β-Lapachone alleviates alcoholic fatty liver disease in rats

Cellular Signalling 26 (2014) 295–305

Contents lists available at ScienceDirect

Cellular Signalling journal homepage: www.elsevier.com/locate/cellsig

β-Lapachone alleviates alcoholic fatty liver disease in rats Sanghee Shin a,1, Jisoo Park a,1, Yuwen Li a,b, Ki Nam Min c, Gyeyeong Kong a, Gang Min Hur a, Jin Man Kim d, Minho Shong e, Min-Suk Jung c, Jong Kook Park c, Kyeong-Hoon Jeong c, Myoung Gyu Park c, Tae Hwan Kwak c, Derek P. Brazil f, Jongsun Park a,⁎ a Department of Pharmacology, Metabolic Diseases and Cell Signaling Laboratory, Research Institute for Medical Sciences, College of Medicine, Chungnam National University, Daejeon 301-474, South Korea b Department of Pharmacy, Xijing Hospital, Fourth Military Medical University, Shaanxi, China c Mazence Inc. R&D Center, Suwon 443-813, South Korea d Department of Pathology, College of Medicine, Chungnam National University, Daejeon 301-131, South Korea e Internal Medicine, College of Medicine, Chungnam National University, Daejeon 301-131, South Korea f Centre for Experimental Medicine School of Medicine, Dentistry and Biomedical Sciences, Queen's University Belfast, BT12 6BA Northern Ireland, UK

a r t i c l e

i n f o

Article history: Received 22 September 2013 Received in revised form 4 November 2013 Accepted 18 November 2013 Available online 22 November 2013 Keywords: β-Lapachone Fatty liver diseases AMPK PPARα Metabolic syndrome

a b s t r a c t Alcohol-induced liver injury is the most common liver disease in which fatty acid metabolism is altered. It is thought that altered NAD+/NADH redox potential by alcohol in the liver causes fatty liver by inhibiting fatty acid oxidation and the activity of tricarboxylic acid cycle reactions. β-Lapachone (βL), a naturally occurring quinone, has been shown to stimulate fatty acid oxidation in an obese mouse model by activating adenosine monophosphate-activated protein kinase (AMPK). In this report, we clearly show that βL reduced alcoholinduced hepatic steatosis and induced fatty acid oxidizing capacity in ethanol-fed rats. βL treatment markedly decreased hepatic lipids while serum levels of lipids and lipoproteins were increased in rats fed ethanol-containing liquid diets with βL administration. Furthermore, inhibition of lipolysis, enhancement of lipid mobilization to mitochondria and upregulation of mitochondrial β-oxidation activity in the soleus muscle were observed in ethanol/βL-treated animals compared to the ethanol-fed rats. In addition, the activity of alcohol dehydrogenase, but not aldehyde dehydrogenase, was significantly increased in rats fed βL diets. βL-mediated modulation of NAD+/NADH ratio led to the activation of AMPK signaling in these animals. Conclusion: Our results suggest that improvement of fatty liver by βL administration is mediated by the upregulation of apoB100 synthesis and lipid mobilization from the liver as well as the direct involvement of βL on NAD+/NADH ratio changes, resulting in the activation of AMPK signaling and PPARα-mediated β-oxidation. Therefore, βL-mediated alteration of NAD+/NADH redox potential may be of potential therapeutic benefit in the clinical setting. © 2013 Elsevier Inc. All rights reserved.

1. Introduction Alcoholic liver disease is a major cause of illness and death in most well-developed countries and is becoming a leading cause of disease in developing countries [1]. In the initial stages of alcoholic liver disease, fat accumulation in hepatocytes leads to the development of fatty liver, which is a reversible condition. With the continuation of alcohol consumption, fatty liver may progress to hepatitis and fibrosis, which may lead to liver cirrhosis [2]. It has been also suggested that patients with fatty liver are more susceptible to fibrotic liver diseases such as hepatitis and fibrosis [3]. Therefore, alcoholic fatty liver has long been considered benign; however, increasing evidence supports the idea that it is a

⁎ Corresponding author at: Department of Pharmacology, College of Medicine, Chungnam National University, Daejeon 301-474, South Korea. Tel.: +82 42 580 8252; fax: +82 42 585 6627. E-mail address: [email protected] (J. Park). 1 These authors contributed equally to this work. 0898-6568/$ – see front matter © 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.cellsig.2013.11.020

pathologic condition. Blunting of the accumulation of fat within the liver during alcohol consumption may block or delay the progression of fatty liver to hepatitis and fibrosis [4]. Therefore, recovery from a fatty liver would decrease the susceptibility to, and prevent the progression of liver fibrosis or cirrhosis. β-Lapachone (βL; also known as ARQ 501), currently in phase II clinical trials for the treatment of pancreatic cancer [5], is an o-naphthoquinone originally isolated from the bark of the lapacho tree (Tabebuia avellanedae) [6]. βL is a prodrug and becomes cytotoxic to cancer cells following bioreduction. The enzyme involved in the bioreduction of quinone-containing drugs is NAD(P)H:quinone oxidoreductase (NQO1) [7–9]. βL is reduced to an unstable hydroquinone that spontaneously reverts to its parent structure using two oxygen molecules [5]. As a result, reactive oxygen species are generated causing DNA damage, γ-H2AX foci formation, poly(ADP-ribose) polymerase-1 (PARP-1) hyperactivation and subsequent loss of ATP and NAD+ [10,11]. βL induced cell death is unique because PARP-1 and p53 proteolysis occurs concomitant with calpain activation [12]. βL mediated cell

