Kinsenoside-mediated lipolysis through an AMPK-dependent pathway in C3H10T1/2 adipocytes: roles of AMPK and PPAR[alpha] in the lipolytic effect of kinsenoside.
Background: Currently, more than one-third of the global population is overweight or obese, which is a risk factor for major causes of death including cardiovascular disease, numerous cancers, and diabetes. Kinsenoside, a major active component of Anoectochilus formosanus exhibits antihyperglycemic, antihyperliposis, and hepatoprotective effects and can be used to prevent and manage obesity.
Purpose: This study examined the catabolic effects of kinsenoside on lipolysis in adipocytes transformed from C3H10T1/2 cells.
Study design/methods: The lipolytic effect of kinsenoside in C3H10T1/2 adipocytes was evaluated by oil-red O staining and glycerol production. The underlying mechanisms were assessed by Western blots, chromatin immunoprecipitation (IP), Co-IP, EMSA and siRNAs verification.
Results: We demonstrated that kinsenoside increased both adipose triglyceride lipase (ATGL)-mediated lipolysis, which was upregulated by AMP-activated protein kinase (AMPK) activation, and the hydrolysis of triglycerides to glycerol and fatty acids that require transportation into mitochondria for further [beta]-oxidation. We also demonstrated that kinsenoside increased the phosphorylation of peroxisome proliferator-activated receptor alpha (PPAR[alpha]) and CRE-binding protein (CREB), and the protein levels of silent information regulator T1 (SIRT1), peroxisome proliferator-activated receptor gamma coactivator-1 alpha (PGC-1[alpha]) and carnitine palmitoyltransferase I (CPT1) through an AMPK-dependent mechanism. S1RT1 deacetylated PGC-1[alpha], facilitating AMPK-mediated PGC-1[alpha] phosphorylation and increasing the interaction of PPAR[alpha] with its coactivator, PGC-1[alpha]. This interaction elevated the expression of CPT1, a shuttle for the mitochondrial transport of fatty acids, in kinsenoside-treated cells. In addition, AMPK-phosphorylation-mediated CREB activation caused kinsenoside-mediated PGC-1[alpha] upregulation.
Conclusion: AMPK activation not only elevated ATGL expression for lipolysis but also induced CPT1 expression for further mitochondrial translocation of fatty acids. The results suggested that the mechanism underlying the catabolic effects of kinsenoside on lipolysis and increased CPT1 induction was mediated through an AMPK-dependent pathway.
AMP-activated protein kinase
Obesity, a common problem in developed countries, may be due to metabolic disorders or energy overconsumption, which is closely associated with several diseases, including stroke, heart attack, and diabetes mellitus (DM). Anoectochilus formosanus Hayata (Orchidaceae), a species endemic to Taiwan, has been traditionally used in herbal medicine for patients with hypertension, DM, pulmonary tuberculosis, and nephritis (Tseng et al. 2006). Kinsenoside, 3-0-[beta]-D-glucopyranosyl-(3R)-hydroxybutanolide, is a major active component of A. formosanus and exhibits immunostimulating (Tseng et al. 2006), antihyperglycemic (Liu et al. 2013; Zhang et al. 2007), antihyperliposis (Du et al. 2001; Du et al. 2008), and hepatoprotective (Hsieh et al. 2011) activities. However, these studies have indicated that kinsenoside might alleviate complications associated with carbohydrate and lipid metabolism disorders. Six-week oral treatment with kinsenoside markedly reduced body and liver weights in aurothioglucose-induced obese mice (Du et al. 2008). Histopathological examinations revealed no substantial damage. However, the mechanism underlying the effects of kinsenoside requires further investigation.
