Hydrogen sulfide regulates circadian-clock genes in C2C12 myotubes and the muscle of high-fat-diet-fed mice

Accepted Manuscript Hydrogen sulfide regulates circadian-clock genes in C2C12 myotubes and the muscle of high-fat-diet-fed mice Rajesh Parsanathan, Sushil K. Jain PII:

S0003-9861(19)30441-2

DOI:

https://doi.org/10.1016/j.abb.2019.07.019

Reference:

YABBI 8054

To appear in:

Archives of Biochemistry and Biophysics

Received Date: 7 June 2019 Revised Date:

18 July 2019

Accepted Date: 24 July 2019

Please cite this article as: R. Parsanathan, S.K. Jain, Hydrogen sulfide regulates circadian-clock genes in C2C12 myotubes and the muscle of high-fat-diet-fed mice, Archives of Biochemistry and Biophysics (2019), doi: https://doi.org/10.1016/j.abb.2019.07.019. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Rajesh Parsanathan and Sushil K. Jain* Department of Pediatrics and Center for Cardiovascular Diseases and Sciences, Louisiana State University Health Sciences Center-Shreveport, 1501 Kings Highway, Shreveport, Louisiana 71130, USA

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Keywords: Hydrogen sulfide (H2S), glutathione (GSH), oxidative stress (OS), core clock genes (CCG)

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Address correspondence to: Dr. Sushil K. Jain 1501 Kings Highway Shreveport, Louisiana 71130, United States Phone: +1-318-675-6086 Fax: +1-318-675-6059 [email protected]

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Abbreviations: Bmal1, Brain and Muscle ARNT-Like 1; CBS, Cystathionine-Beta-Synthase; Clock, Clock Circadian Regulator; Cry2, Cryptochrome Circadian Regulator 2; CSE, Cystathionine Gamma-Lyase; GCLC, Glutamate-Cysteine Ligase Catalytic Subunit; GCLM, Glutamate-Cysteine Ligase Modifier Subunit; GSH, glutathione; GSR, GlutathioneDisulfide Reductase; GSS, Glutathione Synthetase; H2S, hydrogen sulfide; HFD, high-fat diet; HG, high glucose; KD, knockdown; MCP-1, monocyte chemoattractant protein 1; MPST, Mercaptopyruvate Sulfurtransferase; OS, oxidative stress; Per2, Period Circadian Regulator 2; PGC-1α, PPARG Coactivator 1 Alpha; PPARα, Peroxisome Proliferator-Activated Receptor Alpha; PPARγ, Peroxisome Proliferator-Activated Receptor Gamma; Rev-erbα, Nuclear Receptor Subfamily 1 Group D Member 1; RORα, RAR Related Orphan Receptor A; ROS, reactive oxygen species; RXRα, Retinoid X Receptor Alpha; siRNA, short interference RNA; TNF, tumor necrosis factor;

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Hydrogen sulfide regulates circadian-clock genes in C2C12 myotubes and the muscle of high-fat-diet-fed mice

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ABSTRACT Hydrogen sulfide (H2S) is an endogenous novel gasotransmitter which is implicated in the pathophysiology of the metabolic syndrome. Core clock genes (CCG) and its controlled genes disruption is implicated in the progression of metabolic syndrome. We examined whether H2S has any effect on CCG in the skeletal muscle of mice fed a highfat diet (HFD) and in myotubes. In the muscle of HFD-mice, the expression of H2S biosynthesis enzyme genes (CSE, CBS, and 3-Mpst) along with antioxidant genes (GCLC, GCLM, GSS, and GSR) involved in GSH biosynthesis and recycling were reduced significantly, but the oxidative stress (OS) increased. Expression of the CCG (Bmal1, Clock, RORα, Cry2, Per2) and clock-controlled genes (PPARγ, PGC-1α, RXRα) was downregulated, whereas the levels of PPARα mRNA were upregulated. Similar to that in the muscle of HFD-mice, in vitro myotubes exposed to high glucose or palmitate to mimic metabolic syndrome, showed an increased OS and decreased in CSE mRNA, H2S production and CCG mRNA levels were also downregulated.TNF and MCP-1 treatment on the myotubes was similar to that observed in HFD-muscle, with that the Rev-erbα mRNA was upregulated. Inhibition (siRNA/pharmacological inhibitors) of both CSE and GCLC (the rate-limiting enzyme in GSH biosynthesis) decreased H2S, and increased OS; Bmal1 and Clock mRNA levels were downregulated, while Rev-erbα increased significantly in these conditions. CSE KD myotubes were post-treated with an H2S donor partially restored the mRNA levels of core clock genes. These findings report that the deficiencies of H2S/GSH impair expression of CCG and treatment with H2S donor or GSH precursor exert a positive effect over CCG. Thus, suggest that H2S as a new endogenous factor for regulating circadian clock, and its donors could provide a novel chronopharmacological therapy to manage metabolic disorders.

