Magnesium deficiency prevents high-fat-diet-induced obesity in mice
Hypomagnesaemia (blood Mg2+ <0.7 mmol/l) is a common phenomenon in individuals with type 2 diabetes. However, it remains unknown how a low blood Mg2+ concentration affects lipid and energy metabolism. Therefore, the importance of Mg2+ in obesity and type 2 diabetes has been largely neglected to date. This study aims to determine the effects of hypomagnesaemia on energy homeostasis and lipid metabolism.
Mice (n = 12/group) were fed either a low-fat diet (LFD) or a high-fat diet (HFD) (10% or 60% of total energy) in combination with a normal- or low-Mg2+ content (0.21% or 0.03% wt/wt) for 17 weeks. Metabolic cages were used to investigate food intake, energy expenditure and respiration. Blood and tissues were taken to study metabolic parameters and mRNA expression profiles, respectively.
We show that low dietary Mg2+ intake ameliorates HFD-induced obesity in mice (47.00 ± 1.53 g vs 38.62 ± 1.51 g in mice given a normal Mg2+-HFD and low Mg2+-HFD, respectively, p < 0.05). Consequently, fasting serum glucose levels decreased and insulin sensitivity improved in low Mg2+-HFD-fed mice. Moreover, HFD-induced liver steatosis was absent in the low Mg2+ group. In hypomagnesaemic HFD-fed mice, mRNA expression of key lipolysis genes was increased in epididymal white adipose tissue (eWAT), corresponding to reduced lipid storage and high blood lipid levels. Low Mg2+-HFD-fed mice had increased brown adipose tissue (BAT) Ucp1 mRNA expression and a higher body temperature. No difference was observed in energy expenditure between the two HFD groups.
Mg2+-deficiency abrogates HFD-induced obesity in mice through enhanced eWAT lipolysis and BAT activity.
Keywordsβ-Adrenergic receptor Brown adipose tissue Energy homeostasis Hypomagnesaemia Lipid metabolism Lipolysis Magnesium Obesity White adipose tissue
Brown adipose tissue
Epididymal white adipose tissue
Inguinal white adipose tissue
Low Mg2+ (diet)
Normal Mg2+ (diet)
Respiratory exchange ratio
White adipose tissue
Hypomagnesaemia (blood Mg2+ concentration <0.7 mmol/l) affects approximately 30% of individuals with type 2 diabetes [1, 2]. Hypomagnesaemia is an important risk factor for the development and progression of type 2 diabetes [3, 4, 5]. Low dietary Mg2+ intake and reduced serum Mg2+ concentrations have also been associated with obesity, although with conflicting results [1, 6, 7, 8]. Moreover, reduced blood Mg2+ levels have been correlated with elevated glucose and triacylglycerol concentrations in individuals with type 2 diabetes, suggesting that hypomagnesaemia is associated with insulin resistance and dyslipidaemia .
Mg2+ fulfils many roles including cell growth, membrane stability, enzyme activity and energy metabolism . It is a cofactor for numerous enzymes, primarily because it stabilises ATP and facilitates phosphate transfer reactions [10, 11]. Mg2+ is essential for glycolysis and the citric acid cycle [12, 13]. Because Mg2+ is critical for insulin receptor tyrosine kinase activity, hypomagnesaemia has also been implicated in insulin resistance [14, 15, 16]. Recently, hypomagnesaemia in mice was shown to contribute to enhanced catabolism, but no in-depth metabolic phenotype analysis was performed .
In type 2 diabetes, restoring serum Mg2+ values by oral Mg2+ supplementation improves insulin sensitivity, decreases fasting glucose levels  and corrects the lipid profile [19, 20, 21]. Although Mg2+ is essential for key enzymes in lipid metabolism, including hepatic lipase and lecithin-cholesterol acyltransferase [22, 23], the effects of chronic Mg2+ deficiency on adipocyte function and lipid metabolism remain largely unknown.
In this study, we explored the role of Mg2+ in energy homeostasis, insulin sensitivity and lipid metabolism, by feeding mice a low-fat diet (LFD) or a high-fat diet (HFD) combined with low or normal Mg2+ for 17 weeks. The resulting metabolic effects were extensively characterised. Data were confirmed by an independent replication experiment.