296

S. Shin et al. / Cellular Signalling 26 (2014) 295–305

death exhibited classic features of apoptosis, but was not dependent on typical apoptotic mediators, such as p53, Bax/Bak, or caspases [5]. Interestingly, it has recently been proposed that βL plays an important role in regulating metabolic syndrome such as obesity [13] and atherosclerosis [14] through NQO1-mediated changes of NAD+/NADH redox potential. In the current study, we have investigated the mechanism of βL action on the improvement of alcohol-induced fatty liver in a rat model and primary hepatocytes. Our results suggested that βL prevents the onset of steatosis and plays a critical role in lipid metabolism and lipid mobilization by directly or indirectly modulating NAD+/NADH ratio in the injured liver of rats.

50 mM Tris–HCl (pH 7.4), 0.25 M sucrose and 1 mM EDTA, and centrifuged for 10 min at 12,000 ×g. Lipids in the liver homogenate were extracted using chloroform/methanol (1:2 vol/vol), evaporated, and dissolved in 2-propanol. The amounts of triglyceride, total cholesterol, and phospholipids were assayed enzymatically, using kits obtained from Wako Pure Chemicals Co. (Osaka, Japan) with a spectrophotometer DU-650 reader (Beckman, Fullerton, CA) at 600 nm. For histological analysis, liver tissues were fixed with formalin, dehydrated with ethanol, embedded in paraffin, cut at a thickness of 5 μM, and stained with hematoxylin and eosin. Alternatively, hepatic lipids were stained by an Oil red O method.

2. Materials and methods

2.5. Analysis of fatty acid oxidation

2.1. Chemical and reagents RPMI 1640 medium, fetal bovine serum (FBS), L-glutamine, penicillin, streptomycin and other cell culture reagents were obtained from Invitrogen (USA). Anti-AMPK, anti-pT179 (AMPK), anti-ACC and antipS79 (ACC) antibodies were obtained from Cell Signaling Technology (Danvers, MA). Anti-PPARα, anti-PPARγ and anti-RXR antibodies were purchased from Perseus Proteomics Inc. (Tokyo, Japan). Lipid extraction kit was purchased from Wako Pure Chemical (Osaka, Japan). Alcohol dehydrogenase (ADH) assay kit was obtained from Biomedical Research Service. Oil red O and 6-methoxy-2-napthaldehyde were obtained from Sigma (St. Louis, MA).

To determine fatty acid oxidation in the muscle, 14C-palmityl-CoAoxidation was measured in unfrozen soleus muscle using 14CO2 and 0.2 ml of benzethonium solution as described previously [15]. Briefly, soleus muscle was mixed with the assay buffer containing 1.25 M NaCl, 0.1 M MgSO4·7H2O, 1.2 mM KH2PO4, 0.5 M KCl, 0.2 M glucose, 1 M NaHCO 3, 0.25 M CaCl 2 and 1 mM [14C]-palmityl-CoA (1 mCi/mmol; Perkin Elmer; Chicago, IL), gassed for 30 s under humidified 95/5% O2/CO2, covered with filter paper moistened with 0.2 ml of benzethonium solution and then incubated for 1 h at 37 °C. The reaction was stopped with 6% TCA solution and the radioactivity trapped in the filter paper was determined. Fatty acid β-oxidation activity was expressed as nmol/min/liver.

2.2. Synthesis and formulation of compounds

2.6. Measurement of enzymatic activity; ADH and ALDH

β-Lapachone (βL) and other related 1,2-naphthoqunes were prepared by a two step synthetic process, resulting in a purity N 99.9% as described previously [6]. Briefly, the lithium salt of 2-hydroxy-1,2naphthoquinone was treated with several allyl-halides, such as 3-bromo-1-propene, 1-bromo-3-methyl-2-butene, 3-bromo-2-methyl-1-propene, and 1-bromo-2-butene, to give several 2-hydroxy-3allyl-1,4-naphthoquinone derivatives. Then each 2-hydroxy-3-allyl1,4-naphtho-quinone derivative was treated with H2SO4 and purified by recrystallization to give several pure 1,2-naphthoquinones, including βL.