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1. INTRODUCTION Clock genes are critical for the generation and regulation of circadian rhythms [1-3]. Circadian rhythms are ~24-hour physiological and behavioral rhythms synchronized to the environmental cycles of light/dark and fasting/feeding [1] and with the metabolic events that occur within peripheral tissues, including the skeletal muscle. Subjects with metabolic syndrome and obesity exhibit decreased skeletal muscle strength and function compared to healthy weight subjects [4], as well as impaired skeletal muscle mitochondrial function and contributed reactive oxygen species (ROS) generation [5]. In particular, obese individuals present higher ratios of type II-to-type I skeletal muscle fibers; type II fibers have been shown to generate 2- to 3-fold more ROS production than type I fibers [5, 6]. Also, TNF is exclusively expressed by type II muscle fibers and catalyzes skeletal muscle-derived oxidative stress (OS) [7, 8]. Animals fed high-fat diets (HFD) showed disrupted circadian-controlled clock genes, and human subjects demonstrated altered sleep cycles, including interrupted sleep. Deterioration of sleep quality results in increased appetite and a tendency to develop metabolic syndrome, obesity, and diabetes [2, 9, 10] along with altered endogenous gasotransmitter reductant hydrogen sulfide (H2S) and glutathione (GSH) [11, 12]. The current literature suggests that the perturbations in the circadian system and clock genes signaling may be causal in the development of metabolic syndrome. H2S affects the intracellular redox state [13, 14]. Diurnal physiological fluctuations of plasma H2S levels have been reported in rats and mice [15, 16]. In cultured hepatocytes, H2S can affect the intracellular redox status (NAD+/NADH) and upregulate the core clock genes (CCG) such as Clock, Per2, Bmal1, and Rev-erbα [14]. As far as we know, no previous study has examined the effect of H2S/GSH antioxidant systems on the core clock genes (Bmal1, Clock, Rev-erbα, RORα, Cry2, and Per2) and clock-controlled genes (PPARα, PPARγ, PGC-1α, and RXRα) in C2C12 myotubes or in the peripheral tissue (skeletal muscle) of HFD-fed mice. This study reports simultaneous downregulation of both H2S/GSH synthesis and CCG and its controlled genes in the skeletal muscle of mice fed an HFD. Cell culture studies showed that the downregulation/upregulation of physiological levels of H2S and GSH exerted inhibitory/beneficial effects by tuning the peripheral CCG and glucose homeostasis genes. Thus, suggest that H2S as a new endogenous factor for regulating circadian clock, and its donors or precursors have the potential to be used as a novel chrono-pharmacological therapeutic approach in the management of metabolic disorders.

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2. MATERIALS AND METHODS 2.1. Materials All chemicals and reagents used in the study, which were of molecular and analytical grade, were purchased from Sigma Chemical Co. (St. Louis, MO) unless otherwise mentioned.

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2.2. Animal treatment Male C57BL/6J mice (5 weeks old, 20-24 g) were purchased from The Jackson Laboratory (Bar Harbor, ME, USA) and housed for one week of acclimation in a temperature-controlled room (22±2 °C) with 12/12 h light/dark cycles (lights on at 06:00 am; Zeitgeber time (ZT) 0) and free access to food and water. After the week of acclimation, the mice were divided into two groups. They were fasted overnight and then weighed. The mice were tested for hyperglycemia by measuring their blood glucose concentrations before starting the treatment plan. Blood glucose was assessed by tail prick using a glucometer (Accu-Chek, Boehringer Mannheim Corp., Indianapolis, IN, USA). To induce diabetes, the mice were continuously fed either a high-fat diet (HFD, Harlan TD.88137, providing 42% calories as fat) or a standard chow diet (Control, Harlan TD.08485, providing 5.2% calories as fat) for 16 weeks, a period based on data from a recent study [17]; the detailed composition of these diets is given in a previous publication [18]. At the end of 16 weeks, the animals were fasted overnight and then euthanized for analysis by exposure to isoflurane (Webster Veterinary Supply Inc., Devens, MA). The animals were then perfused with cold saline to free them of residual blood. Skeletal muscle (gastrocnemius) was collected immediately, quickly diced, and frozen in liquid nitrogen at -80 °C. All of the procedures mentioned above were completed in both of the groups between ZT 2 and 3. This model of dietary-induced insulin resistance created both fasting hyperglycemia and hyperinsulinemia and thus represented a reasonable model of the human condition. All procedures followed the guidelines of the ethical standards of the institution and were performed after receiving approval from the Institutional Animal Committee.