Seventeen-week mouse study: Radboud university medical center
This study was approved by the animal ethics board of the Radboud University Nijmegen (RU DEC 2015-0073) and the Dutch Central Commission for Animal Experiments (CCD, AVD103002015239). Forty-eight male C57BL6/J mice (Charles River Laboratories, Sulzfeld, Germany), aged 9–10 weeks, were randomly allocated to four experimental groups of n = 12 mice. Experimental diets consisted of 10% or 60% energy from palm oil plus 0.03% or 0.21% wt/wt magnesium oxide. Researchers and animal caretakers were blinded for Mg2+ content. On days −1, 84 and 112, mice were housed individually in metabolic cages for 24 h. Blood was collected via cheek puncture at days −1, 28, 56 and 84. At weeks 14 and 15, ITT and GTT, respectively, were performed. After 17 weeks on the diets, mice were anaesthetised by 4% vol./vol. isoflurane and exsanguinated via orbital sinus bleeding. See electronic supplementary material (ESM) for full methods.
Intraperitoneal insulin and glucose tolerance tests
After 14 weeks on the diets, mice underwent an intraperitoneal ITT. After 6 h of fasting, from 08:00 to 14:00, mice were injected with 0.75 U/kg body weight of human insulin (Novorapid, Novo Nordisk, Bagsværd, Denmark). Blood glucose levels were measured at 0, 20, 40, 60, 90 and 120 min. After 15 weeks on the diets, mice underwent an IPGTT. After an overnight fast, from 18:00 to 09:00, mice were injected with 2 g/kg body weight of d-glucose (Invitrogen, Bleiswijk, the Netherlands). Blood glucose was measured at 0, 15, 30, 60 and 120 min. See ESM for full methods.
Quantitative real-time PCR
Total RNA was extracted using TRIzol reagent (Invitrogen, Paisley, UK), subjected to DNase (Promega, Fitchburg, WI, USA) treatment and measured using the Nanodrop 2000c spectrophotometer (Thermo Scientific, Waltham, MA, USA). RNA was reverse transcribed using Moloney murine leukaemia virus (M-MLV) reverse transcriptase (Invitrogen, Bleiswijk, the Netherlands). Gene expression levels were quantified by SYBR-Green (Bio-Rad, Veenendaal, the Netherlands) on a CFX96 real-time PCR detection system (Bio-Rad) and normalised for Gapdh. Primer sequences for Acadl, Adrb3, Atgl (also known as Pnpla2), Cact, Cd36, Cpt1-l (also known as Cpt1a), Cpt1-m (also known as Cpt1b), Cpt2, Fbp1, G6pase (also known as G6pc), Gapdh, Glut1 (also known as Slc2a1), Glut2 (also known as Slc2a2), Glut4 (also known as Slc2a4), Gs, Gyk, Hmgcs1, Hsl (also known as Lipe), Mgll, Pepck1, Pklr, Ppar-α (also known as Ppara), Ppar-γ (also known as Pparg), Srebp1 (also known as Srebf1) and Ucp1 are provided in ESM Table 1.
Epididymal fat and liver tissues were fixed in 10% vol./vol. neutral-buffered formalin (KLINIPATH, Duiven, the Netherlands) in PBS. Samples were dehydrated through alcohol, embedded in paraffin and cut into 4 μm sections. Sections were stained with H&E using standard procedures. The average cell size of 100–300 cells per mouse was determined manually using ImageJ software (v1.48, NIH, Bethesda, MD, USA, RRID:SCR_003070). Liver samples were snap frozen in liquid nitrogen, cut into 10 μm sections, stained with Oil Red O (Sigma-Aldrich, St. Louis, MO, USA) and counterstained with haematoxylin.
Five randomly selected samples of each group were analysed, with no technical replicates, by RNA sequencing. Quality control and RNA sequencing were performed by the Beijing Genomics Institute (BGI), Hong Kong, China. Per sample, 13 million reads were sequenced using the Hiseq 4000 platform (Illumina, San Diego, CA, USA) using a 50 bp single-end module. Clean reads were mapped to Mus musculus transcriptome (GRCm38/mm10) using the HISAT/Bowtie2 tool (RRID:SCR_005476) [24, 25]. RSEM software v1.2.31 (RRID:SCR_013027) was used to quantify gene expression levels (fragments per kilobase million [FPKM] values) . FPKM values were log2 transformed and further analysed in R (www.r-project.org, v3.4.1., RRID:SCR_001905). Heatmaps for individual GO terms were created using the ggplot2 library (r-project) . See ESM for full method details.
