Diet-induced vitamin D deficiency reduces skeletal muscle mitochondrial respiration

in Journal of Endocrinology
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  • 1 School of Sport, Exercise and Rehabilitation Sciences, University of Birmingham, Birmingham, UK
  • | 2 MRC-Versus Arthritis Centre for Musculoskeletal Ageing Research, Clinical, Metabolic and Molecular Physiology, University of Nottingham, Royal Derby Hospital Centre, Derby, UK
  • | 3 Mitochondrial Metabolism and Ageing Laboratory, Garvan Institute of Medical Research, Sydney, New South Wales, Australia
  • | 4 St Vincent’s Clinical School, UNSW Medicine, UNSW Sydney, New South Wales, Australia
  • | 5 Institute of Metabolism and Systems Research, University of Birmingham, Birmingham, UK
  • | 6 ANZAC Research Institute, University of Sydney, Sydney, New South Wales, Australia
  • | 7 Centenary Institute, Royal Prince Alfred Hospital, Camperdown, New South Wales, Australia
  • | 8 UTS Centenary Centre for Inflammation, University Technology Sydney, New South Wales, Australia

Correspondence should be addressed to A Philp: a.philp@garvan.org.au
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Vitamin D deficiency is associated with symptoms of skeletal muscle myopathy including muscle weakness and fatigue. Recently, vitamin D-related metabolites have been linked to the maintenance of mitochondrial function within skeletal muscle. However, current evidence is limited to in vitro models and the effects of diet-induced vitamin D deficiency upon skeletal muscle mitochondrial function in vivo have received little attention. In order to examine the role of vitamin D in the maintenance of mitochondrial function in vivo, we utilised an established model of diet-induced vitamin D deficiency in C57BL/6J mice. Mice were either fed a control diet (2200 IU/kg i.e. vitamin D replete) or a vitamin D-deplete (0 IU/kg) diet for periods of 1, 2 and 3 months. Gastrocnemius muscle mitochondrial function and ADP sensitivity were assessed via high-resolution respirometry and mitochondrial protein content via immunoblotting. As a result of 3 months of diet-induced vitamin D deficiency, respiration supported via complex I + II (CI + IIP) and the electron transport chain (ETC) were 35 and 37% lower when compared to vitamin D-replete mice (P < 0.05). Despite functional alterations, citrate synthase activity, AMPK phosphorylation, mitofilin, OPA1 and ETC subunit protein content remained unchanged in response to dietary intervention (P > 0.05). In conclusion, we report that 3 months of diet-induced vitamin D deficiency reduced skeletal muscle mitochondrial respiration in C57BL/6J mice. Our data, when combined with previous in vitro observations, suggest that vitamin D-mediated regulation of mitochondrial function may underlie the exacerbated muscle fatigue and performance deficits observed during vitamin D deficiency.

Abstract

Vitamin D deficiency is associated with symptoms of skeletal muscle myopathy including muscle weakness and fatigue. Recently, vitamin D-related metabolites have been linked to the maintenance of mitochondrial function within skeletal muscle. However, current evidence is limited to in vitro models and the effects of diet-induced vitamin D deficiency upon skeletal muscle mitochondrial function in vivo have received little attention. In order to examine the role of vitamin D in the maintenance of mitochondrial function in vivo, we utilised an established model of diet-induced vitamin D deficiency in C57BL/6J mice. Mice were either fed a control diet (2200 IU/kg i.e. vitamin D replete) or a vitamin D-deplete (0 IU/kg) diet for periods of 1, 2 and 3 months. Gastrocnemius muscle mitochondrial function and ADP sensitivity were assessed via high-resolution respirometry and mitochondrial protein content via immunoblotting. As a result of 3 months of diet-induced vitamin D deficiency, respiration supported via complex I + II (CI + IIP) and the electron transport chain (ETC) were 35 and 37% lower when compared to vitamin D-replete mice (P < 0.05). Despite functional alterations, citrate synthase activity, AMPK phosphorylation, mitofilin, OPA1 and ETC subunit protein content remained unchanged in response to dietary intervention (P > 0.05). In conclusion, we report that 3 months of diet-induced vitamin D deficiency reduced skeletal muscle mitochondrial respiration in C57BL/6J mice. Our data, when combined with previous in vitro observations, suggest that vitamin D-mediated regulation of mitochondrial function may underlie the exacerbated muscle fatigue and performance deficits observed during vitamin D deficiency.

Introduction

Vitamin D deficiency, characterised by serum 25-hydroxyvitamin D (25(OH)D) levels of < 50 nmol/L, remains prevalent across both Europe and the United States of America (Forrest & Stuhldreher 2011, Cashman et al. 2016). Although the classical actions of vitamin D in the maintenance of bone health are well established (Ham & Lewis 1934, Rizzoli et al. 2008, Bhan et al. 2010), a number of non-classical actions have recently been identified including the maintenance of skeletal muscle function (Girgis et al. 2013).

Within human populations, multiple observational studies have reported a positive association between serum 25(OH)D, skeletal muscle strength and lower extremity function in older individuals (Bischoff-Ferrari et al. 2004, Wicherts et al. 2007, Houston et al. 2012). Furthermore, the supplementation of vitamin D has been reported to increase muscle strength within this population (Stockton et al. 2011, Beaudart et al. 2014). Despite these associations, studies of this design are unable to infer causality. In addition, isolating the effects of vitamin D status within older populations is often difficult given that individuals may suffer from a number of pre-existing conditions that may interfere with the vitamin D status (Duncan et al. 2012). These difficulties highlight the importance of model systems that allow for the manipulation and isolation of vitamin D status in order to study the precise role of vitamin D within skeletal muscle.

