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Departments of
* Anesthesiology and
Endocrinology, Mayo Clinic College of Medicine, Rochester, Minnesota, USA; and
Paul F. Glenn Laboratories for the Biological Mechanisms of Aging, Harvard Medical School, Boston, Massachusetts, USA
1Correspondence: Department of Anesthesiology, Mayo Clinic College of Medicine, Rochester, MN, 55905 USA. E-mail: chini.eduardo{at}mayo.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONREFERENCES |
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axis, a well-established cascade, involved in the regulation of mitochondrial biogenesis and energy homeostasis. Thus, together these results identify a novel pathway regulating body weight and clearly show that CD38 is a nearly obligatory component of the cellular cascade that led to diet-induced obesity.—Barbosa, M. T. P., Soares, S. M., Novak, C. M., Sinclair, D., Levine, J. A., Aksoy, P., Chini, E. N. The enzyme CD38 (a NAD glycohydrolase, EC 3.2.2.5) is necessary for the development of diet-induced obesity.
Key Words: sirtuins • SIRT1 • PGC1 • nicotinamide adenine dinucleotide • knockout mice • liver
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONREFERENCES |
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coactivator, PGC1, plays a key role on the development of obesity, and energy metabolism (1
2
3
4
5)
. Activation of PGC1 by the NAD-dependent deacetylase SIRT can protect laboratory animals from high-fat diet-induced obesity and its metabolic effects (3
, 4)
. The protective effect of PGC1 appears to be mediated by increased energy expenditure due to increased mitochondrial biogenesis (3
, 4)
. In fact, the drug resveratrol, an activator of the SIRT enzymes (also known as sirtuins), has been shown to increase levels of PGC1, cellular mitochondrial numbers, and energy expenditure leading to protection of animals against high fat diet-induced obesity (3
, 4)
. However, how SIRT-PGC1 axis is modulated by endogenous pathways has not been clarified. We have previously shown that CD38 is the major NADase in key mice tissues involved in energy metabolism, including liver, brain, heart, and kidney (6
, 7)
. CD38 is a ubiquitous enzyme that catalyzes the synthesis of second messengers (1)
and has been implicated in the regulation of a wide variety of signaling pathways (8)
. Thus, in view of these, we investigated the role of CD38 as a regulator of SIRT-PGC1 axis, as well as its role in the regulation of body weight and protection against high-fat diet-induced obesity. |
MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONREFERENCES |
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and were maintained in the Mayo Clinic Animal Breeding facility in accordance with all Institutional Animal Care and Use Committee Guidelines.
Metabolic studies, animal and diet
CD38 (–/–) mice on a C57BL/6J were obtained as described previously (18)
. At 1 yr of age, mice were placed on either a normal control diet (NCD; diet number 3807, KLIBA-NAFAG), or high-fat diet (HFD) ad libitum (AIN-93G modified to provide 60% of calories from fat; Research Diets, New Brunswick, NJ, USA) and monitored for 6 wk. Body weight was recorded weekly, and food intake was measured for 7 consecutive days. Oxygen consumption and RER measurements were performed in mice fed HFD for 4 wk, and quantification of blood metabolites was performed as described in the supplemental material. For the resveratrol and sirtinol experiments, the HFD was supplemented with daily intraperitoneal injections of 30 mg/kg of drug (or vehicle) for 2 wk.
Cultured smooth muscle cells from CD38 wild-type and knockout mice
Smooth muscle cells were isolated using techniques previously described by Thompson et al. (10)
. Myometrium was minced in Hanks’ balanced salt solution (HBSS) containing 10 mM glucose and 10 mM HEPES (pH 7.4). The tissue was then suspended in fresh HBSS, aerated with 95% O2-5% CO2, and incubated in a 37°C water bath with gentle shaking for 2 h in the presence of 20 U/ml papain and 2,000 U/ml DNase. Subsequently, the tissue was incubated for an additional 2 h at 37°C, with the addition of 1 mg/ml type IV collagenase. Myometrial cells were released by trituration, centrifuged, and suspended in smooth muscle cell basal medium (SmBM) (CC 3181; Clonetics, East Rutherford, NJ, USA) containing 5% FCS, 100 U/l penicillin, 100 µg/l streptomycin, 0.25 µg/l amphotericin B, 0.05 mg/ml insulin, and 5 ng/ml human (h) EGF. Cultures were grown and maintained in 75 cm2 plastic flasks in a humidified incubator supplied with 5% CO2-95% air at 37°C. Subcultures were obtained as needed by detaching the cells with a Ca2+/Mg2+-free HBSS solution containing 0.25% trypsin and 5 mM EDTA. Only cultures between passages 2 and 10 were used. Cells isolated by this procedure stain positive for -smooth muscle actin and negative for keratin. For experiments, cells were made quiescent by replacing the growth medium with SmBM without serum or growth factors. Cell medium was again replaced with SmBM containing testing agents solubilized in 0.1% DMSO or water added to the final concentrations.