For the measurement of alcohol dehydrogenase (ADH) activity, unfrozen liver was homogenized by homogenizer in 0.25 M sucrose at 4 °C and cleared by centrifugation at 600 ×g for 10 min. The resultant supernatant was mixed with the reaction buffer containing 50 mM sodium phosphate, 300 μM formazan, 11.4 mM NAD, and 12 mM 4-methylpyrazol. The reaction was stopped by the addition of 3% acetic acid. The relative ADH activity was measured at O.D. 492 nm and normalized with total amount of protein. In the case for aldehyde dehydrogenase (ALDH) activity, 100 μg of lysates was incubated with the reaction buffer containing 11.4 mM sodium phosphate (pH 8.5), 300 μM NAD, 4-methylpyrazol (ADH inhibitor) and 6 mM 6-methoxy2-naphthaldehyde as a substrate. The fluorescence of reaction was measured with a CytoFluor II fluorescence plate reader (excitation — 310 nm, emission — 360 nm; Applied Biosystems, Foster City, CA).

2.3. Rat model of alcoholic fatty liver All animal procedures were in accordance with the guidelines issued by the Institutional Animal Care and Use Committee of the Chungnam National University School of Medicine. Four-week-old male Sprague– Dawley (SD) rats obtained from Central Lab Animal (Seoul, Korea) and housed individually in temperature- and light-controlled rooms were randomly assigned to four groups: (a) rats fed isocaloric liquid diets without ethanol for 60 days (n = 4); (b) rats fed ethanol-containing liquid diets for 30 days and then fed daily with vehicle (SLS; Sodium Lauryl Sulfate) for the next 30 days (n = 6); (c) rats pair-fed ethanol containing liquid diets for 30 days and then fed daily with SLS for the next 30 days (n = 6) and (d) rats fed ethanol-containing liquid diets for 30 days and then fed daily with 150 mg or 250 mg of βL in SLS per kilogram of body weight for the next 30 days (n = 6). The calorie distribution of liquid diet components is as follows: 17% as protein, 36% as fat, 11% as carbohydrate and 36% as either ethanol or isocaloric maltose dextrin in the isocaloric liquid diets. Body weight and food intake were measured at every 5 days. At the end of the experiment, the rats were sacrificed and blood and liver tissue were collected. 2.4. Biochemical and histopathological analyses For measurement of hepatic lipid content, the liver was homogenized at 4 °C with a Polytron (Polytron PT-MR 2100, Kinematica, Switzerland) homogenizer in the homogenization buffer containing

2.7. Measurement of lipids in cultured primary hepatocytes from ethanol-fed rats Primary hepatocytes were prepared from ethanol-fed SD rats using the collagenase perfusion method as previously described [16]. Briefly, the isolated hepatocytes were placed on a culture plate coated with type I collagen at a density of 5 × 105 cells/ml and culture for 4 h in Williams' medium E supplement with 10% FBS, 1 nM dexamethasone. After serum-starvation, the cells were treated with either DMSO or 10 μM βL for 8 days, washed two times with cold PBS and then solubilized in lysis buffer containing 50 mM Tris–HCl (pH 7.4) and 0.5% SDS. Triglyceride in the cell lysates was extracted and measured. For histological detection of lipids, the cells were fixed with 10% formaldehyde for 1 h and subjected to lipid staining using Oil red O staining. 2.8. Electrophoretic mobility shift assays Binding and electrophoresis were performed as previously described [17]. Briefly, cells were placed on ice, harvested and extracted with nuclear lysis buffer containing 10 mM HEPES (pH 7.9), 10 mM KCl, 1.5 mM MgCl2, 0.05% NP-40 0.5 mM DTT and 0.5 mM PMSF by passing 20 times with a 1 ml Dounce Homogenizer by passing.

S. Shin et al. / Cellular Signalling 26 (2014) 295–305

Homogenates were centrifuged at 3000 ×g for 10 min, pellet was solubilized in 9 volumes of dialysis buffer containing 5 mM HEPES (pH 7.9), 26% glycerol, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM DTT, 0.5 mM PMSF and 300 mM NaCl, and used as a nuclear extract. The following oligonucleotides were used for the probes: PPRE motif (5′-GGG CAT TCT AGG TCA AAG GTC ATC CCC TCA-3′, 5′-GGG TGA GGG GAT GAC CTT TGA CCT AGA ATG-3′) and PPRE mutant motif (5′-GGG CAT TCT AGA ACA AAG AAC ATC CCC TCA-3′, 5′-GGG TGA GGG GAT GTT CTT TGT TCT AGA ATG-3′). Nuclear extracts were preincubated for 15 min at room temperature in the buffer containing 12 mM HEPES (pH 7.5), 5 mM MgCl2, 0.2 mM EDTA, 12% glycerol, 60 mM KCl, 0.2 mM DTT and 2 μg poly(dI-dC), followed by the incubation of mixture with the labeled probe for 15 min, the complexes were separated on a 5% native polyacrylamide gel. Gels were dried and visualized by autoradiography. 2.9. Immunoblot analysis Cells were collected and lysed in the lysis buffer containing 50 mM Tris–HCl (pH 7.6), 1% Nonidet P-40, 120 mM NaCl, 25 mM sodium fluoride, 40 mM beta-glycerol phosphate, 0.1 mM sodium orthovanadate, 1 mM phenylmethylsulfonyl fluoride, 1 mM benzamidine and 2 μM microcystin-LR. Fifty micrograms of the cell lysates was subjected to sodium dodecyl sulfate (SDS) polyacrylamide gel and blotted onto PVDF membrane. After blocking with 5% skim milk in PBST, the membrane was probed with the relevant antibody and visualized by enhanced chemiluminescence, according to the manufacturer's instruction (Amersham, Piscataway, NJ).