In order to study the impact of vitamin D deficiency on skeletal muscle function, a number of animal models have been utilised. A dysregulation of vitamin D status can be achieved via dietary means (Rodman & Baker 1978, Pleasure et al. 1979, Pointon et al. 1979, Girgis et al. 2015), a reduction in sunlight exposure (Pleasure et al. 1979) or by the administration of ethane-1-hydroxy-1,1-diphosphonate which blocks the production of 1α,25-dihydroxyvitamin D3 (1α,25(OH)2D3) (Pointon et al. 1979). Diet-induced vitamin D deficiency has been shown to result in symptoms of skeletal muscle myopathy including impaired contraction kinetics, skeletal muscle weakness as well as decrease in muscle force in both chicks and rats (Rodman & Baker 1978, Pleasure et al. 1979, Schubert & DeLuca 2010). In order to isolate the effects of vitamin D alone and offset the observed hypocalcaemia and hypophosphataemia that are associated with the induction of vitamin D deficiency (Schubert & DeLuca 2010), diets with increased calcium and phosphate content have been utilised (Girgis et al. 2015). However, despite the administration of this rescue diet, mice still display reduced grip strength and an increase in Myostatin gene expression (Girgis et al. 2015), a known negative regulator of muscle mass (McPherron et al. 1997). Similarly, mice fed with this diet chronically (8–12 months) show similar impairments in physical performance including reduced grip endurance, sprint speed and stride length (Seldeen et al. 2018).

The observed impairments in physical performance with vitamin D deficiency may be linked to skeletal muscle mitochondrial function (Bouillon & Verstuyf 2013, Sinha et al. 2013). In vitro, vitamin D-related metabolites are able to increase mitochondrial function in both immortalised and primary skeletal muscle cell lines (Ryan et al. 2016, 2018, Schnell et al. 2019, Romeu Montenegro et al. 2019). Furthermore, we recently observed significant impairments in mitochondrial function in vitamin D receptor (VDR) loss-of-function C2C12 myoblasts (Ashcroft et al. 2020). In humans, the supplementation of vitamin D within a cohort of severely deficient individuals resulted in a reduced phosphocreatine (PCr) recovery time, as measured non-invasively by 31-phosphorous magnetic resonance spectroscopy (31-P MRS) (Sinha et al. 2013). Whilst skeletal muscle mitochondrial content seems to remain unchanged following diet-induced vitamin D deficiency in mice (Seldeen et al. 2018), we have also observed a downregulation of mitochondrial genesets follow VDR knock down in rat skeletal muscle (Bass et al. 2020). Despite these observations, the functional characteristics of the mitochondria remain largely underexplored in vivo. Therefore, we aimed to determine the effects of diet-induced vitamin D deficiency upon skeletal muscle mitochondrial function in C57BL/6J mice.

Methods

Ethical approval

Ethical approval was granted by the Garvan Institute and St. Vincent’s Hospital Animal Experimentation Ethics Committee (approval number 18/19), fulfiling the requirements of the NHMRC and the NSW State Government, Australia. All animal handling was carried out by trained personnel and all procedures were carried out according to the Australian code of practice for the care and use of animals for scientific purposes 8th edition (Health et al. 2013). Male C57BL/6JAusb mice were received at 10 weeks of age and housed communally in a temperature-controlled environment (22 ± 0.5°C) with a 12 h light:12 h darkness cycle.

Composition of diet

Following 1-week acclimation in which mice were fed a standard chow diet, mice were placed on either a vitamin D-control diet (SF085-034, Speciality Feeds, Glen Forest, NSW) or a vitamin D-deplete diet (SF085-003, Speciality Feeds, Glen Forest, NSW) for periods of 1 (n = 10/group), 2 (n = 6/group) or 3 months (n = 6/group). The vitamin D-deplete diet contains no vitamin D (cholecalciferol 0 IU/kg) but increased calcium (2%) and phosphorous (1.2%) content in order to maintain normal mineral homeostasis. Previously, this dietary intervention has been shown to successfully induce vitamin D deficiency following 1 month of dietary intervention (Girgis et al. 2015). The vitamin D-control diet contains vitamin D (cholecalciferol 2200 IU/kg), calcium (1%) and phosphorous (0.7%).

Assessment of food intake

Food intake was assessed on a monthly basis at 1, 2 and 3 months of dietary intervention. The weight of the food within the cage was recorded and subsequently re-weighed following a period of 24 h. The amount of food consumed was then divided by the number of mice within the cage and reported as food intake in grams per mouse.

Assessment of body composition

Body weight was obtained on a weekly basis throughout the dietary intervention periods. In addition, prior to each measurement of body composition, mice were briefly weighed. Body composition was assessed upon arrival (10 weeks of age) and then following 1, 2 and 3 months of dietary intervention using the EchoMRI (EchoMRI LLC, Houston, USA).

Tissue collection

Tissue collections were completed following 1, 2 and 3 months of dietary intervention. All samples were excised from fasted (2 h) mice following isoflurane (5%) anaesthetisation. Following collection, a blood sample was taken via cardiac puncture and animal was terminated via cervical dislocation. All tissues were rinsed in sterile saline, blotted dry, weighed, and frozen in liquid nitrogen. A small portion (∼20 mg) of the red gastrocnemius was removed before freezing and used for high-resolution respirometry. All further tissue samples were stored at −80°C for subsequent analysis.

Tissue processing

Small portions of red gastrocnemius muscle (∼20 mg) were removed and placed in ice-cold BIOPS buffer (2.77 mM CaK2EGTA, 7.23 mM K2EGTA, 5.77 mM Na2ATP, 6.56 mM MgCl2-6H2O, 20 mM taurine, 15 mM Na2-phosphocreatine, 20 mM imidazole, 0.5 mM dithiothreitol, 50 mM MES hydrate, pH 7.1, 290 mOsm). Blood samples were allowed to coagulate at room temperature for 10 min before being placed on ice. Blood samples were then centrifuged at 14,000 g for 10 min and the resulting supernatant was removed and stored at −80°C for further analysis.