Western blot for P53 and SIRT1
Wild-type and CD38 knockout liver, brain, heart, and skeletal muscle were surgically removed, washed three times in ice-cold Hanks’ balanced salt solution (HBSS), and homogenized in 40 mM Tris–HCl (pH 7.4) containing 0.25 M sucrose using a Dounce homogenizer). Homogenates from smooth muscle cells in culture were prepared by scraping the cells in the presence of homogenizing buffer. The homogenates were centrifuged at 10,000 g for 10 min, and the resultant supernatant assayed for protein using the DC protein assay (Bio-Rad, Hercules, CA, USA). The lysates (1000 µl) were adjusted to contain 1 mg protein and 10 µg mouse monoclonal antip53 antibody conjugated to agarose beads (Santa Cruz Biotechnology, Santa Cruz, CA, USA) added overnight at 4°C with gentle rocking. The antibody–protein complex was centrifuged at 1000 g for 2 min, the pellet washed 4x in sucrose buffer and resuspended in 30 µl of sucrose buffer and 30 µl Laemmli buffer. The supernatants were denatured at 100°C for 3–5 min, and 50 µl of sample was subjected to SDS–PAGE using the Criterion Gel System (Bio-Rad) and a 4–15% gradient gel. The gels were run at a constant current of 200 V for 70 min followed by transfer to PVDF membranes (Bio-Rad). The membranes were blocked for 1 h in 5% nonfat dried milk in TBS containing 0.1% Tween 20 followed by incubation with antiacetylated p53 rabbit polyclonal antibody (1:1000) (Abcam, Cambridge, MA, USA) overnight at 4°C with gentle rocking. The membrane was then probed with an HRP-conjugated donkey anti-rabbit antibody (1:10,000) (Santa Cruz Biotechnology) for 1 h at room temperature. Blots were visualized by exposing them to BioMax MR film (Eastman Kodak Co, Rochester, NY, USA) using Supersignal Substrate (Pierce Biotechnology, Rockford, IL, USA). SIRT 1 Western blot analysis was performed using a SIRT1-specific antibody from Upstate Biotechnology (Charlottesville, VA, USA).
Immunostaining for acetylated p53 using confocal microscopy
Smooth muscle cells were obtained as described above. Cells were fixed in suspension in PBS plus 2% paraformaldehyde for 20 min at room temperature with constant agitation, and cells were washed three times for 10 min with TBS–Triton X-100 0.1%. After that, cells were incubated in blocking buffer (TBS, 30 mM glycine, and 5% BSA) for 60 min. Then incubated with antiacetylated p53 rabbit polyclonal antibody (1:1000) (Abcam) overnight at 4°C with constant agitation. Primary antibody was removed, and cells were washed three times for 10 min with TBS-T. Finally, cells were incubated for 1 h with secondary antibody (donkey anti-rabbit antibody; Molecular Probes, Eugene, OR, USA) in blocking buffer at room temperature.
Laser confocal images were obtained using the Olympus Fluoview laser scanning confocal microscopy, with objective Olympus UplanApo oil, x100, 1.35 numerical aperture. Nuclei were excited at 543 nm, and their emission was recorded at 570 nm.
