tissue was chopped using a scalpel into small pieces and sonicated in a modified RIPA buffer with EDTA (150 mM NaCl, 1% NP40, 0.5% Na-deoxycholate, 0.1% SDS, 25 mM Tris, pH 7.4) and centrifuged at maximum speed for 15 min at 4°C. The supernatant was collected and proceeded with the assay according to the manufacturer's protocol.
Histological analysesSkeletal muscle tissue (TA) for histology was collected from 7-week-old WT and frozen in isopentane chilled in liquid nitrogen. Ten-micrometre sections were cut using a cryotome (Thermo Scientific). These sections were stained with periodic acid-Schiff (PAS) according to the manufacturer's protocol (Sigma). The muscle sections were blindly analysed by a pathologist, and the pics were taken at ×400 magnification (scale bar = 20 μm). For succinate dehydrogenase (SDH) staining of muscle sections, frozen sections of skeletal muscles were incubated for 50 min in a humidified chamber at 37°C with incubation media [100 mM phosphate buffer, pH 7.6, 1.2 mM nitroblue tetrazolium (NBT), and 100 mM sodium succinate] followed by imaging.
Serum calcium, lactate, growth hormone, and insulin-like growth factor 1 estimationSerum lactate was estimated using lactate estimation kit (Abcam #ab65331) and calcium estimation kit (Abcam #ab102505) according to the manufacturer's instructions. Growth hormone (GH) and Igf1 were estimated using ELISA kits as per the manufacturer's protocols (Krishgen Biosystems, Mumbai, India).
Muscle succinate dehydrogenase and pyruvate dehydrogenase estimationsAssays were done from muscle lysates prepared in the assay buffer as per the manufacturer's protocol (Sigma-Aldrich).
StatisticsDensitometric quantification of western blots was performed using ImageJ software. All experiments were repeated at least three times. Number for samples per each figure is indicated as data points in the figure. For western and real-time PCR quantifications, pairwise comparisons were made using parametric, unpaired T-test with unequal variance. For all other experiments, a parametric, paired T-test with equal variance was used. F test was performed to determine if samples had equal variance or unequal variance. Graphs show the mean of the indicated number of replicates, and error bars represent standard error of the mean (SEM). All graphs were prepared using GraphPad Prism software Version 9.
Results vdr−/− mice exhibit higher muscle protein degradationAs reported previously,1, 16 body weights of the vdr−/− mice were comparable with that of WT littermates in the pre-weaning stage, whereas in the post-weaning stage, they exhibited progressive growth retardation as indicated by reduced body weight at 5 and 7 weeks of age (Supporting Information, Figure S1a). These mice exhibit a high rate of mortality beyond 8 weeks of age.17 Additionally, vdr−/− mice also exhibited reduced weights of TA, gastrocnemius (GAS), and quadriceps (QUAD) muscles at 7 weeks of age (Figure S1b–S1d) as documented in previous studies.1 Histopathology analysis of cross sections of TA and GAS muscles also shows reduced fibre size in these muscles (Figure S1e–S1g).
Next, we asked if the reduced muscle mass in vdr−/− is indicative of active muscle wasting or the failure of these mice to grow. Because the GH–insulin-like growth factor (IGF) 1 axis controls the growth as well as muscle mass in mammals,10, 18 we checked their levels in the sera of these mice and found reduced GH levels (Figure ). Because GH release by pituitary is pulsatile, there can be temporal variations in its levels. So, we checked the levels of IGF1 as it is induced by GH, more stable in serum, and acts as the major downstream effector of GH.19 Intriguingly, IGF1 levels were unchanged in the serum of these mice (Figure 1B), suggesting that the GH–IGF1 axis may not be drastically affected in these mice. Irrespective of the inducing factor, severe muscle atrophy is almost always associated with increased protein degradation and the up-regulation of muscle-specific E3 ubiquitin ligases.20 We analysed protein ubiquitination levels in both 3- (no atrophy) and 7-week-old (severe atrophy) WT and vdr −/− mice as a proxy for proteasomal degradation. After 2 days of intraperitoneal injection with the proteasome inhibitor MG132, both total ubiquitination and K48-linked ubiquitination, which is specific to proteasomal degradation pathway, were high in 7-week-old vdr−/− skeletal muscle (Figure 1C). Two major muscle-specific E3 ubiquitin ligases Fbxo32 and MuRF1 are well-known atrophy markers and are up-regulated in the majority of skeletal muscle atrophies.21 Indeed, the mRNA levels of both Fbxo32 and MuRF1 were up-regulated in the 7-week-old vdr−/− compared with WT (Figure 1D and 1E), highlighting that the up-regulation of proteasomal protein degradation is associated with the muscle atrophy in these mice.