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2.3. Cell culture Mouse C2C12 myoblasts (ATCC® CRL-1772™, Manassas, VA) were cultured at 37 °C in an atmosphere of 5% CO2 in growth medium (GM) consisting of Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum and antibiotics (penicillin, streptomycin). Differentiation of myoblasts into myotubes was

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induced when the cells had achieved 90-95% confluence by switching the medium from GM to differentiation medium (DM) consisting of DMEM supplemented with 2% horse serum (5 days). Reduction of serum allowed cellto-cell fusion and formation of myotubes. The cells were examined each day to evaluate the degree of differentiation, which was determined under a phase-contrast microscope as the percentage of nuclei present in the multinucleated myotubes.

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2.7. Relative gene expression Total RNA was extracted from cells or tissue using TRIzol Reagent (Invitrogen) following the manufacturer’s instructions. The concentration and quality of the extracted RNA were determined on a NanoDrop spectrophotometer (Thermo Scientific). RNA (1 µg) from each sample was reverse transcribed according to the manufacturer’s instructions using a High Capacity RNA-To-cDNA kit (Applied Biosystems) to synthesize cDNA. QPCR was performed using Applied Biosystems™ TaqMan™ Gene Expression Assays with primer/probe sets Nrf2 (Mm00477784_m1), GCLC (Mm00802655_m1), GCLM (Mm00514996_m1), GSS (Mm00515065_m1), GSR (Mm00439154_m1), CSE (Mm00461247_m1), CBS (Mm00460654_m1), 3-MPST (Mm00460389_m1), Bmal1 (Mm00500226_m1), Clock (Mm 00455950_m1), Rev-erbα (Mm01310356_g1), RORα (Mm01173766_m1), Cry2 (Mm01331539_m1), Per2 (Mm00478099_m1), PPARα (Mm00440939_m1), PPARγ (Mm00440940_m1), PGC-1α (Mm01208835_m1), and RXRα (Mm00441185_m1). The relative amount of mRNA was calculated using the relative quantification (∆∆CT) method. The relative amount of each mRNA was normalized to the housekeeping gene GAPDH. In compliance with MIQE guidelines, technical replicates (n=3) and biological replicates (n=4) were

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2.4. RNA interference of CSE and GCLC siRNA were purchased from Santa Cruz Biotechnology, Inc. (Dallas, TX): CTH siRNA (m) (sc-142618), γ-GCSc siRNA (m) (sc-41979), and Control siRNA-A (sc-37007), a scrambled nonspecific RNA duplex that shares no sequence homology with any of the genes, which was used as a negative control. Cells were transiently transfected with 50 or 100 nM siRNA complex using Lipofectamine™2000 transfection reagent (Invitrogen, Carlsbad, CA) following the method described earlier [19]. The next day the cells were harvested or used for different parameters [20].

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2.5. Cell culture treatments Differentiated myotubes were treated with either high glucose (HG; 25 mM) or palmitate (0.6 mM), for 24 h in basal medium (without serum or any growth factors), respectively to mimic the micro-environment that muscles experience in vivo. Mannitol (19.5 mM) was used as an osmolarity control for the HG group since the control group received a glucose concentration of 5.5 mM. Blood glucose levels can sometimes become elevated to levels as high as 30 mM under uncontrolled diabetic conditions [21]. Equimolar BSA was used as a control for the BSA conjugated palmitate group. As a guide, TNF can be detected in the serum of patients with metabolic syndrome at levels ≥100 pg/mL [22], at levels 8-fold higher than those of healthy controls in cardiomyopathy patients [23], or at levels of 250 pg/mL (equivalent to the TNF used here) immediately after injection of Escherichia coli endotoxin [24]. Serum MCP-1 increased by 37.278 pg/mL for every 1% rise in HbA1c, and the levels that increased (>500 pg/mL) corresponded to HbA1c values of more than 12.2% in uncontrolled diabetic subjects [25]. Hence, the MCP-1 concentrations (250 pg/mL) used in this study precisely reflect those found under diabetic conditions. Myotubes treated with pharmacological inhibitors propargylglycine (PPG), aminooxyacetate (AOA) 100µM for 12 h either separate or together to inhibit H2S producing enzymes (CSE and CBS). Differentiated myotubes were treated with either an H2S donor (NaHS; 10 or 20 µM) or a precursor of GSH (Lcysteine; 300 µM) to boost the level of H2S production and GSH cellular content following the methods used in our previous published studies [17, 19, 20, 26]. In another set of experiments, CSE knockdown cells (100 nM siRNA) were post-treated with H2S donor (NaHS 20 µM) for 6 h.