SIRT 1 activity
SIRT1 activity was determined using the SIRT1 Fluorimetric Kit (Biomol International, LP, Plymouth Meeting, PA, USA), according to the manufacturer’s instructions. Briefly, nuclei prepared from wild-type or CD38 knockout mice (1 µg protein/well) were incubated in 40 mM Tris–HCl (pH 7.4) containing human recombinant SIRT1 (1 U/assay), 500 µM NAD+, and 100 µM Fluor de Lys-SIRT1 substrate for 30 min at 37°C. Following incubation, the reaction was terminated by the addition of a solution containing Fluor de Lys Developer and 2 mM nicotinamide. Values were determined by reading fluorescence on a fluorimetric plate reader (Spectramax Gemini XPS, Molecular Devices, Sunnyvale, CA, USA) with an excitation wavelength of 360 nm and emitted wavelength of 460 nm. Calculation of net fluorescence included the subtraction of a blank consisting of buffer containing no NAD+ and expressed as a percentage of control. Endogenous SIRT activity was determined as described above with the exception that no recombinant SIRT enzyme was added to the reaction mixture and that the nuclear preparations were sonicated before the assay. The data are expressed as NAD stimulated deacetylation. No deacetylation was observed in the absence of NAD in both wild-type and CD38 knockout mice nuclei.
Nuclei isolation
Mouse nuclei were isolated as described before (7)
with minor modifications. All of the steps of the preparation were performed at 4°C. The tissues were excised and washed 5 times with 20 ml of ice-cold TKM solution (50 mM Tris–HCl, pH 7.4, 25 mM KCl, and 5 mM MgCl2) to remove blood cells. Tissues were then cut into small pieces and thoroughly homogenized (10 strokes) in 5.0 ml TKM solution supplemented with 0.25 M sucrose (homogenizing medium) using a Dounce homogenizer. The homogenate was filtered through three layers of cheesecloth and centrifuged at 800 g for 10 min. The pellet was homogenized in the same volume of medium (five strokes) and centrifuged again at 800 g for 10 min. The resulting pellet was resuspended in 1.0 ml of medium (five strokes) and added to the top of a sucrose gradient containing (from top to bottom) 0.5 ml each of TKM solution with the following concentrations of sucrose, respectively: 1.0, 1.5, and 2.1 M. The tubes were centrifuged in an SW 55 Beckman rotor at 70,000 g for 60 min. The resulting pellet was resuspended in homogenizing medium and centrifuged at 800 g for 10 min. The final pellet containing the purified nuclei was resuspended in homogenizing medium (five strokes) at a protein concentration of 2.5 mg/ml and stored at –70°C until used. Protein concentration was determined using the Dc protein assay (Bio-Rad). The purity of nuclear preparations was determined as described before (7)
; nuclear preps were found to be nearly 100% pure.
Chromatin immunoprecipitation assay
Chromatin immunoprecipitation assay (ChIP) was performed in cultured smooth muscle cells as previously described (11)
. Briefly, cells from wild-type and CD38 knockout animals were cross-linked with formaldehyde for 20 min at 25°C, harvested in SDS-lysis buffer (Upstate Biotechnology), and sheared to fragment DNA (<500 bp). Samples were then immunoprecipitated using an agarose-conjugated antip53 antibody (Santa Cruz Biotechnology) or IgG control together with agarose beads at 4°C overnight. Following immunoprecipitation, samples were washed and eluted using the chromatin immunoprecipitation kit (Upstate Biotechnology) according to the manufacturer’s instructions. Cross-links were removed at 65°C for 6 h and immunoprecipitated DNA was purified using phenol/chloroform extraction and ethanol precipitation. A 260 bp region flanking p53 binding site in the mouse p21 promoter (12
, 13)
was detected in immunoprecipitated samples by PCR. PCR products were visualized on a 2% agarose gel.
ADP-ribosyl cyclase and NADase activity
ADP-ribosyl cyclase activity was measured using the NGD technique as described previously, and NADase activity was determined using etheno-NAD as described before (6)
. Enzyme preparations were incubated in a medium containing 0.2 mM NGD, 0.25 M sucrose, and 40 mM Tris–HCl (pH 7.2) at 37°C. Activity was determined by measuring the change in fluorescence over time at 300-nm excitation and 410-nm emission.