Analysis of serum vitamin D and calcium

Serum concentrations of vitamin D metabolites were analysed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) as previously described (Jenkinson et al. 2016). Briefly, serum was extracted prior to analysis by protein precipitation followed by supportive liquid–liquid extraction. Analysis was performed on a SCIEX 6500 coupled 191 to a Shimadzu Exion liquid chromatography mass system. Analysis was carried out in multiple reaction monitoring (MRM) for the following analytes: 25(OH)D3, 3-epi-25-hydroxyvitamin D3 (3-epi-25(OH)D3), 24,25-dihydroxyvitamin D3 (24,25(OH)2D3), 20,24-dihydroxyvitamin2D3 (20,24(OH)2D3) and 1α,25(OH)2D3. The LC-MS/MS method for vitamin D quantification was validated for serum analysis as previously described for accuracy, precision, recovery and matrix effects (Jenkinson et al. 2016). Vitamin D metabolites were purchased from Supleco Sigma-Aldrich. LC-MS grade methanol and water were purchased from Thermo Fisher and Chem Supply, respectively. Supportive liquid–liquid extraction plates were purchased from Phenomenex. Serum calcium was measured using a Calcium Detection Assay kit (Abcam). Serum samples were diluted 1:10 and manufacturer's instructions were followed. The assay plate was read at 575 nm using a CLARIOstar microplate reader (BMG Labtech, Victoria, Australia).

High-resolution respirometry

High-resolution respirometry was conducted in MiR05 (2 mL) with the addition of blebbistatin (25 μM) using the OROBOROS Oxygraph-2k (Oroboros Instruments, Corp., Innsbruck, AT) with stirring at 750 rpm at 37°C. Oxygen within the chamber was maintained between 150 and 220 μM for each experiment. Prior to the addition of the fibre bundles to the chamber, bundles were blotted dry and weighed. Bundles totalling 2.5–5.0 mg were added to each chamber. First, pyruvate (10 mM) and malate (2 mM) were added in the assessment of complex I-related leak (CIL). ADP was then titrated in step-wise increments (100–6000 μM) followed by the addition of glutamate (10 mM) to assess phosphorylating respiration (CIP). The addition of succinate (10 mM) followed to assess respiration support via complex II (CI + IIP). Cytochrome c (cyt c) (10 μM) was added in order to check outer mitochondrial membrane integrity. The partial loss of cyt c during fibre preparation may limit respiration, however, no fibre preparation exhibited an increase of > 10%. Carbonyl cyanide 3-chlorophenylhydrazone (CCCP) was titrated in a step-wise manner (0.5 to 2.5 μM) until the maximal capacity of the electron transport chain (ETC) was reached. Finally, antimycin A (2.5 μM) was injected in order to determine non-mitochondrial oxygen consumption.

The apparent Km for ADP was determined through the Michaelis–Menten enzyme kinetics – fitting model (Y = Vmax × X/(Km + X)), where X = (free ADP; ADPf), using Prism (GraphPad Software, Inc.) as previously described (Perry et al. 2011). Flux control ratios (FCR) was calculated by setting CCCP stimulated respiration as 1 and antimycin A respiration as 0.

Citrate synthase activity

Gastrocnemius samples were homogenised in a ten-fold mass of ice-cold sucrose homogenisation buffer via shaking in a FastPrep 24 5G (MP Biomedicals, Santa Ana, California, USA) at 6.0 m/s for 80 s as previously described (Stocks et al. 2017). Protein concentrations were determined via a DC protein assay (Bio-Rad) and an equal volume of protein (10 μg) was loaded onto a 96-well microtiter plate in triplicate. Ten microlitres of sample was diluted in 235 μL of reaction buffer (64 mM tris pH 8.0, 0.13 mM 5,5-dithio-bis-(2-nitrobenzoic acid), 0.13 mM acetyl CoA). Five microlitres of 5 mM oxaloacetate was added to start the reaction and absorbance was read at 412 nm for 3 min in a CLARIOstar microplate reader (BMG Labtech, Victoria, Australia). Citrate synthase activity in nmol/min/mgwas determined from absorbance values and corrected for differences in path length (Spinazzi et al. 2012, Stocks et al. 2017).

Immunoblotting

Gastrocnemius samples were powdered on dry ice using a Cellcrusher™ tissue pulverizer (Cellcrusher Ltd, Cork, Ireland) and homogenised via shaking in a FastPrep 24 5G (MP Biochemicals, Santa Ana, California, USA) at 6.0 m/s for 80 s in a ten-fold mass of ice-cold sucrose lysis buffer (50 mM tris pH 7.5; 270 mM sucrose; 1 mM EDTA; 1 mM EGTA; 1% triton X-100; 50 mM sodium fluoride; 5 mM sodium pyrophosphate decahydrate; 25 mM beta-glytcerolphosphate). Inhibitors were added fresh on the day of use and included one cOmplete™ protease inhibitor cocktail EDTA free tablet (Roche) and Phosphatase Inhibitor Cocktail 3 both purchased from Sigma-Aldrich). Samples were then centrifuged for 10 min at 8000 g at 4°C to remove any insoluble material. Protein concentrations were determined using the DC protein assay as per manufacturer’s instructions (Bio-Rad). An equal volume of protein (30 μg) was separated by SDS-PAGE on 12.5% gels at a constant current of 23 mA per gel for ∼60 min. Proteins were then transferred on to BioTrace NT nitrocellulose membranes (Pall Life Sciences, Pensacola, Florida, USA) using a wet transfer system at 100 V for 1 h. Membranes were then stained in Ponceau S (Sigma-Aldrich) and imaged to check for even loading and transfer. Membranes were then blocked for 1 h in 3% dry-milk in tris-buffered saline with tween (TBS-T). Membranes were incubated overnight in primary antibodies at 4°C. Following primary antibody incubation, membranes were washed three times in TBS-T and subsequently incubated in the appropriate horseradish peroxidase-conjugated secondary antibody at room temperature for 1 h. Membranes were again washed three times in TBS-T prior to imaging. Images were captured using the ChemiDoc (Bio-Rad) and quantified using ImageJ.