AMP-activated protein kinase (AMPK) acts as a master regulator of cellular energy homeostasis. Under energy stress, AMPK is activated in cells by an increase in the AMP/ATP ratio, initiating metabolic and genetic events to restore ATP levels (i.e., fatty acid [FA][beta]-oxidation) and reduce ATP-consuming processes (i.e., triglyceride and protein syntheses). Extra energy stored in adipocytes causes lipid accumulation. Several studies have investigated whether natural products inhibit adipogenesis or enhance lipolysis to reduce the amount of adipose tissue. A major component of flavonoids in green tea, (-)-epigallocatechin gallate (EGCG), was shown to significantly reduce insulin-stimulated glucose uptake through the 67LR, known as the EGCG receptor, and AMPK pathways at 5-10 [micro]M in 3T3-L1 and C3H10T1/2 adipocytes (Hsieh et al. 2010). Other natural compounds, such as genistein in soy, xanthohumol in beer hops, resveratrol in grapes, and curcumin in turmeric, were reported to exhibit adipogenesis inhibition activities by reducing peroxisome proliferator-activated receptor gamma (PPAR[gamma]) expression and/or activating AMPK in adipocytes (Andersen et al. 2010). In adipose tissue, AMPK activation is concomitant with lipolysis (Yin et al. 2003) and increased FA oxidation (FAO) (Minokoshi et al. 2002). A previous study showed that AMPK inactivated acetyl-CoA carboxylase (ACC) and reduced lipogenesis (Matejkova et al. 2004). This induces an increased capacity for FAO that could be due to a decreased concentration of malonyl-CoA, attenuating the inhibition of carnitine palmitoyltransferase I (CPT1) that catalyzes the entry of FAs in mitochondria, a rate-limiting enzyme of FAO (Assifi et al. 2005).
Sirtuins are a family of [NAD.sup.+]-dependent histone/protein deacetylases that are also fuel-sensing molecules. AMPK and silent information regulator T1 (SIRT1) regulate each other and share many common target molecules such as peroxisome proliferator-activated receptor gamma coactivator-1 alpha (PGC-1 a) (Ruderman et al. 2010). Adipose tissue is essential for energy regulation in the body and provides free FAs as an energy source to other tissues when energy is scarce. The adipose tissue metabolism is subjected to various regulatory mechanisms including those involving hormones and neurotransmitters. For example, catecholamine rapidly activates the [beta]-adrenergic receptor and the cAMP-dependent protein kinase A (PKA) axis (McKnight et al. 1998), thus stimulating lipolysis through a complex regulatory mechanism that appears to involve hormone sensitive lipase (HSL) and perilipin phosphorylation (Ducharme and Bickel 2008; Granneman and Moore 2008; Zimmermann et al. 2009).
Evidence increasingly suggests a causative relationship between PPAR activity and metabolic syndromes, including insulin resistance, diabetes, obesity, and dyslipidemia. Notably, PPAR[alpha] activators (i.e., the fibric acid class of hypolipidemic drugs) and PPAR[gamma] agonists (i.e., antidiabetic thiazolidinediones) are effective for improving metabolic syndrome (Mansour 2014; Zhang et al. 2005). The PPAR family seems essential for preventing type-Il-diabetes-associated complications and therefore may serve as a potential therapeutic target for treating metabolic syndrome and its related complications. The deacetylation and phosphorylation of PGC-1[alpha] by SIRT1 and AMPK, respectively, are essential for PPAR transactivation (Canto et al. 2009). Mammalian sirtuins (e.g., SIRT1) exhibiting HDAC activity not only inhibit 3T3-L1 adipogenesis through their interaction with PPAR[gamma] (Rosen and MacDougald 2006) but also control lipolysis in adipocytes through Foxol mediated ATGL1 expression (Chakrabarti et al. 2011). Adiponectin, an antidiabetic adipokine, induces AMPK, S1RT1, and PGC-1[alpha] expression and reduces PGC-1[alpha] acetylation, thus increasing the amount of mitochondria in myocytes, insulin sensitivity, and exercise endurance (Iwabuetal. 2010).
In this study, we examined the mechanism underlying the catabolic effects of kinsenoside on lipolysis, the signaling pathways in the modulation of gene regulation involved in lipolysis, and a mitochondrial transporter of FAs, CPT1.