vdr−/− mice exhibit higher muscle protein degradation. (A and B) Serum levels of growth hormone (A) and IGF1 (B) measured using ELISA. (C) Anti-ubiquitin and anti-ubiquitin–K48-linkage western blots of hindlimb skeletal muscle extracts of mice treated with proteasome inhibitor MG132. Ponceau staining of the blot is shown for equal loading. (D and E) Expression levels of Fbxo32 (D) and MuRF1 (E) mRNAs in hindlimb muscles of 7-week-old WT and vdr−/− quantified using qRT–PCR (n ≥ 4). (F) Representative western blot for Foxo1 protein levels in hindlimb skeletal muscles of 3- and 7-week-old WT and vdr−/− mice. (G and H) Western blot showing the levels of phospho-Foxo1 (Ser256) (G) and the ratio of phospho-Foxo1 to total Foxo1 (H). (I) Western blot analysis showing nuclear localization of Foxo1 in WT and vdr−/− mice. Anti-TBP antibody was used as a nuclear marker, and anti-GAPDH was used as a cytoplasmic marker. All graphs show mean ± SEM. Statistical significance was determined by unpaired T-test (*: P < 0.05, **: P < 0.01, ***: P < 0.001, ****: P < 0.0001).
FoxO proteins are the key transcription factors that drive muscle atrophy in response to a variety of stimuli. Furthermore, overexpression of Foxo3 and Foxo1 induces atrophy in skeletal muscles.22 Intriguingly, neither the total protein levels nor the nucleo-cytoplasmic localization of Foxo3 protein were altered in vdr−/− compared with WT (Figure S2a–S2c). On the other hand, as observed previously,15 Foxo1 protein levels were significantly increased in the 7-week-old vdr−/− muscles (Figure 1F). The ratio of Foxo1 phosphorylated at Ser256, which is the inactive cytoplasmic form, to the total Foxo1 was dramatically reduced in these mice (Figure 1G and 1H), suggesting that abnormal activation of Foxo1 is responsible for the increased transcript levels of Fbxo32 and MuRF1 as well as the increased protein ubiquitination levels. In support of this, the nuclear localization of Foxo1 was also higher in vdr−/− skeletal muscles (Figure 1I).
mTORC1 pathway and fasting protein synthesis are down-regulated in the skeletal muscles of vdr−/− miceApart from the increased proteasomal protein degradation, defective regulation of protein synthesis is also associated with several skeletal muscle atrophy models.18, 23 Muscle protein synthesis is up-regulated by amino acids as well as insulin during the absorptive state. On the other hand, during the post-absorptive state, only basal rates of protein synthesis are maintained.6 To address if there are defects in protein synthesis in absorptive and fasting states in the skeletal muscles of vdr−/− mice, we performed in vivo SUnSET assay. WT and vdr−/− mice were either fasted for 7 h (fasting protein synthesis) or fed a liquid diet after fasting by oral gavage (absorptive state protein synthesis) followed by an intraperitoneal injection of puromycin. Western blot analysis of hindlimb skeletal muscles harvested 30 min after puromycin injection showed similar levels of puromycin incorporation in 3-week-old WT and vdr−/− fasted mice (Figure 2A). On the contrary, 7-week-old vdr−/− mice exhibited a drastic decrease in fasting protein synthesis compared with their WT counterparts (Figure 2B). Interestingly, there was no difference in the rates of absorptive state protein synthesis in the muscles of 7-week-old WT and vdr−/− mice (Figure 2C). These results show that muscle atrophy in vdr−/− mice is associated with a proteostasis defect characterized by a general increase in protein degradation and a decrease in post-absorptive protein synthesis.