Antibodies

All primary antibodies were used at a concentration of 1:1000 in TBS-T. MitoProfile OXPHOS antibody cocktail (110413) and mitofilin (110329) were purchased from Abcam. Total AMPKα (2603), phospho-AMPKThr172 (2535) and OPA1 (80471) were purchased from Cell Signaling Technology. Anti-mouse (7076) and anti-rabbit (7074) secondary antibodies were used at a concentration of 1:10,000 in TBS-T and were from Cell Signaling Technology.

Statistical analyses

Statistical analyses were performed using Prism version 7 (GraphPad Software Incorporated). Differences between 1, 2 and 3 month vitamin D-replete and -deplete mice were determined by two-way ANOVA with Bonferroni correction for multiple comparisons. Differences between vitamin D-deplete and -replete mice in mitochondrial respiration in response to ADP titration were determined by multiple t-test and corrected for multiple comparisons via Bonferroni method. Differences between vitamin D-replete and vitamin D-deplete groups in citrate synthase activity and vitamin D metabolite concentrations at the 3-month time point were determined via independent t-tests. All values are presented as mean ± s.d. Statistical significance was set at P < 0.05.

Results

Body composition following diet-induced vitamin D deficiency

Body weight increased across time when mice were compared at the 1-, 2- and 3-month time points (main effect of time, P < 0.001) (Fig. 1A). However, no statistically significant differences in body weight were observed when comparing vitamin D-replete and -deplete mice (P > 0.05) (Fig. 1A). Further assessment of body composition revealed no significant differences in absolute lean mass (P > 0.05) (Fig. 1C). However, when expressed as a percentage of body weight, lean mass was lower when compared across dietary intervention time points (group × time interaction, P < 0.001) (Fig. 1D). In particular, lean mass as a percentage of body weight in vitamin D-replete mice remained stable across the dietary intervention; however, it became gradually lower in vitamin D-deplete mice. For example, lean mass as a percentage of body weight was 16% lower at 3 months when vitamin D-deplete mice were compared with 1-month vitamin D deplete (P < 0.001) (Fig. 1D). Although this is indicative of a loss of lean mass with vitamin D deficiency, this effect was potentially driven by a higher lean mass as a percentage of body weight at the 1-month time point in vitamin D-deplete mice (P = 0.039) (Fig. 1D). Both absolute and percentage of body weight fat mass increased across time in both dietary groups (main effect of time, P < 0.001) (Fig. 1E and F). Despite differences in body composition, no statistically significant differences in food intake were observed between time points (P > 0.05) or dietary intervention (P > 0.05) (Fig. 1B).

Figure 1
Figure 1

Body composition and food intake in vitamin D-replete and -deplete C57BL/6J mice. (A) Increase in body weight across dietary period with no differences between differing vitamin D diets (n = 6–10/group). (B) No difference in food intake was observed following dietary intervention (n = 5–6 cages/group). (C and D) Absolute lean mass remained unchanged whilst lean mass as a percentage of body weight was significantly lower in the 3-month vitamin D-deplete mice when compared to the 1-month deplete mice (n = 6–10/group). (E and F) Absolute and percentage of body weight fat mass increased across the dietary period irrespective of dietary intervention (n = 6–10/group). Data mean ± s.d. bMain effect for time (P < 0.05), cGroup × time interaction (P < 0.05), eSignificantly different from 1-month time point of the same group (P < 0.05), *(P < 0.05).

Citation: Journal of Endocrinology 249, 2; 10.1530/JOE-20-0233

Skeletal muscle mass

Given the alterations in total lean mass, we determined whether individual skeletal muscle mass was effected by diet-induced vitamin D deficiency. Crude analysis of skeletal muscle wet weight revealed no significant differences in the mass of the gastrocnemius in response to either dietary intervention or dietary time point (P > 0.05) (Fig. 2A). Simlarly, no significant changes in gastrocnemius weight were observed when normalised to body weight (P > 0.05) (Fig. 2B). The mass of the quadriceps increased over time (main effect of time, P = 0.004); however, this was not changed by dietary intervention (P = 0.951) (Fig. 2C). Similar changes were observed when normalised to body weight (main effect of time, P = 0.017) (Fig. 3D). Collectively, tricep mass was higher when vitamin D-replete mice were compared with deplete (main effect of dietary intervention, P = 0.041), although post-hoc analysis revealed no significant differences between groups at individual time points (P > 0.05) (Fig. 2E). When normalised to body weight, the mass of triceps were reduced across time (main effect of time, P = 0.017) (Fig. 2F).

Figure 2
Figure 2

Skeletal muscle mass in response to the induction of diet-induced vitamin D deficiency. (A) Gastrocnemius wet weight (milligrams) remains unchanged in response to vitamin D deficiency (n = 6–10/group). (B) Gastrocnemius wet weight normalised to body weight (milligrams/gram) remains unchanged in response to vitamin D defieceny (n = 6–10/group). (C) Higher quadricep wet weight (milligrams) across time but no influence of vitamin D deficiency (n = 6–10/group). (D) Higher body quadricep mass (milligrams/gram) when normalised to body weight across dietary time period with no effect of vitamin D deficiency (n = 6–10/group). (E) Lower tricep mass (milligrams) in reponse to diet-induced vitamin D deficiency (n = 6–10/group). (F) Higher tricep mass when normalised to body weight (milligrams/gram) across dietary time period (n = 6–10/group). Data mean ± s.d. aMain effect of dietary intervention (P < 0.05), bMain effect for time (P < 0.05).