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2.6. Cell viability detection Cell viability was determined using the Alamar Blue reduction bioassay. This method is based on Alamar Blue dye reduction by live cells. Briefly, cells were plated into 96-well plates after treatment per the above-described protocols, AlamarBlue® Cell Viability Reagent (DAL1100, ThermoFisher Scientific, Waltham, MA) was added, and the cells were incubated at 37 °C in the dark for 4 h. Absorbance was read at 590 nm using a plate reader. Data are expressed as a percentage of the viable cells counted compared to the total cells counted.

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included in all of our experiments. Data were analyzed using the comparative CT method, and the fold change was calculated using the 2−∆∆CT method using a 7900HT Real-Time PCR system and software (Applied Biosystems). The results were expressed either as the relative quantification (RQ) or log2 FC (fold change) relative values.

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2.8. Preparation of tissue or whole cell extracts Muscle tissue (∼100 mg) was homogenized in RIPA buffer on ice using a rotor-stator to extract total protein. RIPA buffer (50 mM Tris pH 8, 150 mM NaCl, 1% NP-40, 0.5% deoxycholic acid, and 0.1% SDS) was supplemented with protease and phosphatase inhibitors (1 mM PMSF, 5 µg/mL leupeptin, 2 µg/mL aprotinin, 1 mM EDTA, 10 mM NaF, and 1 mM NaVO4). For whole cell extraction, after treatment, the myotubes were washed twice with icecold PBS and lysed in RIPA buffer. Lysates were then centrifuged for 10 min at 10,000 x g at 4 °C. Supernatants were collected, and the protein concentrations determined using a BCA assay kit (Pierce/Thermo Scientific, Rockford, IL) for various assays.

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2.9. H2S production assay H2S production capacity (Lead Sulfide Method) was measured as previously described [27]. Briefly, frozen skeletal muscle was homogenized in passive lysis buffer (Promega), and the volume normalized to protein content. An equal volume/protein amount was added to a reaction master mix containing PBS, 1 mM pyridoxal 5'-phosphate (PLP) (Sigma), and 10 mM Cys (Sigma) and placed in a well-format (96-well) plate. Lead acetate H2S detection blot paper, saturated with lead acetate and then dried, was placed above the plate and incubated for 1-2 h at 37 °C until H2S in the gas phase reacted with the paper to form dark lead sulfide. In vitro studies were carried out using live myotubes in 96-well plates and a growth medium supplemented with 10 mM Cys and 10 mM PLP. Lead acetate H2S detection blot paper was placed over the plate for 2-24 h at 37 °C in a CO2 incubator. Results are presented as the average H2S production capacity relative to the control group.

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2.10. Glutathione and oxidative stress evaluation Levels of total GSH from muscle and cultured cells were quantified using a fluorimetric method (CS1020, SIGMA; Glutathione Assay Kit, Fluorimetric) and expressed as GSH (nmol/mg protein). Oxidative stress was assessed by measuring the quantification of protein carbonyls, and MDA using a Protein Carbonyl Colorimetric and TBARS Assay Kits (Cayman Chemical, Ann Arbor, Michigan, USA). Both protein carbonyl and MDA are expressed as nmol/mg protein. H2O2 was measured using an Amplex Red Hydrogen peroxide/peroxidase assay kit (A22188, Invitrogen, Eugene, Oregon, USA) and expressed as H2O2 (nmol/mg protein). Intracellular reactive oxygen species (ROS) levels were measured in myotubes using the oxidant-sensitive probe 2′,7′ dichlorofluorescein diacetate [H2DCFDA] (D6883 SIGMA). The change in intracellular ROS levels was plotted as mean fluorescence intensity (MFI), a fold change compared with the control. Protocols, as provided in the manufacturer’s instructions, were followed using appropriate controls and standards.

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2.11. Statistical analyses All statistical analyses were conducted using GraphPad Prism 7 for Windows, version 7.04 (GraphPad Software, La Jolla, CA). Data were generated from multiple repeats of different biological experiments to obtain the mean values and standard errors of the mean. Statistical differences were measured using Student's t-test comparing two averages (means) and one-way ANOVA with Bonferroni or Dunnett corrections for multiple comparisons when appropriate. Significance was set at p < 0.05. 3. RESULTS 3.1. HFD attenuates antioxidant system and alters core clock and its controlled genes in mouse skeletal muscle HFD-fed mice (following 16 weeks of consuming an HFD) showed a metabolic phenotype similar to that of obese type 2 diabetic human subjects. The imbalance between high oxidative stress and decreased antioxidant response is thought to be the primary reason for high-fat-mediated function deterioration. Based on this information, experiments were performed to analyze the alteration of antioxidant capacity and the oxidative stress marker reactions. First, H2S production capacity and the mRNA levels of H2S biosynthesis enzymes (CSE, CBS, and 3Mpst) were decreased significantly in the muscle of HFD-treated mice (Fig. 1A-C). Also, critical skeletal muscle antioxidant genes (GCLC, GCLM, GSS, and GSR) involved in GSH biosynthesis and recycling were significantly downregulated, along with fundamental components of the antioxidant system (Nrf2), in the skeletal muscle of HFD-fed mice compared to those of the control group (Fig. 1D-E). In comparison, levels of oxidative products such as H2O2, MDA, and protein carbonyl content were increased in HFD-treated muscle (Fig. 1F-H). Surprisingly, the