Serum Mg2+ was determined using a spectrophotometric assay at 600 nm (Roche/Hitachi, Tokyo, Japan) according to the manufacturer’s protocol. Liver samples were weighed and lysed in lysis buffer (10% wt/vol.) containing 50 mmol/l Tris-HCl pH 7.5, 1 mmol/l EGTA, 1 mmol/l EDTA, 1% vol./vol. Triton X-100, 10 mmol/l glycerophosphate, 1 mmol/l sodium orthovanadate, 50 mmol/l sodium fluoride, 10 mmol/l sodium pyrophosphate and 150 mmol/l sodium chloride. Triacylglycerol concentrations in serum and liver lysate were assayed using an enzymatic kit (Roche Molecular Biochemicals, Indianapolis, IN, USA), according to the manufacturer’s protocol. Serum NEFA (NEFA-C kit, WAKO Diagnostics, Delfzijl, the Netherlands), cholesterol (Human Diagnostics, Wiesbaden, Germany), glucose (Instruchemie, Delfzijl, the Netherlands), leptin (R&D Systems, Minneapolis, MN, USA) and adiponectin (R&D Systems, Minneapolis, MN, USA) concentrations were determined according to manufacturers’ protocols.
3-Methoxytyramine and normetanephrine were analysed by a 6490 LC-MS/MS (Agilent Technologies, Amstelveen, the Netherlands) after solid phase extraction (SPE) Oasis WCX μElution sample cleanup (Waters, Etten-Leur, the Netherlands). A calibration curve was used with 3-methoxytyramine-HCl and normetanephrine-HCl (Sigma-Aldrich, St. Louis, MO, USA) as calibrators. 3-Methoxytyramine-d4-HCl and normetanephrine-d3-HCl (Medical Isotopes, Pelham, NH, USA) were used as internal standards. An ethylene bridged hybrid (BEH) Amide 1.7 μm 100A, 2.1 × 100 mm column (Waters) was used as an analytical column.
Nine-week replication mouse study: MRC Harwell Institute
All experimental procedures were conducted in compliance with the UK Animals Scientific Procedures Act (1986) and University of Oxford ethical guidelines. Thirty-nine male C57BL6/J mice (Medical Research Council [MRC], Harwell, UK) were randomly allocated to four groups of n = 10 mice (n = 9 in the low Mg2+[LowMg]-LFD group). At 8 weeks old, mice were put on experimental diets identical to the Radboud university medical center experiment for 9 weeks. At day 14, mice were housed individually in metabolic cages (Tecniplast, Buguggiate, Italy). Blood was collected via tail bleed at days −1 and 14. Respiration metabolic cages (TSE PhenoMaster Cages, Bad Homburg, Germany) were used at days 28 and 56 and body temperatures were measured by rectal probe (ATP-instrumentation, Ashby, UK). Data were averaged per hour and plotted from 18:30 to 09:30 h. See ESM Methods for full details.
Lipolysis in 3 T3-L1 adipocytes
Differentiated 3 T3-L1 cells (mycoplasma-free, ATCC, Manassas, VA, USA) were incubated for 20 h in DMEM, containing 0 or 1 mmol/l MgCl2. Aliquots of 50 μl medium were taken hourly and heated for 8 min at 65°C. The concentration of NEFA was assessed using the WAKO NEFA-C kit (Instruchemie, Delfzijl, the Netherlands). See ESM Methods for full details.
For the animal experiments, a two-way ANOVA was used to look for a significant interaction effect between the two main variables (dietary fat and Mg2+ content). If there was none, significant differences between the groups were assessed using a two-way ANOVA approach with a Tukey’s multiple comparisons test. If the two-way ANOVA demonstrated a significant interaction effect between the two main variables, an unpaired multiple t test approach using the Holm–Sidak method for multiple comparisons was used. Statistical significance was determined using Graphpad Prism v7 (La Jolla, CA, USA, RRID: SCR_002798). For the lipolysis assays, an unpaired Student’s t test was used.
Differences with a p value of <0.05 were considered statistically significant. Results are presented as mean ± SEM.