Materials and methods
Collection of plant material and isolation of kinsenoside
A voucher specimen of A. formosanus was deposited in the herbarium of the School of Pharmaceutical Sciences, Taipei Medical University. Entire plants of freshly cultivated A. formosanus, collected from Chutung, Hsinchu, Taiwan, were extracted with 10 vol methanol at room temperature. After the filtrate was evaporated under reduced pressure, the obtained crude extract was partitioned with water and ethyl acetate (1:1). The water-soluble fraction was then partitioned with water and n-butanol (1:1). Chromatographing the n-butanol fraction on a silica gel column eluted with ethyl acetate/methanol/trifluoroacetate yielded seven fractions. To isolate kinsenoside in fraction 1, a semipreparative reverse-phase highperformance liquid chromatography (HPLC) column (VP 250/10 NucleodurC1810 mm x 250 mm) equipped with a refractive index detector was used at a flow rate of 3 ml/min, with water used as the mobile phase. Kinsenoside was eluted at 19.3 min, and the purified kinsenoside was subjected to NMR analysis. The results are summarized in Table SI; 13C NMR (pyridine-d5): 8 36.5 (C-2), 63.3 (G-6), 72.1 (G-4), 75.5 (G-2), 75.8 (C-4), 76.2 (C-3), 78.9 (G-5), 79.1 (G-3), 104.6 (G-l), 177.3 (C-l). The 1H and 13C spectrum (Figs. SI and S2) was consistent with the results published by Zhang et al. (2014); therefore, the sample was identified as kinsenoside.
The purity of kinsenoside was determined using LC-UV (Fig. S3). Specifically, the purified kinsenoside was dissolved in MeOH to 3.5 [micro]g/ml as the test sample. The absorbance of the sample was determined at 210 nm by using ultraviolet spectrometry. Finally, the purity of kinsenoside was calculated to be 92.0%.
Preparation of adipocytes from C3H10T1/2 cells
Mouse embryonic bone marrow stem cells (C3H10T1/2) (ATCC) were maintained in a low-glucose Dulbecco's modified Eagle medium (DMEM) supplemented with 10% (v/v) fetal bovine serum (FBS) (Atlanta Biologicals), 100 U/ml penicillin, and 100 mg/ml streptomycin (Invitrogen, Carlsbad, CA, USA) and were transformed into adipocytes in an adipogenic induction medium supplemented with 0.5 m M 3-isobutyl-l-methylxanthine (a phosphodiesterase inhibitor), 1 [micro]M dexamethasone, and 10 [micro]g/ml insulin in high-glucose DMEM supplemented with 10% FBS for 3 d. For lipid droplet accumulation, the cells were maintained in 10 [micro]g/ml insulin in high-glucose DMEM supplemented with 10% FBS for an additional 6 d and then subjected to insulin withdrawal for 2 d to exhaust insulin signaling. Subsequently, the cells became round and exhibited lipid droplets, as observed under an Olympus CX40 microscope (Center Valley, PA, USA) at 200 x magnification. After cell differentiation, the accumulated fat was stained using oil-red O (Sigma, St. Louis, MO, USA), the optical density was measured at 510 nm by using a spectrophotometer, and the ADP/ATP ratio was determined using the ADP/ATP ratio assay kit (Abeam, Cambridge, England, UK) according to the manufacturer's protocol.
Evaluation of fat accumulation using oil-red O staining and lipolysis using a glycerol assay
Adipocytes cultured in 24-well petri dishes containing DMEM at 500 [micro]l/well were pretreated with the indicated activators or inhibitors of the proposed signaling molecules in lipolysis. Kinsenoside treatment was then performed. The effects of the activators or inhibitors on kinsenoside-mediated lipolysis were determined according to the amounts of fat deposited and glycerol secreted from the cells. In the oil-red staining assay, the cells subjected to the indicated treatments were fixed with 10% formalin for 10 min at room temperature, washed with 100% isopropanol, dried completely, and incubated with 200 [micro]l of the oil-red O working solution (0.35% stock oil-red 0 and distilled water; 3:2) for 30 min in the dark. The cells were then washed four times with double-distilled [H.sub.2]O (dd[H.sub.2]O), dried, and subjected to oil-red extraction using 200 [micro]l of isopropanol. A colorimetric assay was performed at 500 nm, with 100% isopropanol used as a blank. To quantify the amount of lipid uptake, the oil-red O stain was eluted with 100% isopropanol for 10 min, and the absorbance was measured at 510 nm on a microplate spectrophotometer by using Gen5 software (BioTek Instruments, Winooski, VT, USA). Six microliters of a culture medium were aspirated and examined for the secreted glycerol by using quantitative enzymatic colorimetric assay kits (Randox Laboratories Ltd. Co., Antrim, UK). The color development was measured at 520 nm and presented as folds of induction compared with the control.