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chronic stress due to consumption of a high-fat diet caused a decrease in the expression of circadian core clock genes (Bmal1, Clock, RORα, Cry2, and Per2) and its controlled genes (PPARγ, PGC-1α, and RXRα) and an increase in their transcription factor (PPARα) (Fig. 1I). Although Rev-erbα showed an increasing trend in HFDtreated muscle, this was not statistically significant (Fig. 1I). Altogether, this information indicates that HFD stress attenuates antioxidant systems (H2S and GSH), escalates oxidative stress, and significantly alters CCG and its controlled genes expression in skeletal muscle.

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3.2. In vitro myotubes, glucolipotoxicity affects antioxidant gene expression and modulates core clock and its controlled genes Our in vivo findings were further supported by results obtained in vitro using the myotubes in high glucose-mediated glucotoxicity and palmitate-mediated lipotoxicity models. Compared to the control group, under glucolipotoxicity in vitro conditions, the mRNA levels of the genes Nrf2, GCLC, and CSE were downregulated, along with those of H2S and glutathione, but the levels of H2O2 and ROS production were elevated (Fig. 2A-F). Similarly, the level of expression of core clock and its controlled genes was downregulated in high glucose- and palmitate-treated myotubes (Fig. 2G); an effect that was also observed in the muscle of mice fed an HFD. However, compared to that in the control group, the expression of Rev-erbα and PPARα was not altered by HG or PA treatments (Fig. 2G). Cell viability was not affected under any of the conditions. These findings support the idea that in vitro glucolipotoxicity evoked oxidative stress attenuates the antioxidant system and modulates CCG and its controlled genes.

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3.3. Inflammatory cytokines (MCP-1 and TNF) decreases antioxidant capacity and upregulates Rev-erbα Oxidative stress and the inflammation response are thought to be the primary reasons for high-fatmediated function deterioration in different tissues. In vitro, myotubes were exposed to inflammatory cytokines to mimic low-grade inflammation, which is observed in both obesity and diabetes. The myotube antioxidant genes, H2S, and GSH, were significantly downregulated, and the oxidative stress markers increased by MCP-1 and TNF (Fig. 3A-F). The levels of circadian core clock genes and their regulated transcription factors were downregulated by inflammatory cytokines (Fig. 3G). Surprisingly, treatment with MCP-1 and TNF elevated the expression of Reverbα compared to that in the control group (Fig. 3G). These results indicate that, while inflammatory cytokines in vitro induce stress like that of glucolipotoxicity, they may also contribute to the upregulation of Rev-erbα.

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3.4. Altered profile of circadian core clock and its controlled clock genes expression in CSE and GCLC knockdown myotubes Myotube knockdown (KD) with CSE and GCLC siRNA caused decreased expression of antioxidant genes and their status along with increased levels of oxidative stress markers (Fig. 4A-H). CSE and GCLC KD cells showed attenuated levels of Bmal1, Clock, PPARγ, PGC-1α, and RXRα mRNA, but levels of Rev-erbα increased significantly compared to those of the control group (Fig. 4I). The mRNA levels of RORα, Cry2, and PPARα decreased in CSE KD but not in GCLC KD myotubes (Fig. 4I). Per2 mRNA was upregulated only in the GCLC KD cells; no alteration was observed in CSE KD cells (Fig. 4I). Pharmacological inhibition of H2S producing enzymes alters CCG similar to that of CSE KD cells (Fig. 4J). Collectively, these data illustrate that a deficiency in H2S or GSH alters the expression of the CCG and its controlled genes differently. 3.5. H2S and GSH induces regulation of core clock genes and its regulated transcription factors genes in vitro The possible effect of H2S or GSH on the circadian core clock and its controlled genes was explored with the antioxidant precursors NaHS (an H2S donor) or L-cysteine (a GSH/H2S precursor) following the methods used in previous publications [17, 19]. Results showed that the mRNA levels of antioxidant genes, including Nrf2, GCLC, and CSE, were increased by NaHS/LC treatment, and that these precursors increased cellular H2S production, GSH levels, and decreased the levels of oxidative stress markers (Fig. 5A-F). Compared to levels in the control group, the mRNA levels of Bmal1, Clock, Cry2, Per2, PPARα, PPARγ, PCG-1α, and RXRα were significantly increased in response to treatment with NaHS and LC, indicating that H2S and GSH may have a direct or indirect effect on these genes (Fig. 5G). Interestingly, treatment with NaHS and LC resulted in significantly decreased Rev-erbα and RORα mRNA levels compared to those in control myotubes (Fig. 5G). CSE KD cells post-treated with an H2S donor partially restored the mRNA levels of core clock genes (Fig. 5H). These data suggest that H2S and GSH regulate the CCG and its regulated transcription factors in vitro.