mTORC1 pathway and fasting protein synthesis are down-regulated in the skeletal muscles of vdr−/− mice. (A–C) Western blots using anti-puromycin antibody for estimation fasting protein synthesis in 3-week-old (A) and 7-week-old (B) and absorptive state protein synthesis in 7-week-old (C) hindlimb muscles of WT and vdr−/−. Image of a Ponceau stained blot is used as loading control. (D) Western blots for phospho and total mTORC1 protein in 3- and 7-week-old mice. (E) Western blots for S6 and 4EBP levels in 3- and 7-week-old mice. (F–H) Quantitation of mTOR protein (F), phospho-S6 (G), and 4EPB (H) bands from (D) and (E). All graphs show mean ± SEM. Statistical significance was determined by unpaired T-test (*: P < 0.05, **: P < 0.01, ***: P < 0.001, ****: P < 0.0001).
To investigate the molecular mechanisms underlying altered protein synthesis, we examined the mammalian target of Rapamycin Complex 1 (mTORC1) signalling pathway, which acts as the central regulator of cell growth partly by increasing protein synthesis.24 Recent studies have shown evidence for dysregulation of mTORC1 in a vitamin D-deficient rat model system.12, 13 When compared with WT, mTOR protein levels were unchanged in 3-week-old vdr−/− mice (Figure 2D). On the other hand, 7-week-old vdr−/− mice exhibited significantly lower levels of mTOR protein, suggesting that skeletal muscle atrophy is associated with down-regulation of mTORC1 pathway (Figure 2D and 2F). Concomitantly, levels of mTOR phosphorylated at Ser2448 and ribosomal protein S6 phosphorylated at Ser240/244, both performed by the mTORC1 target p70-S6K, were reduced in 7-week-old vdr−/− muscles, confirming the down-regulation of the mTORC1 pathway (Figure 2E and 2G). Additionally, western blot analysis of 4EBP1, a repressor of cap-dependent translation and an mTORC1 target,25 displayed increased protein levels in 7-week-old vdr−/− muscles, with the majority of protein belonging to the fast migrating, hypophosphorylated band (Figure 2E and 2H), further confirming reduced mTORC1 activity. Taken together, our observations strongly indicate that reduced protein synthesis driven by defective mTORC1 activity and increased protein degradation induced by Foxo1 activation could be driving skeletal muscle atrophy in vdr−/− mice.
vdr−/− skeletal muscles exhibit energy deprivationNext, we asked what is the root cause of the defective proteostasis in vdr−/− mice. Various triggers ranging from inflammation to metabolic disorders and energy deprivation can affect skeletal muscle proteostasis and mTORC1 activity. Insulin signalling as well as growth factor signalling represses Foxo1 and activates mTORC1 activity while energy deprivation conditions such as starvation do the opposite.26 Interestingly, we observed an up-regulation of insulin receptor (IR) expression in the vdr−/− skeletal muscles compared with WT muscles (Figure 3A). However, phosphorylation of IR at Tyr1345, which is induced by activation of IR, was reduced, suggesting lower IR pathway activity (Figure 3B and 3C). Moreover, the levels of the adapter protein IRS1 were significantly low in vdr−/− muscles, strongly indicating reduced insulin signalling (Figure 3B). The activating Ser473 phosphorylation of AKT/PKB, an effector of the IR pathway and an activator of mTORC1 complex, was also reduced in vdr−/− muscles compared with WT (Figure 3D and 3E). In agreement with this, similar to the mice with mutations in the VDR DNA binding domain,27 7-week-old vdr−/− mice displayed defective insulin response following intraperitoneal glucose injection (Figure 3F). So, it is likely that the increased IR protein levels could be a compensatory mechanism to counteract the absence of insulin response, but this increase could not restore the IR pathway.