Lactate dehydrogenase cytotoxicity assay
To determine the optimal concentration required for lipolysis, adipocytes grown in 24-well petri dishes were treated with increasing kinsenoside concentrations from 10 [micro]g/ml to 1 mg/ml for 24 h. Fifty microliters were aspired from 500 [micro]l/well and mixed with 50 [micro]l of the lactate dehydrogenase (LDH) substrate mixture supplied in the CytoTox 96 NonRadioactive Cytotoxicity Assay Kit (Promega, Madison, WI, USA) for 30 min in the dark. LDH released in the medium catalyzed lactate dehydration with the conversion of NAD+ to NADH. The released hydrogen ions reduced the tetrazolium salt to form a red formazan product. The reaction was stopped by adding 50 [micro]l of a stop solution, and the level of the formazan product was determined by measuring its absorbance at 490 nm by using a spectrophotometer in 96-well plates (Tecan, Grodig, Austria).
Western blotting of whole cell lysates, nuclear-cytosolic fractionation, and coimmunoprecipitation
Adipocytes grown in 10-[cm.sup.2] dishes were pretreated with an AMPK inhibitor (cpdC) or a SIRT1 inhibitor (sirtinol) for 1 h and then treated with kinsenoside for 1 h. Cell lysates were separated into cytosolic and nuclear fractions by using NE-PER[TM] nuclear extraction reagents (Pierce Biotechnology, Inc., Rockford, IL, USA) supplemented with protease inhibitors. The PGC-1[alpha] protein was immunoprecipitated in samples containing 200 pg of total protein by using 2 [micro]g of an antiPGC-1 a antibody and 20 [micro]g of protein A plus G agarose beads to determine whether the SIRT1 and PPAR[alpha] proteins coprecipitated with the PGC-1[alpha] protein. The precipitates were then washed five times with a lysis buffer and once with phosphate-buffered saline; resuspended in a sample buffer containing 50 mM Tris, 100 mM bromophenol blue, and 10% glycerol at pH 6.8; and incubated at 90 [degrees]C for 10 min before electrophoresis was used to release the proteins from the beads. Western blotting (Lee et al. 2010) was performed using the following: anti-PPAR[alpha], anti-ATGL, anti-PGC-1[alpha], anti-SIRTI, anticarnitine palmitoyltransferase I (anti-CPT1), anti-PPAR[gamma], anti-(p)ERKl/2, antiGAPDH, and anti-Lamin A/C antibodies (Santa Cruz Biotechnology, Dallas, TX, USA); an anti-pPPAR[alpha] (Ser21) antibody (ABR Affinity Bioreagents, Rockford, IL, USA); and anti-AMPK, anti-pAMPK (Thrl72), anti-CREB, and anti-pCREB (Seri 33) antibodies (Millipore, Burlington, MA, USA). Aliquots of the nuclear and cytosolic fractions containing 50 [micro]g of the total protein were separated on a 10% acrylamide gel by using sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Hybond-P membranes (GE Healthcare Life Sciences, Waukesha, WI, USA) containing the electrophoretically transferred protein bands were probed using the various primary antibodies. Band intensities were normalized to the GAPDH or Lamin A/C (control) band intensity by using an IS-1000 digital imaging system (ARRB, Victoria, Australia).
Chromatin immunoprecipitation and electrophoretic mobility shift assay
To examine the kinsenoside-mediated genomic regulation of PGCl[alpha], a chromatin immunoprecipitation (ChIP) assay was performed according to the instructions of Upstate Biotechnology (Lake Placid, NY, USA) with minor modifications. In brief, 6 x [10.sup.5] cells cultured in 10-[cm.sup.2] dishes and subjected to the indicated treatments were harvested. The resulting cell lysates were subjected to overnight coimmunoprecipitation (Co-IP) by using an anti-CREB antibody. DNA filtrates were amplified using PCR with the following primers flanking the PCC-l[alpha] gene promoter containing the putative CREB-binding sites: PGC-1[alpha] forward primer5'-CAAAGCTGGCTTCAGTCACA-3' and reverse primer 5'-CTTGCTGCACAAACTCCTGA-3'. The template was replaced with dd[H.sub.2]O for use as a negative internal control. The PCR products were electrophoresed on a 2% agarose gel; products of the expected size, 155 bp, were observed and quantified using Scion Image analysis.