297

3. Results 3.1. βL prevents the accumulation of hepatic lipids in ethanol-treated rats βL has been shown to improve key symptoms of metabolic syndrome in rodent models [13,14]. In order to investigate the direct effects of βL on hepatic steatosis, we experimentally developed an animal model of ethanol-induced fatty liver. SD-rats were fed ethanolcontaining diets for 30 days, then further fed daily either with vehicle Sodium Lauryl Sulfate (SLS) or βL (150 mg/kg). Histological analysis of the liver reveals that lipid droplets had accumulated in hepatocytes preferentially in perivenular and periportal areas, while lipid droplets were rare in control animal (Fig. 1A, first two panels), validating this model. Interestingly, treatment of rats with βL for 30 days led to a decrease in the formation of lipid droplets, similar to levels of the control group livers (Fig. 1A, third panel). This observation was further confirmed by Oil-red O staining (Fig. 1B). Mean body weights of rats given ethanol/SLS diets are lower than control. Furthermore, mean body weights of ethanol/βL group were even lower than ethanol/ SLS group. This observation was also confirmed with the pair-fed ethanol/SLS group (Fig. 1C and D), suggesting that βL administration contributed to weight loss of these mice. Furthermore, all these metabolic changes including decreased food-intake (Fig. 1C), reduced body weight (Fig. 1D), decreased subcutaneous fat (Fig. 1E) and reduced lipid accumulation in the liver (Fig. 1F) were similarly observed in ethanol/βL group, indicating that βL plays an important role in the prevention of ethanol-induced fat accumulation in the liver.

2.10. Determination of NADH levels in H4IIE cells NADH level in H4IIE cells was measured by using Cell Counting Kit 8 (Dojindo, Kumamoto, Japan), which is based on the color development of reduced compound by NADH. After serum-starvation, the cells were treated with either 5 μM or 10 μM βL for the indicated time. Cells were incubated with 10% Dojindo's highly water-soluble tetrazolium salt for 2 h. Dojindo's highly water-soluble tetrazolium is reduced by dehydrogenases in cells to give a yellow-colored product (formazan) and the observance of reaction was measured with a Sunrise Observance plate reader at 450 nm (Tecan, Switzerland). Relative NADH levels were calculated by comparing the values with control sample (0 min) and expressed as a %. 2.11. RT-PCR analysis for the components of lipid metabolism Unfrozen tissues were homogenized by homogenizer in Easy Blue solution (Invitrogen, Carlsbad, CA). Homogenates were mixed in 20% chloroform and 30% isopropanol and then centrifuged at 10,000 ×g for 10 min at 4 °C. The pellet was washed with 70% ethanol and dissolved in DEPC-distilled water. Reverse-transcription was performed using 100 μg of RNA with 5 units of the MMLV reverse transcriptase, 1 × MMLV RT buffer (50 mM Tris–HCl (pH 8.3), 75 mM KCl and 10 mM MgCl2), 10 mM dithiothreitol, 0.25 mM dNTP mix, and 0.5 μg oligo(dT)12–18, and then was incubated at 37 °C for 60 min. The PCR protocol consisted of 27 cycles of amplification. GAPDH was used as an internal control in all reaction. The PCR products were analyzed by electrophoresis on 1% agarose gel. The detail information of each primer set is shown in Supplementary Table 1. The photographs were scanned with a Gel Doc 200 (Bio-Rad, Hercules, CA). 2.12. Statistical analysis Data are expressed as the mean ± standard deviation (S.D.) from at least three separate experiments performed in duplicate. The differences between groups were analyzed using a Student's t test and p b 0.05 was considered statistically significant. Statistical analyses were carried out using SPSS software (ver. 11.0; SPSS Inc., Chicago, IL).