Cyclic-ADP-ribose induced Ca2+ release
cADPR-induced Ca2+ release was determined in sea urchin egg homogenates using fluo-3 as a Ca2+ indicator as extensively described before (14)
. Homogenates from Lytechinus pictus egg were prepared as described previously (14)
. Frozen homogenates were thawed in a 17°C water bath and diluted to 1.25% with an intracellular medium containing 250 mM N-methyl glutamine, 250 mM potassium gluconate, 20 mM HEPES buffer, pH 7.2, 1 mM MgCl2, 2 U/ml creatine kinase, 4 mM phosphocreatine, 1 mM ATP, 3 µg/ml oligomycin, and 3 µg/ml antimycin. After incubation at 17°C for 3 h, 3 µM fluo-3 was added. Fluo-3 fluorescence was monitored at 490-nm excitation and 535-nm emission in a 250-µl cuvette, held at 17°C with a circulating water bath and continuously mixed with a magnetic stirring bar, in a Hitachi spectrofluorometer (F-2000).
Detection of NAD by cycling assay
Mouse tissues were frozen in liquid N2, pulverized into a powder, and extracted with 10% trichloroacetic acid (TCA) at 4°C. TCA was removed with water–saturated ether. The aqueous layer containing the NAD was removed and adjusted to pH 8 with 1 M Tris. Detection of NAD was determined as described before (6
, 7)
.
PGC1 analysis was performed by direct Western blot analysis using a rabbit anti-PGC1 antibody (H-300, Santa Cruz Biotechnology).
Fecal analysis
Daily fecal outputs were determined through the use of metabolic cages and were measured by collecting all feces from each individual mouse (3 or 4 mice/day) for 7 days. Feces were allowed to air dry at least 72 h before weighing. Total lipids were extracted by the Folch method (15)
.
Histological analysis
Histological analysis, including hematoxylin & eosin staining (H&E) and electron microscopy (EM) were performed using standard techniques as described before (3)
. Mitochondria in EM images were quantified using Image J version 1.36b. Histological specimens were analyzed by three different pathologists that classified the liver tissue as normal, mild, moderate, or severe steatotic. All 3 independent clinical pathologists agree with the diagnosis of moderate steatosis in liver samples from wild-type mice on a high-fat diet.
Biochemical markers, hormones, and fatty acids
Plasma levels of nonesterified fatty acids were measured using the NEFA C kit (Wako Chemicals, Richmond, VA, USA) according to the manufacturer’s instructions. Leptin was measured with using an ELISA kit (R&D Systems, Minneapolis, MN, USA).
Oxygen consumption, carbon dioxide production, and activity
We acclimated each mouse to a 30 cm x 10 cm cylindrical chamber for 24 h prior to the measurement of 24-h energy expenditure (EE). For measurements of EE, we used a small animal calorimeter, and for measurements of physical activity, we used Opto-M Varimex Minor activity monitor, which uses an infrared beam that breaks in 3 axes to measure activity (Columbus Instruments, Columbus, OH, USA) (16)
. Physical activity data were collected each minute in 3 axes using infrared beam break counts (ambulatory=nonrepetitive horizontal beam breaks; total counts=horizontal+ambulatory+vertical counts; stationary counts=ambulatory-horizontal). We set the calorimeter to deliver room air 0.60–0.65 LPM to the chamber and collect samples every min (with a 5-min reference period every 30 samples) with a sample flow of 0.4 LPM. We calculated resting energy expenditure (REE) by averaging the EE values associated with 0 activity counts for the minute of the EE measurement and the prior 4 min. Activity EE (EEA) was calculated by subtracting REE from TEE.
Oxygen consumption and carbon dioxide production were measured by using a customized, high-precision, single-chamber indirect calorimeter (Columbus Instruments) as we have reported previously (16)
. Thermogenesis was calculated from oxygen consumption and carbon dioxide production. Calibration of the calorimeter was performed at the beginning of each measurement day. The animal was placed inside the cylindrical calorimeter chamber (acrylic; diameter 30 cm, height 20 cm, volume 15 l). The chamber lid was attached and sealed, and room air was pumped at atmospheric pressure through the chamber at 3.4–3.7 l/min. Data on oxygen consumption and carbon dioxide production were then collected every minute and stored on a PC. Each data-point was identified by a time stamp. Ambulation was measured simultaneously with the oxygen consumption and carbon dioxide production measurements. Measurements were performed using customized, high-precision racks of collimated infrared activity sensors (Columbus Instruments) placed around the acrylic chamber. There were 45 collimated beams of infrared light crossing the 30-cm-diameter cage, allowing the detection of 1 inch of movement in three orthogonal axes. Photosensors registered an activity unit each time a beam was interrupted. In this fashion, activity was simultaneously detected in all three axes: forward-and-backward, side-to-side, and up-and-down. Data for ambulation were summed for every minute and stored on the PC with use of the time stamp for identification. Data were thereby derived simultaneously for oxygen consumption and ambulation, for each animal, every minute over the 24-h measurement period. Animals were acclimated for 24–48 h before the measurements.