An electrophoretic mobility shift assay (EMSA) was performed as described previously (Sue et al. 2009) with minor modifications. For nuclear protein extraction, adipocytes in 10-[cm.sup.2] dishes pretreated with kinsenoside at 50 [micro]g/ml for 30 min were treated
with NE-PER[TM] nuclear extraction reagents (Pierce Biotechnology, Inc.) supplemented with protease inhibitors. The oligonucleotide sequences used for the putative PGC-1[alpha] cAMP response element (CRE) wild type and mutant were TGCCTTGGAGTGACgtCAGGAGTTT and TGCCTTGGAGGTCAtgCAGGAGTTT, respectively (the conserved and mutated sequences are bolded and underlined, respectively). The oligonucleotides were end-labeled with biotin according to the manufacturer's protocol (Pierce Biotechnology, Inc.). In brief, unlabeled oligonucleotides (1 [micro]M) were incubated with a TdT reaction buffer containing biotin-11-dUTP (0.5 [micro]M) and TdT (0.2 U/[micro]T) at 37 [degrees]C for 30 min; subsequently 2.5 [micro]l of EDTA (0.2 M, pH 8.0) were added to stop the reaction and 50 [micro]l of chloroform/isoamyl alcohol were added to extract the TdT. The extracted nuclear protein (10 [micro]g) was incubated with biotin-labeled (1 pmol) probes at 15"C for 30 min in a binding buffer containing 1 [micro]g of poly deoxyinosinedeoxycytidine (dl-dC) (Panomics, Inc., Redwood City, CA, USA). For competition with unlabeled oligonucleotides, a 100-fold molar excess of the unlabeled oligonucleotides relative to the biotin-labeled probes was added to the binding assay mixture, which was separated on a 6% nondenaturing polyacrylamide gel at 4 [degrees]C in 1 x TBE (90 mM Tris borate, 2 mM EDTA, pH 8.3) and then transblotted onto a Hybond [N.sup.+] membrane (Amersham Pharmacia Biotech, Freiburg, Germany). The blots were incubated with a blocking buffer. Streptavidin-horseradish peroxidase conjugates were then added, and the blots were imaged using an enhanced chemiluminescence system.
The data are expressed as the mean [+ or -] the standard deviation (SD), representing the results of at least three experiments. Using a one-way analysis of variance, the means of the experimental and control groups were compared. P < 0.05 was considered statistically significant.
Concentration-dependent lipolytic effect of kinsenoside in C3H10T1/2 adipocytes
To examine the catabolic effects of kinsenoside in adipocytes, the C3H10T1/2 cells were treated with an adipogenic induction medium for cell differentiation. The C3H10T1/2 adipocytes transformed by the adipogenic induction medium after 10-d incubation became oval shaped, and fat droplets appeared in red after oil-red O staining. Western blotting showed that PPAR[gamma], an adipocyte differentiation marker, was induced after 9-d incubation in the adipogenic induction medium (Fig. 1A). After cell differentiation, the amount of fat accumulation was 30 [+ or -] 6% assayed by oil-red O staining, and the ADP/ATP ratio was 0.22-0.32. To determine the optimal and effective kinsenoside concentration for lipolysis that had no cytotoxic effect, a broad concentration range of 10-1000 [micro]g/ml was examined using the LDH assay. Because no considerable cytotoxicity was observed for the kinsenoside concentrations not exceeding 50 [micro]g/ml (Fig. IB), the cells were treated with kinsenoside at 10-50 [micro]g/ml to examine the concentration-dependent effect on lipolysis. A kinsenoside concentration of less than 50 [micro]g/ml was used because the cells showed no alteration in the ratio of Bcl-xS to Bcl-xL and no appearance of the active form of caspase 3 compared with the control group (Fig. S4). By contrast, a 10-20-fold increased concentration of kinsenoside (500-1000 [micro]g/ml) increased the ratio of Bcl-xS to Bcl-xL, the activated caspase 3, and LDH release from adipocytes, indicating the occurrence of cell apoptosis at these high concentrations. However, whether this is involved in disrupting the redox system, leading to ATP depletion, remains to be determined. The effects of increased concentration of kinsenoside (50-1000 [micro]g/ml) in the cell apoptosis, induction of lipolysis-related proteins, and levels of mitochondrial-specific protein, voltage-dependent anion channel (VDAC) were shown in the Fig. S4 in comparison with undifferentiated and differentiated cells without kinsenoside treatment. Among them, the ratio of VDAC to GAPDH increased by 24%, suggesting the increased mitochondrial number after cell differentiation. Additionally, treating adipocytes with kinsenoside at 50-1000 [micro]g/ml concentration-dependently elevated the ratios of VDAC to GAPDH by approximately 24,32 and 43%, as compared to adipocytes without treatment (Fig. S4).