Citation: Journal of Endocrinology 249, 2; 10.1530/JOE-20-0233

Figure 3
Figure 3

Analysis of serum vitamin D metabolites and calcium in response to the induction of diet-induced vitamin D deficiency. (A) Successful induction of vitamin D deficiency was observed via significant reductions in serum 25-hydroxyvitamin D3 (25(OH)D3), 3-epi-25-hydroxyvitamin D3 (3-epi-25(OH)D3), 24,25-dihydroxyvitamin D3 (24,25(OH)2D3), 20,24-dihydroxyvitamin2D3 (20,24(OH)2D3) and 1α,25-dihydroxyvitamin D3 (1α,25(OH)2D3) following 3 months of dietary intervention (n = 6/group). (B) No change in serum calcium following the 1, 2 and 3 months of diet-induced vitamin D deficiency in C57BL/6J mice (n = 5–10/group). Data mean ± s.d. *P < 0.05.

Citation: Journal of Endocrinology 249, 2; 10.1530/JOE-20-0233

Serum vitamin D and calcium

Similar to previous reports (Girgis et al. 2015), we observed a significant decrease in serum 25(OH)D3 in response to 3 months of dietary intervention (P < 0.0001) (Fig. 3A). Furthermore, we also observed significant decreases in multiple vitamin D-related metabolites following 3 months of dietary intervention including epi-25(OH)D3, 24,25(OH)2D3, 20,24(OH)2D3 as well as the active metabolite form of vitamin D, 1,25(OH)2D3, which was undectable in serum samples from vitamin D deplete mice (P < 0.005) (Fig. 3A). We also observed no significant change in serum calcium irrespective of dietary group or time point (P > 0.05) (Fig. 3B).

Mitochondrial respiration

In response to both pyruvate and malate alone, we observed no significant differences in CIL (P > 0.05) (Fig. 4A). However, signicificant effects were observed for CIP (group × time interaction; P = 0.035), CI + IIP (group × time interaction; P = 0.035) and ETC (group × time interaction; P = 0.017) supported respiration (Fig. 4B, C and D). Further analysis revealed CIP respiration was 85 and 96% higher in vitamin D-replete mice at the 2- (P = 0.015) and 3-month (P = 0.006) time points when compared with 1-month vitamin D-replete mice (Fig. 4B). Similarly, higher respiratory capacities during CI + IIP respiration and the maximal capacity of the ETC were observed in the 2- and 3-month vitamin D-replete mice when compared with the 1-month time point (P < 0.05) (Fig. 4C and D). It should also be noted that these increases across time were not apparent in the vitamin D-deplete mice (Fig. 4B, C and D). In addition, 3 months of diet-induced vitamin D deficiency resulted in 35 and 37% lower respiratory rates during CI + IIP (P = 0.035) and maximal ETC capacity (P = 0.015) when compared to vitamin D-replete mice at the same time point (Fig. 4C and D). We also analysed the above data as a flux control ratio whereby maximal and non-mitochondrial respiration are set as 1 and 0, respectivley. Such analysis provides a method for internal normalisation (Pesta & Gnaiger 2012, Canto & Garcia-Roves 2015). Despite the observed changes reported above, flux control ratios revealed no significant differences in mitochondrial respiration supported via CIP alone (P > 0.05) or CI + IIP in combination (P > 0.05) (Fig. 4E and F).

Figure 4
Figure 4

Assessment of skeletal muscle mitochondrial oxygen consumption (JO2) in response to diet-induced vitamin D deficiency. (A) Complex I-related leak (CIL) remains unchanged irrespective of the dietary intervention (n = 4–8/group). (B) Complex I phosphorylating (CIP) respiration is higher in the 2- and 3-month vitamin D-replete mice when compared with the 1-month replete mice (n = 6–8/group). (C and D) Phosphorylating respiration supported via complex I and II (CI + IIP) and the maximal capacity of the electron transport chain (ETC) is increased in the vitamin D-replete mice and is higher at the 3-month time point when compared to vitamin D-deplete mice (n = 6–8/group). (E and F) Alteration in absolute rates of respiration are diminished when internally normalised (n = 6–8/group). Data mean ± s.d.. bMain effect for time (P < 0.05), cGroup × time interaction (P < 0.05), eSignificantly different from 1-month time point of the same group (P < 0.05), *(P < 0.05).

Citation: Journal of Endocrinology 249, 2; 10.1530/JOE-20-0233

ADP sensitivity

Commonly, the assessment of mitochondrial function is performed under saturating concentrations of ADP (Pesta & Gnaiger 2012, Canto & Garcia-Roves 2015) which may not be biologically relevant. Therefore, we assessed mitochondrial function in response to a titration of ADP which is biologically relevant to saturating concentrations (Perry et al. 2011, Miotto et al. 2018). We observed no significant differences in mitochondrial respiration throughout the titration of ADP between vitamin D-replete and -deplete mice (P > 0.05) (Fig. 5A, B and C). Furthermore, no significant differences were observed in the apparent Km for ADP in response to either dietary intervention (P > 0.05) or time point (P > 0.05) (Fig. 5D).

Figure 5
Figure 5

Assessment of skeletal muscle mitochondrial ADP sensitivity in response to diet-induced vitamin D deficiency. (A, B and C) No change in respiratory capacity in response to the titration of ADP following 1, 2 and 3 month of vitamin D deficiency (n = 6–8/group). (D) No change in the apparent Km for ADP in response to 1, 2 and 3 month of vitamin D deficiency (n = 5–8/group). Data mean ± s.d.

Citation: Journal of Endocrinology 249, 2; 10.1530/JOE-20-0233

Mitochondrial protein content and enzyme activity

Finally, given the observed decrements in mitochondrial function associated with vitamin D deficiency, we sought to assess mitochondrial protein content following the diet-induced vitamin D deficiency. We observed no significant changes in complex I (NDUFB8), complex II (SDHB), complex IV (UQCRC2) irrespective of time point or vitamin D diet (P > 0.05) (Fig. 6A, B and D). Interestingly, both complex III (MTCO1) and complex V (ATP5A) were decreased across time (main effect of time, P < 0.05) and with vitamin D-deplete dietary intervention (main effect of dietary intervention, P < 0.05) (Fig. 6C and E). Furthermore, no significant changes were observed in citrate synthase activity (Fig. 7A), AMPKThr172 phosphorylation (Fig. 7B), mitofilin or OPA1 protein content (Fig. 7C and D) in response to dietary intervention (P > 0.05). Collectively this data would suggest that mitochondrial protein content remains unchanged through 3 months of vitamin D deficiency (P > 0.05).