3.2. Decreased hepatic lipid levels and increased circulating fatty acid levels after βL administration In rats given ethanol/SLS liquid diets, overall hepatic lipid (triglyceride, cholesterol and phospholipid) levels were slightly increased compared to controls, whereas there was a significant decrease in hepatic lipid levels in rats fed ethanol/βL liquid diets (Fig. 2A). In the case for serum lipid levels, the oppose effect on overall lipid content in serum was observed in βL-administrated rats (Fig. 2B). Since hepatic lipid levels are highly dependent on the mobilization of lipids from the liver to extra-hepatic tissues [18], the mRNA levels of ApoB100, which is an essential apolipoprotein of low-density lipoproteins, were measured. As shown in Fig. 2C, mRNA levels of ApoB100 in the liver were significantly elevated. In order to investigate the effect of βL on hepatic lipid synthesis, the components of fatty acid and cholesterol synthesis (ACC, FAS, SCD, ChREBP, LDL-r and HMG-CoA) were monitored. Overall the key component of lipogenesis was downregulated in EtOH/βL vs EtOH fed rats (Fig. 2D), suggesting that βL has an influence on lipid mobilization by promoting lipid export from, and inhibiting lipogenesis in, the liver. 3.3. Enhanced mitochondrial β-oxidation in βL treated groups Based on the previous observation that βL administration led to a decrease in alcohol-induced hepatic lipid accumulation (Fig. 1), the effect of βL on the lipid utilization of other tissues was determined. RT-PCR analysis of the liver and muscle revealed that the expression levels of carnitine palmitoyltransferase 1 (CPT1; CPT1A for liver, CPT1B for muscle) and CPT2 were markedly enhanced in βLadministrated group compared to controls (Fig. 3A). No significant changes were observed in fatty acid break-down enzymes (long-chain acyl-CoA synthetases and long-chain acyl-CoA dehydrogenase), indicating that βL-mediated improvement of fatty liver may be mediated by the efficient transportation of lipid to mitochondria rather than the enhancement of lipid break-down enzyme activity (Fig. 3A). To further confirm this observation, lipoprotein lipase (LPL) and mitochondrial β-oxidation activities were monitored. Interestingly, mRNA levels

298

S. Shin et al. / Cellular Signalling 26 (2014) 295–305

A

EtOH + βL

EtOH + SLS

60

Numbers of droplets

Control

50

*

40 30 20 10 0 Con EtOH EtOH + SLS + βL

**

B

EtOH + βL

EtOH + SLS

50

Relative density of Oil-Red O staining

Control

**

40 30 20 10 0

Body weight (g)

D

Body Weight 600

400

200

**

Food Intake 150

Control EtOH + SLS EtOH + SLS (Pair Fed) EtOH + βL

** ** ** ** ** ** ** ** Treatment of βL

Food intake (g)

C

0

Control EtOH + SLS EtOH + SLS (Pair Fed) EtOH + βL

100

50

**

*

30

40

** **

Treatment of βL

0 0

10

20

30

40

50

0

10

Days

E

Con EtOH EtOH + SLS + βL

EtOH +SLS

EtOH + βL 150mg 250mg

20

50

Days

F EtOH +SLS

EtOH + βL 150mg 250mg

Fig. 1. βL-induced metabolic changes in rats with alcoholic fatty liver. Histological sections of the liver from male rats fed with isocaloric liquid diets without ethanol for 60 days (n = 4; control), ethanol-containing liquid diets for 30 days and then fed daily with vehicle for the next 30 days (n = 6; EtOH + SLS), or ethanol-containing liquid diets for 30 days and then fed daily with 150 mg/kg βL for the next 30 days (n = 6; EtOH + βL) were stained (A) with hematoxylin and eosin. (B) Alternatively, hepatic lipids were stained with Oil red O. Arrows indicated the lipid droplet. Semi-quantification of lipid droplet and Oil-red O staining was performed. *, p b 0.05. **, p b 0.01. Scale bars represent 50 μm. Body weight (C) and food intake (D) were also monitored with an additional pair-fed group (n = 6; EtOH + SLS (pair fed)). (E) The photographic images of EtOH + SLS and EtOH + βL (150 mg/kg and 250 mg/kg)treated rats after 60 days of treatment. (F) Representative photographic images of the liver from the above groups. Scale bars represent 5 cm.

of LPL in aorta were also elevated (Fig. 3B) and mitochondrial β-oxidation activity in the soleus muscle was significantly enhanced in rats fed βL diets (Fig. 3C), confirming that βL promoted lipolysis by enhancing the mobilization and utilization of lipid in the peripheral tissues.

3.4. βL-mediated enhancement of lipid secretion in primary hepatocytes To test the possibility that the recovery from fatty liver and the stimulation of lipid mobilization from the liver by βL were exerted through direct effects on hepatocytes, primary hepatocytes from rats fed

S. Shin et al. / Cellular Signalling 26 (2014) 295–305

A 1.0

Triglyceride cholesterol phospholipid

8

0.8

6

0.6

**

4

**

2

0.4 0.2 0

0 Control

EtOH + SLS

EtOH + βL

C

D EtOH + SLS

Serum triglyceride, total cholesterol or phospholipids (mg/dl)

10

Total cholesterol or phospholipids (μg/mg liver protein)

B Hepatic Lipid

Triglyceride (μg/mg liver protein)