Fig. 2A shows a reciprocal relationship between the amount of fat deposited and glycerol secreted from the kinsenoside-treated cells, indicating the lipolytic effect of kinsenoside. The ratio of ADP/ATP was increased to 0.56-0.62 in adipocytes treated with kinsenoside at 50[micro]g/ml, an approximately 2.2-fold increase compared with the control. In addition, western blotting revealed that in the cells treated with kinsenoside at 10-50 [micro]g/ml for 1 h, kinsenoside concentration dependently induced AMPK and PPAR[alpha] phosphorylation and increased the ATGL protein level (Fig. 2B). In addition to the AMPK pathway, another axis involved in lipolysis is the PKC-ERK1/2 pathway. Therefore, the effect of kinsenoside in ERK1/2 phosphorylation was analyzed using western blot analysis. The results showed no apparent alteration in ERK1/2 phosphorylation and the protein level (Fig. 2B). Thereafter, a kinsenoside concentration of 50 ([micro]g/ml was used to examine the signaling pathway and gene regulation involved in lipolysis.
Lipolytic effect of kinsenoside in C3H10TI/2 adipocytes through an AMP-activated protein-kinase-dependent pathway
A panel of lipolysis-related proteins was analyzed using western blotting in the cells treated with kinsenoside at 50 [micro]g/ml (Fig. 3A). Kinsenoside elevated the phosphorylation of AMPK (a signaling molecule) and the transcriptional factors PPAR[alpha] and CREB. Furthermore, kinsenoside upregulated modifying proteins such as SIRT1 (a deacetylase) and PGC-1[alpha] (a PPAR[alpha] coactivator). These effects might upregulate ATGL (a lipolysis mediator) and CPT1 (a mitochondrial transporter of FAs). Moreover, kinsenoside phosphorylated and inactivated ACC, which might have inhibited the formation of malonyl-CoA, a CTP1 inhibitor. This observation suggested that the mitochondrial transport of FAs was increased, and conversely, FA synthesis was lowered in the kinsenoside-treated adipocytes.
The involvement of AMPK activation in kinsenoside-mediated lipolysis was examined using an AMPK agonist (5-aminoimidazole-4-carboxamide-l-[beta]-D-riboside; AICAR) and inhibitor (cpdC). Fig. 3B shows that AICAR, a cAMP agonist, mimicked the kinsenoside-mediated increase in glycerol secretion. However, additional treatment with cpdC reverted the effect. A similar pattern was observed using western blotting (Fig. 3C); AICAR mimicked the function of kinsenoside in activating the downstream targets of lipolysis, including the phosphorylation and upregulation of these lipolysis-related proteins, which were alleviated by cpdC treatment. Western blotting of the cells after nuclear-cytosolic fractionation demonstrated that similar to kinsenoside, AICAR increased the nuclear translocation of PPAR[alpha], CREB, S1RT1, and PGC-1[alpha] in adipocytes, and cpdC treatment prevented the effects.
Induction of CPT1 by the activation of PPAR[alpha], synergized by the upregulation of PGC-1[alpha] and SIRT1 in kinsenoside-treated adipocytes
To identify the mechanism underlying PGC-1[alpha] upregulation and increased PPAR[alpha] transactivation, a series of assays, including ChIP, EMSA, and Co-IP, were conducted (Fig. 4). The ChIP assay revealed an increased recruitment of CREB to the CRE-binding site in the PGC-1[alpha] gene promoter region (Fig. 4A). However, cpdC reverted the effect, indicating the importance of AMPK activation in this event. EMSA revealed that kinsenoside increased the DNA binding activities of CREB to the oligonucleotides derived from the putative CREB responsive element of the PGC-1[alpha] promoter region, which was abolished by cpdC treatment and the competition of the unlabeled oligonucleotides (Fig. 4B). The Co-IP assay revealed that the increase in PGC-1[alpha] acetylation caused by sirtinol (a SIRT1 inhibitor) considerably reduced the extent of the PGC-1[alpha]-PPAR[alpha] interaction in the kinsenoside-treated adipocytes (Fig. 4C), suggesting that epigenetic modification of PGCla is crucial for PPAR[alpha] transactivation.