Figure 6
Figure 6

Assessment of skeletal muscle ETC protein content following diet-induced vitamin D deficiency. (A, B and D) No changes were observed in expression levels of mitochondrial complex I (NDUFB8), complex II (s.d.HB), complex IV (UQCRC2), following diet-induced vitamin D deficiency (n = 6/group). (C and E) Lower expression levels of mitochondrial complex III (MTCO1) and complex V (ATP5A) across the dietary time period with no differences between vitamin D-replete and -deplete mice (n = 6/group). Data mean ± s.d. bMain effect for time (P < 0.05), cGroup × time interaction (P < 0.05).

Citation: Journal of Endocrinology 249, 2; 10.1530/JOE-20-0233

Figure 7
Figure 7

Assessment of skeletal muscle mitochondrial protein content and enzyme activity following diet-induced vitamin D deficiency. (A, B, C, D, F, G and H) No changes in citrate synthase activity, AMPKThr172 phosphorylation, mitofilin and OPA1 following diet-induced vitamin D deficiency (n = 6/group). Data mean ± s.d.

Citation: Journal of Endocrinology 249, 2; 10.1530/JOE-20-0233

Discussion

Vitamin D deficiency has been linked to reductions in skeletal muscle function; however, the specific role of vitamin D on mitochondrial function in less established. Therefore, the aim of the present study was to directly examine the effects of diet-induced vitamin D deficiency upon skeletal muscle mitochondrial function in C57BL/6J mice. Utilising the current gold-standard method to assess mitochondrial function in permeabilised skeletal muscle fibres (Lanza & Nair 2010, Picard et al. 2011, Cardinale et al. 2018), we report that 3 months of diet-induced vitamin D deficiency reduces mitochondrial respiration supported via CI + IIP and the maximal capacity of the ETC (Fig. 4C and D). Interestingly, despite the functional changes, we observed no differences in mitochondrial protein content following the induction of diet-induced vitamin D deficiency (Figs 6 and 7). In addition, 1 month of diet-induced vitamin D deficiency resulted in an increase in lean mass (Fig. 1D) as a percentage of body weight, although these effects were transient as they did not manifest over 2 and 3 months of dietary intervention. Furthermore, diet-induced vitamin D deficiency resulted in a decrease in lean mass as a percentage of body weight across the 3-month time period (Fig. 1D), whilst no changes in body weight, lean mass, fat mass or food intake were apparent when comparing vitamin D-replete group to the -deplete group following 3 months of dietary intervention (Fig. 1A, B, C, D, E and F).

The ability of vitamin D and related metabolites to increase skeletal muscle mitochondrial function in immortalised and primary cell lines has been well established (Ryan et al. 2016, 2018, Schnell et al. 2019, Romeu Montenegro et al. 2019). Despite this, there is little evidence for the effects of vitamin D status upon skeletal muscle mitochondrial function in vivo. To date, just one study has examined skeletal muscle mitochondrial function, as measured non-invasively via 31-P MRS, in a cohort of severely deficient patients (Sinha et al. 2013). The authors reported a decrease in PCr recovery time following supplementation with vitamin D, indicative of increased oxidative phosphorylation (Sinha et al. 2013). However the study followed an open label design, making the extrapolation of this data unclear. In order to directly assess the effects of vitamin D status upon skeletal muscle mitochondrial function, we utilised a mouse model of diet-induced vitamin D deficiency. Extending previous observations (Girgis et al. 2015) via the analysis of further vitamin D metabolites, we show that this model allows for the manipulation of vitamin D status without altering mineral homeostasis (Fig. 2A and B). Following 3 months of diet-induced vitamin D deficiency, we report that respiration supported via CI + IIP and the maximal capacity of the ETC is lower when compared to vitamin D-replete mice at the same time point. Following internal normalisation, the above alterations in mitochondrial function were no longer apparent, suggesting the changes observed were due to alterations in the mitochondrial quantity as opposed to quality. Therefore, in order to ascertain whether these increases were mediated by an increase in mitochondrial protein abundance, we assessed the protein content of ETC subunits I–V and citrate synthase activity. Despite functional impairments, we observed no differences in the six markers of mitochondrial protein content within skeletal muscle following the 3 months of diet-induced vitamin D deficiency. Therefore, this suggests that vitamin D deficiency alters mitochondrial function independently of mitochondrial ETC subunit I–V content or citrate synthase activity. Similar observations have been made in vitro following either the treatment of skeletal muscle cell lines with vitamin D metabolites or the knock down of the VDR (Ryan et al. 2016, Ashcroft et al. 2020). In addition, within the skeletal muscle of the recently developed skeletal muscle muscle-specific VDR-KD mouse, mitochondrial protein content remained unchanged (Girgis et al. 2019). We have, however, recently observed a downregulation of mitochondrial genesets in response to in vivo electrotransfer-mediated VDR knock down in rat skeletal muscle (Bass et al. 2020). The further examination of mitochondrial function in such mouse models would help to determine whether the effects of vitamin D deficiency upon skeletal muscle mitochondrial function are a direct result of vitamin D-related signalling.