vdr−/− skeletal muscles exhibit energy deprivation. (A) Representative western blot for insulin receptor in 3- and 7-week-old WT and vdr−/−. (B) Western blots for phospho-IR (Tyr1345), IR, and IRS1. (C) Ratio of phospho-IR to total IR (n = 4). (D and E) Western blot for p-AKT in 3- and 7-week-old WT and vdr−/− (D) and quantitation of bands (E) (n = 3). (F) Serum insulin levels measured by ELISA at fasting state and 10 min after intraperitoneal injection of glucose (n = 5). (G–J) Representative western blots for LKB1 and AMPK (G, I) and their densitometric quantification (H, J) (n = 3). (K) Muscle ATP levels in WT and vdr−/−. All graphs show mean ± SEM. Statistical significance was determined by unpaired T-test (*: P < 0.05, **: P < 0.01, ***: P < 0.001, ****: P < 0.0001).
Conditions that lead to energy deprivation activate AMPK pathway and increase IR expression levels,28 and it is well established that the AMPK pathway inhibits the mTORC1 pathway.29, 30 Accordingly, protein levels of LKB1, the upstream activator of AMPK pathway, were significantly higher in the skeletal muscles of vdr−/− mice compared with WT (Figure 3G and 3H). Although total AMPK protein levels did not show any difference, there was a sharp increase in the levels of Thr172 phosphorylated AMPK in vdr−/− muscles (Figure 3I and 3J), indicating activation of the AMPK pathway. Indeed, ATP levels were significantly lower in vdr−/− muscles than WT, clearly indicating energy deprivation (0.58 ± 0.18 μmol/mL vs. 1.6 ± 0.0.14 μmol/mL, P = 0.006) (Figure 3K). Consistent with a systemic metabolic imbalance, we observed lower p-S6 levels in vdr−/− livers, indicative of reduced mTORC1 activity (Figure S3a). However, AMPK activity in the vdr−/− liver was comparable with that of WT (Figure S3b). Additionally, 7-week-old vdr−/− mice did not exhibit any difference in liver fasting protein synthesis, suggesting that protein synthesis in skeletal muscles is more susceptible to energy imbalances in the vdr−/− mice (Figure S3c).
Energy insufficiency in vdr−/− skeletal muscles is not due to diminished mitochondrial activity.
Decreased ATP levels in vdr−/− skeletal muscles may indicate defective mitochondrial biogenesis. On the other hand, energy deficiency conditions that lead to activation of the AMPK pathway are well established to increase mitochondrial biogenesis to meet the energy demands.31 To understand the underlying reason for the energy deprivation in vdr−/− skeletal muscles, it is essential to delineate these two scenarios. We postulated that if the energy deprivation is due to mitochondrial biogenesis defect, there should be a decrease in mitochondrial protein content. To address this, we performed western blot analysis of different classes of mitochondrial proteins in WT and vdr−/− mice. TOMM20, a central component of the mitochondrial protein import machinery located on the outer membrane, was unchanged in the vdr−/− mice (Figures 4A and S4a). TOMM20 is considered as a marker for mitochondrial mass,32 and this datum indicates that despite having lower energy levels, vdr−/− muscles maintain mitochondrial mass comparable with that of WT. To have a better understanding about the functional capacity of the mitochondria in vdr−/− muscles, we checked the levels of different proteins involved in oxidative phosphorylation. UQCRC1, a member of the complex III of mitochondrial inner membrane, and SDHA, a subunit of complex II, did not show any statistically significant change in their protein levels (Figures 4A, S4b, and S4c). On the other hand, COX4, a complex IV subunit, was up-regulated in vdr−/− (Figures 4A and S4d). Western analysis using a combination of antibodies against different components of the ETC showed a decrease in the levels of SDHB in vdr−/− but not in subunits of other complexes (Figure 4B). Interestingly, reduced levels of SDHA, but not SDHB, are known to be associated with reduced OXPHOS.33 Apart from the mitochondrial proteins imported from cytoplasm, the mitochondrial genome expresses 13 proteins that are essential functional components of all ETC and ATP synthase complexes.34 We analysed the levels of MT-ND1, which is part of the complex I and found to be unchanged in vdr−/− muscles (Figures 4C and S4e).