The increased activity of PPAR[alpha] shown in Fig. 4 was validated by the expression level of CPT1, a downstream target of PPAR[alpha]. Notably, western blotting showed that kinsenoside caused CPT1 induction, which conversely was attenuated by PPAR[alpha] silence (Fig. 5A), suggesting that PPAR[alpha] is essential in kinsenoside-mediated CPT1 induction. In addition, in the glycerol assay, cells (with genetically or pharmacologically manipulated PPAR[alpha]) overexpressing PPAR[alpha] or treated with Wy14643 (a PPAR[alpha] agonist) mimicked the effect of kinsenoside in increasing glycerol secretion. However, treatment with GW6471 (a PPAR[alpha] antagonist) and siPPAR[alpha] eliminated kinsenoside-mediated lipolysis (Fig. 5B), suggesting that PPAR[alpha] is essential for kinsenoside-mediated lipolysis.
Kinsenoside has been shown to increase lipolysis and insulin sensitivity in high-fat- and aurothioglucose-induced obese murine animals (Du et al. 2001). However, the molecular mechanism underlying kinsenoside-mediated lipolysis remains unclear. In using C3H10T1/2 adipocytes in vitro to characterize the signaling pathway involved in the kinsenoside-mediated lipolysis and mitochondrial transporter of FAs, CPU, we demonstrated that AMPK is responsible for the roles of kinsenoside in lipolysis and CPT induction. A summarized scheme of the mechanisms underlying the lipolytic effect of kinsenoside is shown in Fig. 6. Our finding is in agreement with the finding of Gaidhu et al. (2009), who showed that AICAR-induced AMPK activation promotes energy dissipation in white adipocytes through ATGL upregulation. In addition, we showed that AMPK inactivated ACC through phosphorylation, which might also have contributed to the increased CPT1. Assifi et al. (2005) and Matejkova et al. (2004) concurred that AMPK inactivated ACC, resulting in a decreased concentration of malonyl-CoA, and subsequently alleviated CPU inhibition, which ultimately leads to inhibition of lipid synthesis and an increase in lipid oxidation.
The cAMP-dependent PKA axis is another well-recognized lipolysis pathway. This axis phosphorylates and inactivates AMPK[alpha] (by phosphorylating Seri 73) to promote efficient lipolysis by eliminating the toxic effect of PKA activation in the production of excessive FAs (Djouder et al. 2010). Incongruously, the results of our study demonstrate that AMPK phosphorylation at Thrl 72 is essential for kinsenoside-mediated lipolysis through the SIRT1/CREB/PGC1[alpha]/ PPAR[alpha]-dependent pathway. Moreover, we showed that CREB activation depends on AMPK activation because the AMPK inhibitor, cpdC, reduced CREB phosphorylation and nuclear translocation in kinsenoside-treated adipocytes (Fig. 3C). This finding differs from that of Delghandi et al. (2005), who showed that PKA increased the nuclear translocation and transactivation activity of CREB. These findings suggest that different signaling pathways initiate lipolysis in various cell types in response to varying chemical treatments. Despite the involvement of AMPK and PKA in lipolysis, a previous study showed that [alpha]1-adrenoceptors elevated lipolysis through an IP3-mediated increase in cytosolic calcium levels, resulting in the activation of [Ca.sup.2+]/calmodulin-dependent kinase II. This caused an increase in the membrane translocation of PKC and the resulting ERK1/2 activation, which phosphorylated HSL and consequently caused lipolysis (Jocken and Blaak 2008). The results of our study preclude the possibility that the PKC-ERK1/2 pathway is involved in kinsenoside-mediated adipocyte lipolysis by showing no apparent alteration in ERK1/2 phosphorylation (Fig. 2B).