In addition, we also sought to determine the effects of diet-induced vitamin D deficiency upon body composition within C57BL/6J mice. Previously, no differences in body weight and lean mass were observed following 12 months of diet-induced deficiency in male C57BL/6J mice (Seldeen et al. 2018). In female mice, however, 12 months of diet-induced vitamin D deficiency resulted in reductions in body weight, lean mass and fat mass (Belenchia et al. 2017). Similar to previous reports in male mice, we observed no differences between the vitamin D-replete and -deplete groups in body weight, lean mass, fat mass or food intake at the 3-month time point. We did, however, observe a reduction in lean mass as a percentage of body weight from 1 to 3 months in vitamin D-deplete mice whereas replete mice remained stable over the same time period. This may in part be driven by the fact that we also observed an increase in lean mass as a percentage of body weight following 1 month of diet-induced deficiency. Despite minimal differences in body composition, previous data suggest that vitamin D deficiency impairs physical function in mice (Girgis et al. 2015, Seldeen et al. 2018). Therefore, functional measures of muscle performance may be more relevant to assess the effects of vitamin D deficiency within skeletal muscle. Reductions in physical performance following vitamin D deficiency may also result in reduced daily activity levels which in turn could influence body composition and energy metabolism. It should also be noted that those with serum concentrations of 25(OH)D < 25 nmol/L are at a greater risk of developing sarcopaenia (Visser et al. 2003). Given that the effects of vitamin D deficiency and supplementation seem to be most potent in older individuals, the induction of vitamin D deficiency in aging mouse models may reveal more potent effects on skeletal muscle mass.

In conclusion, we report that mitochondrial function (CI + IIP and ETC) is reduced in C57BL/6J mice following 3 months of diet-induced vitamin D deficiency. These effects are not mediated by alterations in ETC protein content or citrate synthase activity, suggesting that vitamin D deficiency directly affects mitochondrial respiration independent of ETC protein content. Similar to others, we observed minimal differences in body composition following 3 months of diet-induced vitamin D deficiency in male C57BL/6J mice (Seldeen et al. 2017, 2018). Finally, our data provides evidence that vitamin D status is an important determinant of skeletal muscle mitochondrial function in vivo thereby supporting previous in vitro observations.

Declaration of interest

The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.

Funding

The MRC-ARUK Centre for Musculoskeletal Ageing Research was funded through grants from the Medical Research Council [grant number MR/K00414X/1] and Arthritis Research UK [grant number 19891] awarded to the Universities of Birmingham and Nottingham. S P A was funded by a MRC-ARUK Doctoral Training Partnership studentship, joint funded by the College of Life and Environmental Sciences, University of Birmingham.

Author contribution statement

S P A and A P conceived and designed the research project. S P A, G F, A M P, C J, D J H, S D, P M H, P J A and A P performed experiments and data analysis, interpreted results and prepared figures. S P A, P J A and A P drafted the manuscript. All authors approved the final version of the manuscript.

Acknowledgement

The authors like to thank Prof Martin Hewison (University of Birmingham, UK) for valuable discussion relating to vitamin D analysis.

References

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    Body composition and food intake in vitamin D-replete and -deplete C57BL/6J mice. (A) Increase in body weight across dietary period with no differences between differing vitamin D diets (n = 6–10/group). (B) No difference in food intake was observed following dietary intervention (n = 5–6 cages/group). (C and D) Absolute lean mass remained unchanged whilst lean mass as a percentage of body weight was significantly lower in the 3-month vitamin D-deplete mice when compared to the 1-month deplete mice (n = 6–10/group). (E and F) Absolute and percentage of body weight fat mass increased across the dietary period irrespective of dietary intervention (n = 6–10/group). Data mean ± s.d. bMain effect for time (P < 0.05), cGroup × time interaction (P < 0.05), eSignificantly different from 1-month time point of the same group (P < 0.05), *(P < 0.05).

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    Skeletal muscle mass in response to the induction of diet-induced vitamin D deficiency. (A) Gastrocnemius wet weight (milligrams) remains unchanged in response to vitamin D deficiency (n = 6–10/group). (B) Gastrocnemius wet weight normalised to body weight (milligrams/gram) remains unchanged in response to vitamin D defieceny (n = 6–10/group). (C) Higher quadricep wet weight (milligrams) across time but no influence of vitamin D deficiency (n = 6–10/group). (D) Higher body quadricep mass (milligrams/gram) when normalised to body weight across dietary time period with no effect of vitamin D deficiency (n = 6–10/group). (E) Lower tricep mass (milligrams) in reponse to diet-induced vitamin D deficiency (n = 6–10/group). (F) Higher tricep mass when normalised to body weight (milligrams/gram) across dietary time period (n = 6–10/group). Data mean ± s.d. aMain effect of dietary intervention (P < 0.05), bMain effect for time (P < 0.05).

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    Analysis of serum vitamin D metabolites and calcium in response to the induction of diet-induced vitamin D deficiency. (A) Successful induction of vitamin D deficiency was observed via significant reductions in serum 25-hydroxyvitamin D3 (25(OH)D3), 3-epi-25-hydroxyvitamin D3 (3-epi-25(OH)D3), 24,25-dihydroxyvitamin D3 (24,25(OH)2D3), 20,24-dihydroxyvitamin2D3 (20,24(OH)2D3) and 1α,25-dihydroxyvitamin D3 (1α,25(OH)2D3) following 3 months of dietary intervention (n = 6/group). (B) No change in serum calcium following the 1, 2 and 3 months of diet-induced vitamin D deficiency in C57BL/6J mice (n = 5–10/group). Data mean ± s.d. *P < 0.05.

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    Assessment of skeletal muscle mitochondrial oxygen consumption (JO2) in response to diet-induced vitamin D deficiency. (A) Complex I-related leak (CIL) remains unchanged irrespective of the dietary intervention (n = 4–8/group). (B) Complex I phosphorylating (CIP) respiration is higher in the 2- and 3-month vitamin D-replete mice when compared with the 1-month replete mice (n = 6–8/group). (C and D) Phosphorylating respiration supported via complex I and II (CI + IIP) and the maximal capacity of the electron transport chain (ETC) is increased in the vitamin D-replete mice and is higher at the 3-month time point when compared to vitamin D-deplete mice (n = 6–8/group). (E and F) Alteration in absolute rates of respiration are diminished when internally normalised (n = 6–8/group). Data mean ± s.d.. bMain effect for time (P < 0.05), cGroup × time interaction (P < 0.05), eSignificantly different from 1-month time point of the same group (P < 0.05), *(P < 0.05).