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DISCUSSION The circadian clock coordinates anabolic and catabolic processes in all primary organs/tissues in response to sleepawake and fast-fed cycles in mammals; roughly 40% of protein-coding genes display rhythmic transcription, at least

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in mice [28]. The circadian clock senses the metabolic cues and circadian misalignment induced by metabolic diseases [29]. The obesogenic diet has been shown to induce circadian disruption; conversely, restricting access to high-fat food regulates circadian cycles, and ameliorates diet-induced metabolic syndrome [30-32]. Consumption of an HFD also increases oxidative stress and decreases levels of hydrogen sulfide and glutathione (a major antioxidant) in the liver and skeletal muscle in mice [33-35]. Circadian clock mechanism involves heteromerization of transcription factors, Clock and Bmal1, and binding of these factors to E-boxes in the promoter and induce expression of the feedback loop repressor genes such as Period (Per1-3) and Cryptochrome (Cry1-2) [2, 3]. Current literature suggests that the disruption of the circadian system/clock genes is a risk factor in the development/progression of metabolic syndrome. However, it is not known whether obesity and HFD diet-induced antioxidant deficiency (H2S and GSH) impact the circadian desynchrony on circadian-clock genes (CCG) in the peripheral organ skeletal muscle. The impact of consuming an HFD is tissue specific. Among the peripheral metabolic signals, skeletal muscle plays a significant role in the maintenance of glucose homeostasis and is the predominant site of peripheral glucose utilization under physiological conditions [36]. Metabolic syndrome subjects have higher circulating and cellular biomarkers of oxidative stress and reduced antioxidants compared with control subjects [34, 37] and altered endogenous hydrogen sulfide (H2S) [11, 12]. H2S is one of the degradation products of an endogenous sulfur amino acid [38, 39]. This study reports that the expression of antioxidant genes and the production of H2S and GSH content decreased when the levels of oxidative stress markers increased in the skeletal muscle of HFD-fed mice and in vitro myotubes treated with high glucose, palmitate, TNF, and MCP-1. PPARγ may directly modulate the expression of several antioxidant and prooxidant genes. Catalase, a major antioxidant enzyme that converts H2O2 to H2O and O2, is a transcriptional enzyme regulated by PPARγ through the peroxisome proliferator response element (PPRE) containing the canonical DR-1 in experimental animals and humans. Also, ligand-activated PPARγ promotes the expression of manganese SOD and GPx [40]. PPARγ expression is compromised in Nrf2 null mice. Under both normal and stress conditions it may have a direct effect, and it has been demonstrated that Nrf2 induces PPARγ expression binding to at least two antioxidant response element (ARE) sequences (−916 and −784) in the upstream promoter region of the nuclear receptor PPARγ [41]. Furthermore, by inhibiting pro-inflammatory transcription factors such as nuclear factor Kappa-light-chain-enhancer of activated B cells (NF-κB), PPARγ reveals its antiinflammatory role [42]. Recent studies show that H2S positively regulates the primary cellular antioxidant glutathione (GSH) [19, 43]. HFD increases oxidative stress and decreases GSH in mice skeletal muscle [33, 34]. This data suggest that decreased antioxidant capacity and increased oxidative stress in HFD-fed mice/in vitro cell studies carried out under conditions of metabolic and inflammatory insult may be mediated through a loss of Nrf2/PPARγ signaling. PPAR integrates the mammalian clock and energy metabolism [44]. Of note, the skeletal muscle of chronic HFD-fed mice showed altered CCG and its controlled genes expression, with the majority of genes downregulated except PPARα, which was increased along with a marginal increase in Rev-erbα. The role of PPARα in skeletal muscle lipid and glucose metabolism was shown in PPARα-null mice protected from diet-induced obesity, while transgenic PPARα overexpressing mice developed glucose intolerance [45]. Loss of PGC-1α, RORα, and a minor increase in Rev-erbα may be the possible mechanism behind reduced Bmal1 expression in our study. It was shown that PGC-1α upregulates the expression of CCG, notably Bmal1 and Rev-erbα, through co-activation of the retinoic acid receptor-related orphan receptors (ROR). Mice lacking PCG-1α show abnormal diurnal rhythms of activity [46]. Also, RORα directly enhances Bmal1 transcription [47], whereas Rev-erbα inhibits it [48]. Rev-erbα binds to a ROR-responsive element in the Bmal1 promoter and represses its transcriptional activity [49]. The gene Rev-erbα is a significant regulatory component of the circadian clock [50]. Although it is not significantly altered in our study, circadian rhythms are based on organizing transcription/translation negative autoregulatory feedback loops comprising both activating and inhibiting multiple pathways. Hence, diabetes represents the dysregulation of both oxidative stress and circadian rhythm machinery [51]. These observations suggest that an alteration in metabolic status may influence the antioxidant system and thus disrupt circadian rhythms at the molecular level. In vitro studies on the mouse, myotubes demonstrated that H2S/GSH deficiency induced by CSE/GCLC knockdown could cause increased oxidative stress and decreased GSH levels. H2S affects the intracellular redox state [13, 14]. Previously, we have shown that cellular antioxidant glutathione biosynthesis is regulated by the CSE/H2S system [19]. Also, it has been reported that H2O2 can also inhibit H2S production in a concentrationdependent manner [52]. Surprisingly, H2S/GSH-deficient myotubes showed decreased Bmal1, Clock, and elevated Rev-erbα levels. The in vitro CSE knockdown data on CCG and its controlled genes were in line with our data collected from the muscle of HFD-fed mice. The contribution of H2S to the cecal gut microbes of HFD mice follows diurnal patterns, but fecal pellets showed a loss of rhythmicity [2]. Also, in the literature, diurnal physiological