Low dietary Mg2+ intake reduces diet-induced obesity in mice
Reduced diet-induced obesity in Mg2+-deficient mice is accompanied by improved insulin sensitivity
Both HFD-fed groups demonstrated increased insulin resistance in the ITT compared with their respective controls (Fig. 2f,g). Low dietary Mg2+ content resulted in a significantly lower AUC of the ITT (Fig. 2g, two-way ANOVA for dietary Mg2+ effect p < 0.05). In the LowMg-HFD group compared with the NormMg-HFD-fed mice, the AUC of the ITT was not significantly different (0.91 ± 0.05 vs 0.72 ± 0.05 mol/l × min in NormMg-HFD-fed and LowMg-HFD-fed mice, respectively, Tukey’s test p = 0.07, Fig. 2f,g).
Insulin resistance is often accompanied by hyperlipidaemia, in particular, high triacylglycerol and NEFA levels. As expected, the HFD-fed mice had higher serum triacylglycerol and NEFA levels than LFD-fed mice (Fig. 2h,i). Interestingly, despite their lower body weight and increased insulin sensitivity, LowMg-HFD-fed mice also had high serum triacylglycerol and NEFA (Fig. 2h,i). In contrast, serum leptin levels correlated with body weight; hence, reduced leptin levels were observed in the LowMg-HFD-fed mice compared with NormMg-HFD-fed mice (Fig. 2j). No difference between the two HFD groups was observed in serum adiponectin and cholesterol (ESM Fig. 1a,b).
Mg2+ deficiency prevents diet-induced hepatic lipid storage
Reduced adipose tissue mass in Mg2+-deficient HFD-fed mice is associated with increased mRNA expression of lipolysis genes
To determine the consequences of low Mg2+ on lipid metabolism, we performed RNA sequencing on eWAT. A principal component analysis using the log2 transformed expression values shows that the samples from both LFD groups cluster closely together, indicating the absence of a strong Mg2+ effect, whereas there is a clear separation of NormMg-HFD vs LowMg-HFD gene expression profiles (Fig. 4e). To investigate the effect of Mg2+ on specific biological pathways, the fold changes for groups of genes belonging to the same gene ontology (GO) were analysed. GO term analysis indicated that processes associated with adiposity (e.g. inflammation) were downregulated in LowMg-HFD-fed vs NormMg-HFD-fed mice, in accordance with decreased adipose tissue mass (ESM Table 2). Interestingly, despite the increased insulin sensitivity of the LowMg-HFD-fed mice, several key genes involved in the triacylglycerol catabolism pathway (lipolysis) were upregulated in the LowMg-HFD vs the NormMg-HFD group, which may explain the reduced lipid storage as well as the high serum NEFA levels (Fig. 4f). A modest increase in acyl-CoA dehydrogenase dependent β oxidation was observed in the LowMg-HFD-fed mice vs the NormMg-HFD-fed mice (Fig. 4g). The metabolic effects of Mg2+ in eWAT appear to be specific to lipid homeostasis, as there was no clear effect on glycolysis (ESM Fig. 3g). Although serum cholesterol levels were not different between the experimental groups, cholesterol biosynthesis was greatly reduced in the LowMg-HFD-fed vs the NormMg-HFD-fed mice (ESM Fig. 3h, ESM Table 2).
To investigate whether hypomagnesaemia has a direct effect on lipolysis in eWAT, we examined the effect of Mg2+ on lipolysis in differentiated 3 T3-L1 cells in vitro. Unstimulated lipolysis was not different at 0 or 1 mmol/l Mg2+, indicating that Mg2+ deficiency does not directly induce lipolysis in adipocytes (ESM Fig. 4a).
mRNA expression of the β3-adrenergic receptor is increased in LowMg-HFD mice
Expression of Ucp1 in BAT, which is essential for non-shivering thermogenesis, was upregulated in NormMg-HFD-fed mice compared with NormMg-LFD-fed mice (Fig. 5g). In line with increased β3-adrenergic signalling, Ucp1 expression was further increased in BAT of LowMg-HFD-fed mice. BAT thermogenesis is strongly regulated by fatty acid availability . Indeed, genes involved in NEFA metabolism of BAT are upregulated (Fig. 5h–j) (Atgl, Cpt1-m and Acadl). In contrast, mRNA levels of glucose transporters 1 and 4 (Glut1/4) in BAT were unchanged in LowMg-HFD-fed compared with NormMg-HFD-fed mice (ESM Fig. 5b,c). mRNA expression of the fatty acid transporter Cd36 and of the important metabolic transcription factors peroxisome proliferator-activated receptor alpha (Pparα) and gamma (Pparγ) remained unchanged in BAT between the NormMg-HFD-fed and LowMg-HFD-fed mice (ESM Fig. 5d–f).