299

Serum Lipid 25 Triglyceride cholesterol phospholipid

20

** **

15

*

10 5 0 EtOH + SLS

Control

EtOH EtOH + SLS + βL

EtOH + βL

EtOH + βL

EtOH EtOH + SLS + βL ACC

ChREBP

FAS

LDL-r

SCD

HMG-CoA

ApoB100

3

*

2

1

0 EtOH + SLS

EtOH + βL

GAPDH Relative mRNA abundance

Relative mRNA abundance

GAPDH

1.5

FAS

ACC

SCD

ChREBP

1.0

*

* 0.5

LDL-r

HMG-CoA

*

**

**

** 0.0

SLS

βL SLS

βL SLS

βL SLS βL

SLS βL

SLS βL

Fig. 2. Changes in hepatic and serum lipid contents in rats fed isocaloric diets ethanol/SLS diets or ethanol/βL diets. Rats were fed with isocaloric liquid diets without ethanol for 60 days (n = 4; control), ethanol-containing liquid diets for 30 days and then fed daily with vehicle for the next 30 days (n = 6; EtOH + SLS), or ethanol-containing liquid diets for 30 days and then fed daily with 150 mg/kg βL for the next 30 days (n = 6; EtOH + βL). Triglyceride, cholesterol and phospholipid from (A) liver or (B) serum were measured as described in the Materials and methods. mRNA levels of (C) hepatic ApoB100 and (D) lipid synthesis-associated genes were analyzed by RT-PCR. The results are mean ± S.D. of three independent experiments. *, p b 0.05. **, p b 0.01. (ACC: acetyl-CoA carboxylase, FAS: fatty acid synthase, SCD: stearoyl-coenzyme A desaturase, chREBP: carbohydrate responsive-element binding protein, LDL-r: LDL receptor, HMG-CoA: 3-hydroxy-3-methyl-glutaryl-CoA).

ethanol-diets were prepared. Specific staining with Oil-red O clearly indicated that lipid droplets were retained within hepatocytes during the 8-day culture (Fig. 4A upper panel). However, treatment with 10 μM βL led to a remarkable decrease in lipid droplets in the primary hepatocytes (Fig. 4A lower). Consistent with this result, triglyceride content in the βL-treated hepatocytes was significantly reduced compared to controls (Fig. 4B), suggesting that βL directly decreased lipid components that had accumulated within hepatocytes as a result of longterm feeding of ethanol-diets. 3.5. ADH activity, but not ALDH activity, was enhanced in βL-treated animals Alcohol dehydrogenase (ADH) enzymes occur in many organisms and facilitate the inter-conversion between alcohols and aldehydes or ketones with the reduction of NAD+ to NADH [19]. Recent studies on metabolic syndrome demonstrated that βL regulated the NAD+/NADH ratio, working as a NADH:quinone oxidoreductase 1 (NQO1) substrate [13]. To evaluate the possibility that βL might directly affect ADH or

aldehyde dehydrogenase (ALDH) activity through the changes in the NAD+/NADH ratio, ADH activity from hepatic cell lines was measured. Among cell lines tested, H4IIE cells, which are rat hepatocytes, showed a higher ADH activity (Supplementary Fig. 1A). Therefore H4IIE cells were treated with 10 μM βL for the indicated time. As shown in Supplementary Fig. 1B, there was no apparent effect of βL on ADH activity in H4IIE cells. Contrasting this observation, there was a significant elevation of ADH activity in rats fed ethanol/βL diets (Fig. 5A), while no change in ALDH activity was observed in either group (Fig. 5B). In addition, mRNA and protein levels of ADH and ALDH appeared to be similar in both groups (Supplementary Fig. 1C), suggesting that enhancement of ADH activity in rats fed ethanol/βL diets is specific to the in vivo situation. 3.6. βL-induced NADH oxidation leads to activation of AMPK signaling Based on the previous observation that βL activated ADH activity in vivo (Fig. 5A), the involvement of βL on regulating the NAD+/NADH ratio was investigated in H4IIE cells. The measurement

300

S. Shin et al. / Cellular Signalling 26 (2014) 295–305

A Liver

Muscle

EtOH EtOH + SLS + βL 1

0.9

1

1.7

Soleus

EDL

Heart

EtOH EtOH + SLS + βL

EtOH EtOH + SLS + βL

EtOH EtOH + SLS + βL

1

1.1

1

1.0

1

1.0

1

1.2

1

1.2

1

2.5

1

1.4

1

1.1

1

3.4

VLACS

VLACS

CPT1B

CPT1A 1.2

1

CPT2

CPT2 1

1.0

1

0.9

1

0.8

1

1.1

LCAD

LCAD 1

1.0

1

1.0

1

1.0

1

1.0

GAPDH

GAPDH

B

C Aorta

LPL Relative mRNA abundance

GAPDH 5 4

*

3 2 1 0 EtOH EtOH + SLS + βL

[14C]-palmityl-CoA oxidation (cpm)

EtOH EtOH + SLS + βL 120000

*

100000 80000 60000 40000 20000 0 EtOH + SLS

EtOH + βL

Fig. 3. βL upregulates the expression of numerous genes involved in lipid oxidation. Rats were fed with ethanol-containing liquid diets for 30 days prior to feed daily with vehicle for the next 30 days (n = 6; EtOH + SLS) or with 150 mg/kg βL for the next 30 days (n = 6; EtOH + βL). (A) mRNA levels of lipid oxidation-associated genes from the liver or muscle were analyzed by RT-PCR. The quantifications of PCR products were shown as a fold-induction, compared to control. (VLACS: long-chain acyl-CoA synthetases, CPT1A: carnitine palmitoyltransferase 1A, CPT2: carnitine palmitoyltransferase 2, LCAD: long-chain acyl-CoA dehydrogenase). (B) Aortas were isolated freshly and mRNA of lipoprotein lipase from the aorta was analyzed by RT-PCR. (C) 14C-palmityl-CoA oxidation in the soleus muscle was measured as described in the Materials and methods. The results are mean ± S.D. of three independent experiments. *, p b 0.05.