Materials
All other reagents, of the highest purity grade available, were supplied from Sigma Chemical (St. Louis, MO, USA), except when stated otherwise. L. pictus and Aplysia californica were obtained from Marinus Inc. (Long Beach, CA, USA). Fluo-3 was purchased from Molecular Probes. All other reagents, of the highest purity grade available, were supplied by Sigma, and cADPR were synthesized as described before (12)
.
Statistical analysis
Data are presented as means ± SEM. The main and interactive effects were analyzed by analysis of variance (ANOVA) factorial, repeated measurements or by one-way ANOVA Differences between individual groups means were analyzed by Fisher’s test. Analyses were performed using (sigma plot statistics)
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONREFERENCES |
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pathway
A–C). Using acetylation status and transcriptional activity of the SIRT target p53, as a surrogate marker of endogenous sirtuin activity, we clearly show that the in vivo sirtuin activity is increased not only in muscle (Fig. 1A
), but in liver (Fig. 1B
), brain (Fig. 1D
) and heart (data not shown). Using a combination of expression and chromatin immunoprecipitation and reporter assays, we observed that in CD38 (–/–) mice, cells levels of acetylated p53 are decreased (Fig. 1E
), and its functional activity is impaired as demonstrated by decrease binding to the promoter of one its target genes, p21. Furthermore, cells from CD38 (–/–) show decrease p53-mediated p21 promoter activity (Fig. 2
). Recent reports demonstrate that treatment with resveratrol leads to SIRT activation (3
, 4)
, which, in turn, deacetylates PGC1 and also increases its intracellular levels (3
4
5)
. Consistent with the SIRT activation, levels of PGC1 were increased severalfold in liver from CD38 (–/–) mice (Fig. 1F
). Thus, in view of these findings and the large body of evidences that implicates SIRT-PGC1 as a regulator of obesity (1
2
3
4
5)
, we explored the effect of CD38 knockout on high-fat diet-induced obesity.
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Protective effect of CD38 deficiency against high-fat diet-induce obesity
To determine the role of CD38 on high-fat diet-induced obesity, we study fully developed middle-aged mice. As animals reach adulthood, their growth rate decreases, and fat is stored as an energy reserve in adipose tissue, thus leading to obesity. We have followed wild-type and CD38 (–/–) mice for over one year. Adult wild-type and CD38 (–/–) mice maintained on standard chow diet (4% total calories derived from fat, 3.04 kcal g–1) differ slightly regarding weight. The CD38 (–/–) mice were consistently thinner by 
3g (Table 1
). Furthermore, the abdominal and inguinal adipose tissue was also reduced in CD38 (–/–) mice (Table 1)
. This decrease in fat accumulation in CD38 (–/–) mice was not due to lower food intake or malabsorption (Table 1)
, as food intake, fecal output, and fat content were not significantly different between both mouse genotypes (Table 1)
. In all other tests, biochemical parameters, including insulin, leptin, adiponectin, glucose, ketone body formation, and free fatty acids levels, did not differ between the wild-type and CD38 (–/–) mice.