LowMg-HFD-fed mice have increased body temperature but equal energy expenditure
Hypomagnesaemia has been repeatedly reported in type 2 diabetes and the metabolic syndrome [1, 2, 14], but the role of Mg2+ in lipid metabolism has been largely overlooked. Here, we demonstrate that low dietary Mg2+ intake ameliorates HFD-induced obesity. The lower body weight results in beneficial metabolic effects including improved insulin sensitivity, reduced hepatic steatosis and lower WAT inflammation. Nevertheless, serum triacylglycerol and NEFA concentrations were increased in the low Mg2+ HFD group, corresponding to increased eWAT mRNA expression of lipolysis genes. These findings establish Mg2+ as an important regulator of body weight and lipid metabolism.
In this study, a Mg2+-deficient diet ameliorated HFD-induced weight gain in mice. This was the result of reduced adiposity, because lean body mass was similar between the two HFD groups and both eWAT and iWAT weight were lower in mice fed a LowMg-HFD compared with a NormMg-HFD. The reduced body weight was associated with favourable metabolic effects. IPGTT, IPITT and fasting glucose levels indicated enhanced insulin sensitivity. Moreover, the reduced body weight of the LowMg-HFD mice led to a complete absence of hepatic steatosis and RNA sequencing of the eWAT demonstrated downregulation of pro-inflammatory pathways. Despite these beneficial effects, blood lipid levels remained high. In line with our data, others have demonstrated that low dietary Mg2+ intake reduced body weight in several rat models of Mg2+ deficiency [34, 35, 36, 37]. However, these studies did not address the underlying cause or investigate the effects on lipid metabolism.
Our animal data is strengthened by the results of Chubanov et al  where severe hypomagnesaemia via Trpm6 knockout also resulted in a catabolic phenotype and improved insulin sensitivity . The catabolic phenotype of Mg2+-deficient mice leads to hyperlipidaemia, which has considerable adverse effects in individuals with type 2 diabetes [38, 39]. Nevertheless, the low Mg2+ HFD does not completely mimic the human situation because the hypomagnesaemia induced in mice is more severe . Moreover, an unhealthy human diet consists of both high fat and sugar, whereas the HFD in mice purely depends on palm oil. Indeed, Mg2+-deficiency in high-fructose diets has adversely affected insulin sensitivity and lipid homeostasis in rats. This shows the considerable differences in the role of Mg2+ in the metabolism of lipids vs carbohydrates [40, 41]. Future studies should investigate the role of Mg2+ in combined fat and sugar diets. These differences may explain why, in humans, higher oral Mg2+ intake and Mg2+ supplementation have beneficial effects on metabolic variables, which apparently contrasts with our animal data [18, 19, 20].
In our study, the reduced WAT mass of LowMg-HFD-fed mice was associated with enhanced lipolysis gene expression, causing high serum NEFA and triacylglycerol levels. These findings suggest that LowMg-HFD-fed mice depend more on mitochondrial β-oxidation, rather than glycolysis, for energy production. However, our energy metabolism experiments demonstrated neither differences in energy expenditure nor in RER between the NormMg-HFD and LowMg-HFD groups. It should be noted, however, that both HFD groups mainly depend on lipids for energy metabolism, masking potential RER differences between these groups. Moreover, despite equal energy expenditure, the NormMg-HFD-fed mice are considerably heavier than LowMg-HFD-fed mice and therefore have a higher energy demand. Several studies have discussed the considerable difficulties associated with the interpretation of energy expenditure data and emphasised that body weight differences complicate interpretation [42, 43]. Increased thermogenesis may explain why energy expenditure does not differ between LowMg-HFD-fed and the heavier NormMg-HFD-fed mice. Although the effects are modest, the LowMg-HFD-fed mice had a significantly higher body temperature and increased Ucp1 expression in BAT, indicative of higher thermogenesis. Cold-exposure studies are necessary to further investigate the role of Mg2+ status in BAT activation, WAT browning and thermogenesis.