of NADH levels revealed that the treatment of cells with 50 mM alcohol dramatically induced NADH levels while ethanol-mediated induction of NADH levels was inhibited by βL treatment (Fig. 6A). Furthermore, NADH levels in βL-treated H4IIE cells were reduced in a dosedependent manner (Fig. 6B), indicating that βL directly regulates the NAD +/NADH ratio (possibly via NQO1). The intracellular NAD+/ NADH ratio reflects the energy balance of cells [20,21]. The activation of AMP-activated protein kinase (AMPK) signaling, which acts as an energy sensor, was analyzed in βL-treated H4IIE cells and our rat model. Consistent with previous reports [13,14], βL promoted the activation of AMPK signaling (AMPK and acetyl-CoA carboxylase (ACC) phosphorylation) more than other AMPK activators (metformin and AICAR) in H4IIE cells (Fig. 6C and E). Similar results were also observed in βL-treated rats compared to controls (Fig. 6D), suggesting that βL activates AMPK signaling in vitro and in vivo.

3.7. Enhancement of βL-mediated lipid oxidation is mediated by PPARα Since peroxisome proliferator-activated receptor α (PPARα) mainly regulates the transcription of genes encoding fatty acid oxidation enzymes [18], the possible regulation of PPARα by βL was monitored in our rat model. Western blot analysis of the liver tissues indicated that protein levels of PPARα were significantly upregulated in the EtOH/βL-treated group with no apparent change in PPARγ detected (Fig. 7A). To further investigate the effects of βL on the DNA-binding activity of PPARα, electrophoretic mobility-shift assay was employed. As shown in Fig. 7B, treatment of H4IIE cells with 10 μM βL enhanced the DNA-binding activity of PPARα toward PPAR response elements of the acyl-CoA oxidase gene [22]. To evaluate βL-mediated PPARα activation in the presence of ethanol, H4IIE cells were treated with 50 mM ethanol for 5 h, followed by βL treatment. DNA-binding activity of PPARα was

S. Shin et al. / Cellular Signalling 26 (2014) 295–305

301

B

A

Triglyceride in cells (ug/mg cellular protein)

500

Control

βL

400

*

300 200 100 0 Control

βL

Fig. 4. In vitro effects of βL on lipid mobilization in primary cultured hepatocytes from rats with ethanol-induced fatty liver. Primary hepatocytes were prepared from rat fed ethanolcontaining liquid diets for 30 days and further cultured in either absence (control) or presence (βL) of 10 μM βL for 8 days. (A) Lipid droplets in primary hepatocytes were stained with Oil red O. Scale bars represent 20 μm. (B) Levels of triglyceride in the cells were measured as described in the Materials and methods. The results are mean ± S.D. of three independent experiments. *, p b 0.05.

blocked by ethanol and this ethanol-mediated inhibition of DNAbinding activity was completely recovered in the presence of βL (Fig. 7C), indicating that βL plays an important role in regulating PPARα transcription activity via modulation of DNA-binding activity in the presence of ethanol. 4. Discussion Alcoholic liver disease is the most common liver disease in which fatty acid metabolism is altered. It has been shown that alcohol metabolism alters the cellular NADH concentration which impairs β-oxidation and tricarboxylic acid cycle activity [23], resulting in elevated free fatty acid, increased formation of triacylglycerol, and increased rates of very low density lipoprotein synthesis and secretion [24,25]. The fatty liver persists despite attenuation of the altered redox state after chronic ethanol administration [26]. Here, we provide the first evidence that

B

EtOH EtOH + SLS + βL

ALDH

GAPDH

GAPDH

250

*

200 150 100 50 0 EtOH + SLS

EtOH EtOH + SLS + βL

ADH

EtOH + βL

Relative ALDH activity

Relative ADH activity

A

120 100 80 60 40 20 0 EtOH + SLS

EtOH + βL

Fig. 5. Modulation of ADH activity by βL administration in vivo. Rats were fed with ethanol-containing liquid diets for 30 days prior to feed daily with vehicle for the next 30 days (n = 6; EtOH + SLS) or with 150 mg/kg βL for the next 30 days (n = 6; EtOH + βL). (A) ADH activity and (B) ALDH activity in the liver were measured as described in the Materials and methods. mRNA level of each sample was also analyzed by RT-PCR (top panel). The results are mean ± S.D. of three independent experiments. *, p b 0.05.