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To further evaluate the role of CD38 on obesity, we challenged mice with high caloric-fat diet (60% of calories from fat, 5.05 kcal g–1). Indeed, when CD38 (–/–) mice were fed with a high-fat diet, weight accumulation was nearly absent when compared with wild-type mice during the feeding period (Fig. 3
). To determine the amount of weight gain corresponding to fat, mice were sacrifice at 4 wk of the high-fat diet, and the total abdominal and inguinal fat was dissected (Table 1
, Figs. 3
and 4
). Assessment of total abdominal and inguinal fat pads, revealed that the body fat in CD38 (–/–) mice did not increase over this period, whereas it nearly quadrupled for wild-type mice (Table 1)
. Furthermore, microscopic evaluation of adipocytes in wild-type and CD38 (–/–) cells indicates a larger accumulation of fat in wild-type cells. In addition, liver size increases in wild-type due to fat infiltration but were not altered in CD38 (–/–) mice (Table 1
, Figs. 3
and 4
). In fact, wild-type mice placed on a high-fat diet for 6 wk show moderate liver steatosis (Fig. 4
). In addition, fat accumulation in other organs like skeletal muscle was also prevented by CD38 knockout mice (Fig. 4)
.
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Protective effect of CD38 knockout against high-fat diet-induced glucose intolerance
One of the important metabolic consequences of obesity is the development of diabetes and glucose intolerance (2
3
4)
. CD38 has been implicated in insulin release from ß-cells and the development of diabetes (17)
. Okamoto and colleagues (17)
have proposed that glucose-induced insulin release is mediated by CD38 generation of cADPR, which, in turns, causes intracellular Ca2+ release (17
, 18)
. However, subsequent experiments failed to show an effect of CD38 and cADPR on glucose-induced insulin release from ß-cells (19
, 20)
. Furthermore, it has been shown that in ß cells deficient in CD38 and cAPDR, the second messenger IP3 can modulate intracellular Ca2+ release and insulin secretion (18)
. These data together indicate that CD38 and cADPR may not be necessary for insulin secretion and ß cell-mediated control of glucose metabolism. Some authors have further proposed that the role of CD38 on glucose homeostasis may be indirect (20)
. Here, we explore the role of CD38 on the development of glucose intolerance induced by a high-fat diet (Fig. 5
). In agreement with other studies, we observed that wild-type and CD38 (–/–) mice fed with standard chow did not differ on the glucose tolerance (data not shown) (20)
. In contrast, after 8 wk of a high-fat diet, wild-type mice become glucose intolerant. In contrast, no significant changes were observed on glucose tolerance test in CD38 (–/–) mice (Fig. 5)
. These data support the notion that CD38 is not necessary for insulin secretion or maintenance of normal glucose metabolism. In contrast, we show for the first time a beneficial effect of CD38 (–/–) in glucose metabolism in vivo.
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Enhanced energy expenditure in CD38 null mice
Although food intake in CD38 (–/–) mice was similar to that of wild-type mice, when normalized to body weight, they consume 40% more food (Table 1)
. Moreover, on the basis of similar fecal output and fecal fat content, the protection against weight gain of the CD38 (–/–) mice was not due to malabsorption of fat (Table 1)
. Therefore, these data indicate that the CD38 (–/–) mice may have higher energy expenditure than wild-type mice. Thus we investigated the energy homeostasis in CD38(–/–) and wild-type mice, fed a high-fat diet. Using a high-precision indirect calorimeter, we determine that the CD38 (–/–) mice had significantly higher oxygen consumption (VO2) and energy expenditure (EE) corrected for body weight compared to the wild type mice (Table 2
). Resting energy expenditure (REE) corrected for body weight also significantly greater in the CD38 knockout mice (Fig. 6
). The wild-type mice exhibited greater amounts of physical activity (horizontal, vertical, ambulatory, and total activity counts) compared to the CD38 knockout mice (Fig. 6
, Table 2
). These data are in concordance with recent findings demonstrating that mice treated with resveratrol (4)
, a SIRT1 activator, significantly decreased their activity levels. Respiratory quotient (RQ) was significantly higher in the CD38 (–/–) compared to wild type mice. Energy expenditure of activity (EEA) was similar between the groups of mice, but EEA per activity count (horizontal+vertical) was significantly higher in the CD38 (–/–) mice, almost twice the value of the wild-type mice, suggesting that the calculated efficiency of movement was lower in the CD38 knockout mice (Fig. 6)
. Thus, these changes contribute to the ability of these mice to fend off weight gain during high-fat diet feeding, even with decreased ambulation. Thus, together, these results suggest that CD38 deficiency’s protective effect against high-fat diet-induced obesity is mediated by enhanced energy expenditure.