The increased lipolysis and brown adipose tissue activity were associated with higher β3-adrenergic receptor expression in eWAT and BAT of LowMg-HFD-fed mice. β3-receptor knockout mice have increased lipid stores and impaired WAT browning [44, 45]. Activation of the β3-adrenergic receptors in mice using agonist CL316243 decreases adipose tissue mass, improves insulin sensitivity, increases uncoupling protein-1 (UCP1)-dependent thermogenesis and activates a cycle of concomitant lipolysis and de novo lipogenesis [46, 47]. Interestingly, this is exactly the phenotype that was observed in the LowMg-HFD-fed mice, although to a lesser extent. A link between Mg2+, β-adrenergic signalling and lipolysis is not without precedent. Use of β-adrenergic agonists, which stimulate lipolysis, have been associated with decreased blood Mg2+ levels [1, 48, 49]. Mg2+ has also been shown to reduce catecholamine release from the adrenal medulla  and Mg2+ deficiency is associated with higher urinary levels of adrenaline and noradrenaline (norepinephrine) . Moreover, Mg2+ supplementation has been suggested to regulate lipolysis, as it prevents hyperlipidaemia in diabetic rats and reduces serum triacylglycerol levels in individuals with type 2 diabetes [20, 51]. Further research is required to determine exactly how hypomagnesaemia increases β-adrenergic signalling and how β-adrenergic signalling can induce hypomagnesaemia.
A strength of this study is that the model used to induce type 2 diabetes and low dietary Mg2+ intake closely resembles the human situation. The Western diet contains high amounts of processed foods consisting of high energy and low Mg2+. Moreover, the extensive phenotyping of the animals in this study provides new avenues for research into the pivotal role of Mg2+ in metabolism. The data obtained in this study are robust, as a replicate animal experiment was performed in a separate institution, confirming our results.
Our study has limitations. First, because of the large weight differences induced by the Mg2+ deficient diet, it is difficult to specifically attribute the metabolic changes of the mice to their lower body weight or their Mg2+ deficiency. In addition, our study design did not allow us to study in more depth the contribution of disturbed β-adrenergic signalling to the differences in body weight, eWAT lipolysis, BAT activity and hyperlipidaemia. Although our data and previous studies support a role for Mg2+ in β-adrenergic signalling [37, 50], further studies are required to establish the exact role of Mg2+ in catecholamine secretion and signalling.
In conclusion, our results demonstrate that hypomagnesaemia in mice prevents HFD-induced weight gain by enhanced BAT activity and increased eWAT lipolysis gene expression. Consequently, this led to improved insulin sensitivity and absent hepatic steatosis. These results underline the pivotal function of Mg2+ in maintaining a healthy energy metabolism.
The authors thank M. Voet, F. Krewinkel, T. Peters, K. de Haas-Cremers, M. School, H. Janssen-Wagener, S. Mulder, T. van Herwaarden, A. Hijmans (Radboud Institute for Molecular Life Sciences, Radboud university medical center, Nijmegen, the Netherlands) for their excellent technical support with the animal study and measurements, and H. Cater, M. Rohm and M. Brereton (Department of Physiology, Anatomy & Genetics, University of Oxford, Oxford, UK) for their insights and scientific input. Some of the data were presented as an abstract and poster at the Experimental Biology meeting in Chicago in 2017.
SK, JdB, JH, RB, FA and CT conceived and designed the study; SK, JdB, JvD, CO-B and WA contributed to data acquisition; SK, JdB, JvD and WA analysed the data; all authors interpreted the data, drafted the article, revised it and approved the final version. JdB is the guarantor of this work.
This work was supported by funding from the Radboud Institute for Molecular Life Sciences and by grants from the Netherlands Organization for Scientific Research (J. Hoenderop, the Netherlands Organisation for Scientific Research (NWO) VICI 016.130.668), the Wellcome Trust (884655, 089795) and the European Research Council (ERC; 322620). J. van Diepen is supported by a Veni Grant from NWO (NWO VENI 91616083). J. de Baaij is supported by grants from NWO (Rubicon 825.14.021, NWO VENI 016.186.012) and the Dutch Diabetes Research Foundation (2017.81.014). F. Ashcroft holds an ERC Advanced Investigatorship and a Royal Society Research Wolfson Merit Award.
Duality of interest
The authors declare that there is no duality of interest associated with this manuscript.
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