Mitochondrial function is also modulated by fission and fusion of the mitochondria.35 Western blot analyses of the fission protein Drp1 as well as the inner and outer membrane fusion proteins Opa1 and Mfn2 show that Mfn2 levels were slightly decreased in vdr−/− while others were unchanged, suggesting that there could be functional differences in the mitochondria of these mice (Figure 4D). To address this, we analysed the activity of respiratory complex II (SDH) in GAS muscles of these mice. Surprisingly, we found that the activity of SDH is significantly high in the vdr−/− mice, indicating increased mitochondrial activity (Figure 4E). Taken together, the data shown here suggest that although the vdr−/− skeletal muscles exhibit reduced ATP levels, it is not due to reduced mitochondrial activity. On the contrary, evidence suggests that these muscles may have increased mitochondrial activity. We did not find any significant differences in the levels of mitochondrial uncoupling proteins 2 and 3 (UCP2 and UCP3) in vdr−/− skeletal muscles, indicating that the discrepancy in mitochondrial activity and ATP levels is not due to the proton leakage induced by these proteins (Figure 4F and 4G). Interestingly, prolonged fasting, which leads to skeletal muscle wasting, is reported to result in increased mitochondrial activity in skeletal muscles.36 This raises the likelihood that the observed defects could be due to deficiency in nutrient utilization rather than defective mitochondrial biogenesis or function.
vdr−/− mice exhibit defective glucose homeostasisSkeletal muscle is the major site of glucose disposal in mammals, and insulin resistance in skeletal muscles accounts for 85–90% of glucose clearance impairment in type 2 diabetes.37, 38 The lack of insulin response in these mice in combination with reduced IR signalling in muscle suggests defective glucose clearance, which is expected to cause hyperglycaemia. But, on the contrary, fasting blood glucose levels of vdr−/− mice were significantly lower compared with WT at both 5 weeks (138.18 ± 5.02 mg/dL vs. 78.5 ± 7.69 mg/dL, P = 0.000003) and 7 weeks (95 ± 14.5 mg/dL vs. 148.6 ± 6.1 mg/dL, P = 0.0017) of age (Figure 5A and 5B). Random glucose levels at 7 weeks of age were also found to be lower in vdr−/− (Figure S5a).
vdr−/− mice exhibit defective glucose homeostasis. (A and B) Fasting glucose levels of 5- (A) and 7-week-old (B) mice (n ≥ 6). (C and D) Glucose tolerance test in 5- (C) and 7-week-old mice (D) (n > 6). (E) Hindlimb Glut1 mRNA levels quantified using qRT–PCR (n = 4). (F) Glucose uptake by GAS muscle as measured by the levels of fluorescence of NBDG in muscle (n = 3). (G) Serum lactate levels in WT and vdr−/− mice (n = 5). (H) Representative western blot showing the levels of PDK4 in WT and vdr−/− muscles. (I) PDH activity assay done on GAS muscles of vdr−/− and WT mice (n = 3). (J) Images of SDH stained WT and vdr−/− muscle sections. All graphs show mean ± SEM. Statistical significance was determined by unpaired T-test (*: P < 0.05, **: P < 0.01, ***: P < 0.001, ****: P < 0.0001).
A glucose tolerance test did not show any delay in glucose clearance in vdr−/− mice compared with WT at both 5 and 7 weeks of age (Figure 5C and 5D). The glucose levels remained low for vdr−/− even after glucose administration, but that is likely to be a reflection of lower baseline glucose levels. However, normalization of the values with respect to fasting glucose levels suggested that the kinetics of glucose uptake could be different in these mice (Figure S5b and S5c). Glut4 is the major glucose transporter in response to insulin, but during energy deprivation conditions, AMPK mediated up-regulation of Glut1 transporter accounts for increased glucose uptake independent of insulin.39 qRT–PCR analysis revealed a 10-fold up-regulation of the Glut1 mRNA in the vdr−/− muscle (Figure 5E). Indeed, there was an up-regulation of glucose uptake by vdr−/− skeletal muscles as indicated by the higher accumulation of NBDG, a fluorescent analogue of glucose (Figure 5F). Taken together, these data show that there is indeed a systemic energy deprivation in vdr−/− mice, leading to hypoglycaemia, but glucose disposal in these mice are not dramatically affected. Serum lactate levels were also high in vdr−/− mice (7.42 ± 0.31 nmol/μL vs. 4.95 ± 0.44 nmol/μL, P = 0.0032), further confirming a systemic defect in energy metabolism in vdr −/− mice (Figure 5G).