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The mechanism that underlines the increase in energy expenditure in CD38 (–/–) mice is probably mediated by augmented PGC1 activity and its downstream effects on energy metabolism and mitochondrial biogenesis (1
2
3
4)
. Because in CD38 (–/–) mice, both basal and activity-related metabolic rates are increased, it is possible that more then one tissue may be involved in changes in energy expenditure. Skeletal muscle tissue is one of the likely candidates because of its role in basal and activity-related energy expenditure. We investigate here the morphological changes in mitochondria in skeletal muscle. As shown in Figs. 4
and 7
, the size and number of the mitochondria in gastrocnemius muscle from CD38 (–/–) are increased compared to wild-type mice. In fact, the calculated mitochondrial area was increased 2.5 times in CD38 (–/–) (Fig. 7
), an observation compatible with role of skeletal muscle on the increase in energy expenditure in C38 (–/–) mice. These data further supports the notion that the effect of CD38 on obesity and energy expenditure are mediated by the SIRT1-PGC1 pathway.
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CD38 deficiency protective effect on high-fat diet-induced obesity requires an intact SIRT activity
CD38 has been implicated as an enzyme responsible for the synthesis of the second messenger cyclic-ADP-ribose (9)
. However, recently, it has been reported that CD38 can modulate different signaling cascades by regulating intracellular NAD level (6
, 7)
. To further determine whether the regulation of SIRT by CD38 was mediated by cADPR-induced Ca2+ release or modulation of the activity of SIRT, we determined the effect of pharmacological agonists and antagonists of the SIRT system on cADPR synthesis and cADPR-induced Ca2+ release (Fig. 8
). We observed that neither resveratrol (a SIRT activator) (3
, 4
, 17)
nor sirtinol (a SIRT inhibitor) (21)
had any effect on these parameters. Moreover, we investigated the effect of SIRT and cADPR pharmacologic modulators on in vivo acetylation of p53 as a measure of endogenous SIRT activity. As expected, sirtinol increased acetylated p53 in CD38 (–/–) cells, and resveratrol decreased it in wild-type cells (Fig. 8)
. In contrast, neither the cell-permeable cADPR antagonist (8-br-cADPR) nor the cADPR agonist (3-deaza-cADPR) (9)
had any effect on the levels of p53 acetylation in cells (Fig. 8)
.
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Finally, we treated wild-type mice with 30 mg/kg/day of resveratrol and CD38 (–/–) with the same dose of sirtinol to determine the role of the SIRT enzymes on high-fat diet-induced obesity. In agreement with a recent report (4)
, we observed that wild-type mice treated with resveratrol for 2 wk were protected against high-fat induced obesity (Fig. 9
). In contrast, the protective effect of CD38 knockout on high-fat diet-induced obesity was abrogated by sirtinol. Sirtinol-treated CD38 (–/–) mice gain a statistically significant amount of weight when compared with nonsirtinol (vehicle)-treated CD38 (–/–) mice (Fig. 9)
. These data support the novel notion that CD38 modulates high-fat diet-induced obesity by a sirtuin-dependent mechanism.
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CONCLUSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONREFERENCES |
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, which, in turn, regulates energy metabolism and obesity (22)
. Our studies provide a novel and crucial physiological role for CD38 and also a new mechanism of action for this enzyme. Further, studies should focus on the role of CD38 a sensor and regulator of cellular metabolism. The role of CD38 on glucose tolerance is controversial since both no changes on decrease glucose tolerance have been described in CD38 (–/–). This may be explained by different mice backgrounds (17
, 18
, 20)
. Finally, our studies may have relevance for human physiology, since in a recent study, a genetic linkage has been observed between chromosome 4 near marker D4S403 where the CD38 gene is located and the metabolic syndrome that refers to the clusteringof disease conditions such as obesity, insulin resistance, hyperinsulinemia, dyslipidemia, and hypertension (23)
.
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ACKNOWLEDGMENTS |
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Received for publication March 16, 2007. Accepted for publication May 17, 2007.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONREFERENCES |
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: key regulator of energy metabolism. Adv. Physiol. Educ. 30,145-151. Int. J. Obes. 29,S5-S9[CrossRef]. Cell. 127,1109-1122[CrossRef][Medline] and SIRT1. Nature. 434,113-118[CrossRef][Medline]This article has been cited by other articles:
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