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    Assessment of skeletal muscle mitochondrial ADP sensitivity in response to diet-induced vitamin D deficiency. (A, B and C) No change in respiratory capacity in response to the titration of ADP following 1, 2 and 3 month of vitamin D deficiency (n = 6–8/group). (D) No change in the apparent Km for ADP in response to 1, 2 and 3 month of vitamin D deficiency (n = 5–8/group). Data mean ± s.d.

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    Assessment of skeletal muscle ETC protein content following diet-induced vitamin D deficiency. (A, B and D) No changes were observed in expression levels of mitochondrial complex I (NDUFB8), complex II (s.d.HB), complex IV (UQCRC2), following diet-induced vitamin D deficiency (n = 6/group). (C and E) Lower expression levels of mitochondrial complex III (MTCO1) and complex V (ATP5A) across the dietary time period with no differences between vitamin D-replete and -deplete mice (n = 6/group). Data mean ± s.d. bMain effect for time (P < 0.05), cGroup × time interaction (P < 0.05).

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    Assessment of skeletal muscle mitochondrial protein content and enzyme activity following diet-induced vitamin D deficiency. (A, B, C, D, F, G and H) No changes in citrate synthase activity, AMPKThr172 phosphorylation, mitofilin and OPA1 following diet-induced vitamin D deficiency (n = 6/group). Data mean ± s.d.

  • Ashcroft SP, Bass JJ, Kazi AA, Atherton PJ & Philp A 2020 The vitamin D receptor (VDR) regulates mitochondrial function in C2C12 myoblasts. American Journal of Physiology: Cell Physiology 318 C536C541. (https://doi.org/10.1152/ajpcell.00568.2019)

    • Search Google Scholar
    • Export Citation
  • Bass JJ, Kazi AA, Deane CS, Nakhuda A, Ashcroft SP, Brook MS, Wilkinson DJ, Phillips BE, Philp A & Tarum J et al. 2021 The mechanisms of skeletal muscle atrophy in response to transient knockdown of the vitamin D receptor in vivo. Journal of Physiology 599 963979. (https://doi.org/10.1113/JP280652)

    • Search Google Scholar
    • Export Citation
  • Beaudart C, Buckinx F, Rabenda V, Gillain S, Cavalier E, Slomian J, Petermans J, Reginster JY & Bruyere O 2014 The effects of vitamin D on skeletal muscle strength, muscle mass, and muscle power: a systematic review and meta-analysis of randomized controlled trials. Journal of Clinical Endocrinology and Metabolism 99 43364345. (https://doi.org/10.1210/jc.2014-1742)

    • Search Google Scholar
    • Export Citation
  • Belenchia AM, Johnson SA, Kieschnick AC, Rosenfeld CS & Peterson CA 2017 Time course of vitamin D depletion and repletion in reproductive-age female C57BL/6 mice. Comparative Medicine 67 483490.

    • Search Google Scholar
    • Export Citation
  • Bhan A, Rao AD & Rao DS 2010 Osteomalacia as a result of vitamin D deficiency. Endocrinology and Metabolism Clinics of North America 39 32133 1. (https://doi.org/10.1016/j.ecl.2010.02.001)

    • Search Google Scholar
    • Export Citation
  • Bischoff-Ferrari HA, Dietrich T, Orav EJ, Hu FB, Zhang Y, Karlson EW & Dawson-Hughes B 2004 Higher 25-hydroxyvitamin D concentrations are associated with better lower-extremity function in both active and inactive persons aged > or = 60 y. American Journal of Clinical Nutrition 80 752758. (https://doi.org/10.1093/ajcn/80.3.752)

    • Search Google Scholar
    • Export Citation
  • Bouillon R & Verstuyf A 2013 Vitamin D, mitochondria, and muscle. Journal of Clinical Endocrinology and Metabolism 98 961963. (https://doi.org/10.1210/jc.2013-1352)

    • Search Google Scholar
    • Export Citation
  • Canto C & Garcia-Roves PM 2015 High-resolution respirometry for mitochondrial characterization of ex vivo mouse tissues. Current Protocols in Mouse Biology 5 135153. (https://doi.org/10.1002/9780470942390.mo140061)

    • Search Google Scholar
    • Export Citation
  • Cardinale DA, Gejl KD, Ørtenblad N, Ekblom B, Blomstrand E & Larsen FJ 2018 Reliability of maximal mitochondrial oxidative phosphorylation in permeabilized fibers from the vastus lateralis employing high-resolution respirometry. Physiological Reports 6 e13611. (https://doi.org/10.14814/phy2.13611)

    • Search Google Scholar
    • Export Citation
  • Cashman KD, Dowling KG, Skrabakova Z, Gonzalez-Gross M, Valtuena J, De Henauw S, Moreno L, Damsgaard CT, Michaelsen KF & Molgaard C et al. 2016 Vitamin D deficiency in Europe: pandemic? American Journal of Clinical Nutrition 103 10331044. (https://doi.org/10.3945/ajcn.115.120873)

    • Search Google Scholar
    • Export Citation
  • Duncan A, Talwar D, Mcmillan DC, Stefanowicz F & O'reilly DS 2012 Quantitative data on the magnitude of the systemic inflammatory response and its effect on micronutrient status based on plasma measurements. American Journal of Clinical Nutrition 95 6471. (https://doi.org/10.3945/ajcn.111.023812)

    • Search Google Scholar
    • Export Citation
  • Forrest KY & Stuhldreher WL 2011 Prevalence and correlates of vitamin D deficiency in US adults. Nutrition Research 31 4854. (https://doi.org/10.1016/j.nutres.2010.12.001)

    • Search Google Scholar
    • Export Citation
  • Girgis CM, Clifton-Bligh RJ, Hamrick MW, Holick MF & Gunton JE 2013 The roles of vitamin D in skeletal muscle: form, function, and metabolism. Endocrine Reviews 34 3383. (https://doi.org/10.1210/er.2012-1012)

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