The above-described observations suggested that these muscles use glucose at a higher rate through aerobic glycolysis. We found that vdr−/− muscles have higher levels of PDK4 (Figure 5H), which inhibits pyruvate dehydrogenase (PDH) complex to divert the pyruvate flux to aerobic glycolysis. But, PDH activity of vdr−/− muscles did not vary significantly in vdr−/− (6.91 ± 0.19 mU/mL for WT vs. 9.43 ± 2.52 mU/mL for vdr−/−; P = 0.23) (Figure 5I). Our previous results show increased activity of mitochondrial complexes in vdr−/−, indicating increased mitochondrial function (Figure 4E). Indeed, SDH staining of GAS muscle cryosections clearly shows increased SDH activity in the vdr−/− muscles (Figure 5J). The increased mitochondrial activity argues against oxidative-to-glycolytic metabolic shift in vdr−/− muscle fibres. Instead, we propose that the unchanged PDH and increased SDH activities indicate a compensatory mechanism in vdr−/− muscles to increase energy production, which is known to occur during energy deprivation conditions such as starvation.36, 40-42 The energy deprivation in these muscles despite having increased mitochondrial activity, glucose uptake, and normal levels of uncoupling proteins suggested that these muscles may have a defect in mobilizing stored glucose.
Increased muscle glycogen accumulation is associated with defective glucose homeostasis in vdr−/− miceMajority of glucose injected is disposed in skeletal muscles, and it is intriguing that despite normal glucose clearance, there is an energy deficit in vdr−/− skeletal muscles. So, we asked if there is any deficiency in utilization of glucose in these muscles. Glucose taken up by skeletal muscles is stored as glycogen. We checked whether the energy deprivation in skeletal muscles is associated with differences in glycogen levels. PAS staining of TA muscle sections revealed higher levels of glycogen in vdr−/− compared with WT (Figure 6A). To address whether this increase is associated with an increase in glycogen synthesis, we estimated glycogen synthase (GYS1) and phospho-GYS1 levels in the two groups and found a profound reduction in the inhibitory Ser641 phosphorylation of GYS1 (Figure 6B and 6C). Furthermore, consistent with reduced glycogen utilization, the activating Ser15 phosphorylation of glycogen phosphorylase (PYGM), an enzyme that degrades glycogen, was significantly reduced in vdr−/− (Figure 6D and 6E). Both the glycogen synthase and glycogen phosphorylase are regulated by calcium, and as expected, vdr−/− mice exhibit deficiency in serum calcium levels (Figure 6F). Moreover, the mRNA levels of glycogenin, an enzyme required to initiate glycogen biosynthesis, were up-regulated in vdr−/− muscles (Figure 6G), supporting increased glycogen synthesis. Interestingly, we also found that the mRNA levels of glycogen branching enzyme (GBE1), whose depletion leads to accumulation of abnormal glycogen, were down-regulated (Figure 6H). Taken together, these results suggest that skeletal muscles of vdr−/− mice exhibit a carbohydrate utilization defect characterized by increased glycogen accumulation. Skeletal muscle glycogen storage disorders are known to induce systemic and skeletal muscle energy imbalance and trigger atrophy.43
Dysregulation of glycogen synthase and phosphorylase activities lead to increased glycogen accumulation in vdr−/− skeletal muscles. (A) Micrographs of PAS stained transverse sections of GAS muscles from WT and vdr−/− mice. (B and C) Western blot (B) and densitometric quantitation (C) for GYS1 (n = 3). (D and E) Western blot (D) and densitometric quantitation (E) of PYGM (n = 3). (F) Serum calcium levels in WT and vdr−/− mice (n = 4). (G and H) qRT–PCR quantitation of glycogenin (G) and GBE1 (H) mRNAs in TA muscles (n = 5). All graphs show mean ± SEM. Statistical significance was determined by unpaired T-test (*: P < 0.05, **: P < 0.01, ***: P < 0.001, ****: P < 0.0001).