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fluctuations of plasma H2S was reported in rats and mice [15, 16]. In support of these findings, an in vitro study on isolated mouse hepatocytes showed that treatment with H2S maintained expression of CCG such as Clock, Per2, Bmal1, and Rev-erbα for 48 hours post-shock treatment [14]. These data suggest that antioxidant deficiency desynchronizes CCG; hence, H2S and glutathione may be upstream of CCG and its controlled genes. In this study, treatment with either an H2S donor (NaHS) or a precursor of GSH (L-cysteine) has a positive effect on CCG and its controlled genes. The involvement of H2S during hibernation, a hypometabolic state of ‘suspended animation’ [53-55] and diurnal physiological fluctuations of plasma H2S have been reported in rats and mice [15, 16]. These findings suggest that the upregulation of physiological levels of H2S/GSH can have beneficial effects on muscle physiology by tuning the peripheral CCG and its controlled genes. This study focused on a single time point to examine the regulation of H2S on CCG in both deficient and supplemented in vitro conditions and on in vivo metabolic syndrome mice model. Future, studies on different time points are needed to strengthen the role of H2S on circadian physiology further.

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5. CONCLUSIONS In summary, these observations highlight the critical role of H2S in the muscle peripheral clock genes and thus suggest that H2S as a new endogenous factor for regulating circadian clock. These results may contribute to the understanding of how a small molecular antioxidant (H2S) or its precursors could be used therapeutically as a novel chronopharmacological approach to the management of metabolic disorders.

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DISCLOSURES None.

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ACKNOWLEDGMENTS This study was supported by a Malcolm W. Feist Cardiovascular Research Fellowship to RP and the Endowed Chair in Diabetes to SKJ from the Center for Cardiovascular Diseases and Sciences (CCDS), LSUHSC, Shreveport, as well as grants to SKJ from the National Institutes of Health/National Center for Complementary and Integrative Health (RO1 AT007442, 2013-16). We thank Ms. Paula Polk, Manager, and Dr. Wiola Luszczek, Research Specialist, at the Research Core Facility at Louisiana State University Health Sciences Center in Shreveport for their expert technical assistance. We also thank Mr. William E. McLean for lab assistance. The authors thank Ms. Georgia Morgan for excellent editing.

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FIGURE LEGENDS Figure 1. HFD attenuates the H2S mediated antioxidant system and alters circadian core clock genes (CCG) and its controlled genes expression in mouse skeletal muscle. Male C57BL/6J mice (5 weeks old) were fed either a standard chow diet (Control) or a high-fat diet (HFD) for 16 weeks. (A) The mRNA levels of H2S producing enzyme genes (CSE, CBS, and 3-Mpst) were analyzed using qRT-PCR (n=4). (B, C) The H2S production capacity of muscle extracts was measured using the lead sulfide method (n=7). (D) The mRNA levels of target genes (Nrf2, GCLC, GCLM, GSS, and GSR) were analyzed using qRT-PCR (n=3-4). (E) Glutathione levels (n=6). (F-H) hydrogen peroxide (H2O2), malondialdehyde (MDA) lipid peroxidation, and protein carbonyl levels in the muscle (n=6). (I) The mRNA levels of CCG (Bmal1, Clock, Rev-erbα, RORα, Cry2, and Per2) and its controlled genes (PPARα, PPARγ, PGC-1α, and RXRα) were analyzed using qRT-PCR (n=3-4). The unpaired Student’s t-test was used to compare the control with the HFD group. The asterisks indicate the significant difference between the control and HFD groups; *p≤0.05. Error bars represent values ±SEM.