In addition, the results of our study show that the loss of function of PPAR[alpha] by siPPAR[alpha] eliminated kinsenoside-mediated CPT1 induction, suggesting that CPT1 is a downstream target of PPAR[alpha]. The increased transactivation of PPAR[alpha] resulted from the activation and upregulation of PGC-1[alpha], a PPAR[alpha] coactivator, by AMPK-mediated CREB phosphorylation and the increased binding of CREB to the CRE of the PGC-1[alpha] promoter region in cells treated with kinsenoside. Furthermore, PGC-1[alpha] is deacetylated by the increased SIRT1 level and phosphorylated by AMPK activation, thereby increasing the PPAR[alpha]PGC-1 a interaction and PPAR[alpha] transactivation in kinsenoside-treated adipocytes. The mechanisms underlying the increased PPAR[alpha] transactivation by SIRT1 and PGC-1[alpha] are in agreement with the findings of Jager et al. (2007), showing that AMPK-mediated upregulation and subsequent phosphorylation ofPGC-1[alpha] initiates many of the vital gene regulatory functions of AMPK in skeletal muscles including glucose transporter 4 and mitochondrial genes. Therefore, in this study, the induction of CPT by kinsenoside through AMPK activation might have catalyzed the increased mitochondrial transport of FAs, ultimately contributing to the increased FAO.
The results of our study demonstrated that kinsenoside increased lipolysis through AMPK-mediated upregulation of lipolytic-related genes including ATGL, CPT1, SIRT1, pACC, pCREB, and pPPAR[alpha]. Understanding the molecular mechanisms underlying the lipolytic effect of kinsenoside provides an essential molecular basis for designing new therapeutic strategies to eliminate obesity-associated complications and might enable the development of kinsenoside as a dietary supplement for lipolysis in the future.
Received 11 August 2014
Revised 7 March 2015
Accepted 10 April 2015
Conflict of interest
The authors declare that no competing interests exist.
We thank Professor Yung-Hsi Kao (National Central University, Taiwan) for his excellent technical assistance and invaluable input into this work. This study was funded by a grant from the National Science Council, Taiwan (NSC102-2320-B-038-030-MY3(2-3)).
Supplementary material associated with this article can be found, in the online version, at doi:l 0.1016/j.phymed.2015.04.001.
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Kur-Ta Cheng (a,b), Yu-Shiou Wang (b,c), Hsiu-Chu Chou (b,d), Chih-Cheng Chang (b,c), Ching-Kuo Lee (e), Shu-Huijuan (b,c) *
(a) Department of Biochemistry and Molecular Cell Biology, School of Medicine, College of Medicine, Taipei Medical University, Taipei, Taiwan
(b) Graduate Institute of Medical Sciences, Taipei Medical University, Taipei, Taiwan
(c) Department of Physiology, School of Medicine, College of Medicine, Taipei Medical University, Taipei, Taiwan
(d) Department of Anatomy and Cell Biology, School of Medicine, College of Medicine, Taipei Medical University, Taipei, Taiwan eSchool of Pharmacy, Taipei Medical University, Taipei, Taiwan
Abbreviations: PPAR[alpha], peroxisome proliferator-activated receptor alpha; PGC-1[alpha], peroxisome proliferator-activated receptor gamma coactivator-1 alpha; CPT1, carnitine palmitoyltransferase 1; AMPK, AMP-activated protein kinase; AICAR, 5-aminoimidazole-4-carboxamide-1-[beta]-D-riboside; cpdC, compound C; HSL, hormonesensitive lipase; ATGL, adipose triglyceride lipase; ACC, acetyl-CoA carboxylase; FAO, fatty acid oxidation; CRE, cAMP response element; CREB, CRE-binding protein; SIRT1, silent information regulator T1; HDAC, histone deacetylase; ERK1/2, extracellular signal-regulated protein kinases 1 and 2; EMSA. electrophoretic mobility shift assay; Chip, chromatin immunoprecipitation; Co-IP, coimmunoprecipitation.
* Corresponding author. Tel.: +886 2 27361661; fax: +886 2 23461737.
E-mail address: email@example.com, firstname.lastname@example.org (S.-H. Juan).
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|Author:||Cheng, Kur-Ta; Wang, Yu-Shiou; Chou, Hsiu-Chu; Chang, Chih-Cheng; Lee, Ching-Kuo; Juan, Shu-Hui|
|Publication:||Phytomedicine: International Journal of Phytotherapy & Phytopharmacology|
|Date:||Jun 15, 2015|
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