DiscussionSeveral factors are known to lead to skeletal muscle atrophy, ranging from inactivity and starvation to genetic disorders and other diseases such as diabetes, cancer, and tuberculosis.10, 21 Although the vdr−/− mice are known to exhibit severe skeletal muscle atrophy,1 and vitamin D-deficient individuals exhibit varying degrees of myopathies,44 the underlying mechanisms are not well understood. Because these mice exhibit a growth defect as well, it is likely that the reduced muscle mass could be a manifestation of reduced growth than muscle wasting. But our results as well as others have shown that skeletal muscles of these mice exhibit increased expression of atrophy markers such as MurF1, Fbxo32, and the Foxo1 transcription factor.15, 45 Moreover, our data show that the muscle proteostasis defect is not observed in 3-week-old mice. Increased muscle protein degradation has been reported in vitamin D-deficient rat models as well.12 Further granularity of proteostasis defects was achieved by identifying that the decrease in protein synthesis is limited to the fasting state in vdr−/− mice while absorptive state protein synthesis was not affected. We propose that the availability of nutrients in the postprandial state is responsible for maintaining normal protein synthesis rates while energy deprivation associated with impaired glycogen utilization is responsible for decreased protein synthesis during the fasting state in vdr−/− skeletal muscles.
Based on the evidence presented here, we propose the following model for the development of skeletal muscle atrophy in vdr−/− mice (Figure 7). The abnormal accumulation of glycogen in vdr−/− muscles, driven by increased glycogen synthase activity and decreased glycogen phosphorylase activity, renders it unavailable for energy production during the post-absorptive state leading to a state of constitutive energy deprivation and to the activation of the AMPK pathway. High AMPK activity in turn leads to higher glucose uptake by muscle with the aid of increased Glut1 expression,39 and its chronic activation contributes to increased glycogen accumulation in skeletal muscles.46 This feed forward loop of glucose uptake and glycogen synthesis with a reduction in the glycogen phosphorylase activity could elevate glucose consumption by vdr−/− muscles and subsequent systemic glucose depletion especially during post-absorptive state. The increased serum lactate levels, which could be the result of the body's attempt to increase the liver gluconeogenesis in vdr−/− mice, support this hypothesis. The increased muscle proteolysis observed in vdr−/− could also function to ensure supply of other gluconeogenesis substrates such as alanine. However, this seems insufficient in maintaining fasting glucose levels in vdr−/− mice. The defects in liver metabolism require to be studied further to resolve this issue. The increase in the mitochondrial activity observed in the muscles of these mice indicates that the energy deficiency is not driven by mitochondrial damage. The increased mitochondrial activity may reflect the attempt by the muscles to counterbalance the energy insufficiency. These data clearly suggest that the energy deprivation induced by the glycogen storage defect is causative for the muscle wasting in vdr−/− mice.
Glycogen storage disorder leads to skeletal muscle atrophy in vdr−/− mice. Dysregulation of glycogen synthase and phosphorylase in the absence of VDR signalling leads to increased glycogen accumulation in skeletal muscles and subsequent energy deprivation and glucose homeostasis defects. These defects lead to increased protein degradation and decreased protein synthesis, causing skeletal muscle atrophy.
This model could be applicable to humans as well. Interestingly, skeletal muscles of athletes with severe vitamin D-deficient have been reported to show increased glycogen granules.44 Moreover, the susceptibility of individuals with vitamin D deficiency to develop insulin resistance is also well documented, showing the relationship between vitamin D signalling and glucose homeostasis.
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