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Figure 2. Treatment with high glucose and palmitate affects the antioxidant status and alters circadian core clock genes (CCG) and its controlled genes in myotubes. Differentiated myotubes treated with high glucose (25 mM) or palmitate (0.6 mM) for 24 h. Mannitol was used as an osmolarity control. (A) The mRNA levels of target genes (Nrf2, GCLC, and CSE) were analyzed using qRT-PCR (n=3). (B, C) The H2S production capacity of in vitro myotubes was measured using the lead sulfide method (n=4). (D-F) Glutathione levels, hydrogen peroxide (H2O2), reactive oxygen species (ROS) (n=4). (G) RT-qPCR was performed for assessment of the level of the CCG and its controlled genes expression as indicated (n=3). One-way ANOVA followed by Bonferroni or Dunnett corrections means comparison was performed between the control and treatment groups. *p≤0.05 was considered significant. Data are expressed as mean±SEM.

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Figure 3. Treatment with inflammatory cytokines (MCP-1 and TNF) decreases antioxidant status and alters circadian core clock genes (CCG) and its controlled genes in myotubes. Differentiated myotubes treated with MCP-1 (2.5 ng/mL) or TNF (250 pg/mL) for 6 h. (A) The mRNA levels of target genes (Nrf2, GCLC, and CSE) were analyzed using qRT-PCR (n=3). (B, C) The H2S production capacity of in vitro myotubes was measured using the lead sulfide method (n=4). (D-F) Glutathione levels, hydrogen peroxide (H2O2), reactive oxygen species (ROS) (n=4). (G) RT-qPCR was performed for assessment of the level of the CCG and its controlled genes as indicated (n=3). One-way ANOVA followed by Bonferroni or Dunnett corrections means comparison was performed between the control and treatment groups. *p≤0.05 was considered significant. Data are expressed as mean±SEM.

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Figure 4. Hydrogen sulfide or glutathione deficiency alters circadian core clock genes (CCG) and its controlled genes in myotubes. Myotubes were transiently transfected with CSE siRNA (H2S deficient) or GCLC siRNA (GSH deficient). Scrambled siRNA, a nonspecific RNA duplex with no sequence homology with any of the genes, served as a control. (A) The mRNA levels of target genes (Nrf2, GCLC, and CSE) were analyzed using qRTPCR (n=3). (B-E) The H2S production capacity of in vitro CSE and GCLC knockdown myotubes was measured using the lead sulfide method (n=4). (F-H) Glutathione, hydrogen peroxide (H2O2), and reactive oxygen species (ROS) levels. (I) RT-qPCR was performed for assessment of the level of the CCG and its controlled genes as indicated (n=3). (J) Myotubes have treated with CSE and CBS pharmacological inhibitors propargylglycine (PPG), aminooxyacetate (AOA) 100µM for 12 h either separate or treated together and the mRNA levels of CCG (Bmal1, Clock, Rev-erbα, RORα, Cry2, Per2) were analyzed using qRT-PCR (n=3). One-way ANOVA followed by Bonferroni or Dunnett corrections means comparison was performed between the control and treatment groups. *p≤0.05 was considered significant. Data are expressed as mean±SEM.

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Figure 5. The effect of sodium hydrosulfide and L-cysteine on myotube core clock genes (CCG) and its controlled genes. Differentiated myotubes treated with either sodium hydrosulfide (NaHS; 10 or 20 µM) or Lcysteine (LC; 300 µM) for 6 h. (A) The mRNA levels of target genes (Nrf2, GCLC, and CSE) were analyzed using qRT-PCR (n=3). (B, C) The H2S production capacity of in vitro myotubes exposed to NaHS and L-cysteine were measured using the lead sulfide method (n=4). (D-F) Glutathione, hydrogen peroxide (H2O2), and reactive oxygen species (ROS) levels. (G) RT-qPCR was performed for assessment of the level of the CCG and its controlled genes as indicated (n=3). (H) Myotubes were transiently transfected with CSE siRNA 100 nM, and post-treated with NaHS 20 µM for 6 h and the mRNA levels of CCG (Bmal1, Clock, Rev-erbα, RORα, Cry2, Per2) were analyzed using qRT-PCR (n=3). One-way ANOVA followed by Bonferroni or Dunnett corrections means comparison was performed between the control and treatment groups. *p≤0.05 was considered significant. Data are expressed as mean±SEM.

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