β-Lapachone (βL) administration reduced alcohol-induced hepatic steatosis in a rat model for alcoholic fatty liver. Taken together, these results clearly suggest that improvement of fatty liver by βL administration is mediated by the upregulation of lipoproteins and lipid mobilization from the liver as well as the direct involvement of βL on NAD+/ NADH ratio changes, resulting in the activation of AMPK signaling and the increase in PPARα-mediated β-oxidation, as illustrated in Fig. 8. It has been proposed that a defect in secretion of lipid components such as very low density lipoprotein (VLDL) complex in hepatocytes is one of the major causes of alcoholic liver disease [27,28]. Davis and colleagues showed that the availability of apoB determines the capacity of the hepatocyte to assemble/secrete triacylglycerol-rich VLDL [29]. Consistent with these observations, reduction of hepatic lipid and induction of serum lipid by βL-administration (Fig. 2A and B) appear to be due to the induction of apolipoprotein B100 (Fig. 2C). This observation is further supported by Oil Red O staining of primary hepatocytes from alcohol-fed rats (Fig. 4A), and secreted triglycerides in the medium following βL treatment (Fig. 4B). In addition, increased fatty acid synthesis and decreased fatty acid degradation may be additional factors leading to alcoholic fatty liver [30]. Therefore, another possible explanation for the improvement of alcohol-mediated fatty liver by βL includes the inhibition of lipid synthesis (Fig. 2D) and the enhancement of lipid oxidation in peripheral tissues (Fig. 3). The effects of ethanol on organ function (brain, heart, and liver) are dependent upon the systemic concentrations of ethanol over time. Therefore, pharmacokinetics play a pivotal role in the pharmacodynamic actions of ethanol and of its metabolic product acetaldehyde [31]. Alcohol elimination rate varies as much as 3-fold from person to person [32]. Ethanol metabolic rate is influenced by the genetic variations in the principal alcohol metabolizing enzymes, cytosolic alcohol dehydrogenase (ADH), and mitochondrial aldehyde dehydrogenase (ALDH2). Both ADH and ALDH use NAD+ as a cofactor in the oxidation of ethanol and acetaldehyde. The rate of metabolism as well as the NAD+/NADH ratio in the liver varies with the fed or fasted state [33]. Furthermore, the rate of alcohol metabolism is determined not only by the amount of ADH and ALDH2 enzyme in tissue but also by the concentrations of the cofactors, NAD+ and NADH, and of ethanol and acetaldehyde in the cellular compartments (cytosol and mitochondria of hepatocytes). Interestingly, βL administration of ethanol-fed rats enhanced ADH activity, but not ALDH activity, in the liver (Fig. 5) while stimulation of H4IIE cells with βL did not affect ADH activity (Supplementary Fig. 1B), possibly due to the lower NQO1 activity in the cell culture system. However, there was no significant difference in the rate of alcohol

302

S. Shin et al. / Cellular Signalling 26 (2014) 295–305

susceptibility to modulate the higher NAD+/NADH ratio. Indeed, ethanol-induced upregulation of NADH levels was blocked by βLmediated NADH oxidation (Fig. 6A), which appears to enhance ADH activity (Fig. 5A) as well as AMPK signaling (Fig. 6C and E). As a result, the enhancement of AMPK signaling leads to inhibition of fatty acid synthesis. It is noteworthy that the reduced cytosolic NADH levels induced by

metabolism between control and βL-fed group when rats were acutely administrated with ethanol (Supplementary Fig. 2). NQO1 is a flavoenzyme that uses NAD(P)H as an electron donor for the bioreduction of quinine-containing substrates. Accumulating evidence suggests that βL is a specific and high-affinity substrate of NQO1 in vitro and in vivo [13,34]. Therefore βL might have the

A

B

200

150

** **

100

50

NADH (%)

NADH (%)

150

**

Control βL (10uM) EtOH + SLS EtOH+ βL

0 0

30

60

90

50

* **

**

**

**

**

**

Control βL (5uM) βL (10uM)

30

60

90

0

120 min

C

100

0

120 min

D Rat model

βL

Con

EtOH + βL

Con

H4llE cells

EtOH + SLS α pAMPK (T172)

Met. AICAR α pAMPK (T172)

α AMPK

α AMPK

α pACC (S79)

pAMPK/AMPK

α ACC

*

2.0

pAMPK/AMPK

*

1.5 1.0 0.5 0.0

βL

Con

Met. AICAR

Relative Intensity

Relative Intensity

2.5

2.5

*

2.0 1.5 1.0 0.5 0.0

Con

βL SLS

pACC/ACC 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0

*

Con βL SLS

E H4llE cells βL 15

30

60

120

(min) α pAMPK (T172) α AMPK α pACC (S79) α ACC α Actin

pAMPK/AMPK

pACC/ACC 2.0

2.0

Relative Intensity

0

* *

* 1.5

1.5

*

*

1.0

1.0

0.5

0.5

0.0

0.0 0 15 30 60 120 (min)

βL

0 15 30 60 120

βL

S. Shin et al. / Cellular Signalling 26 (2014) 295–305

303

A 1#

2#

EtOH + βL

EtOH + SLS 1#

2#

1#

α PPARα α PPARγ α p84

2.5

*

2# 2.0

**

1.5

1.0

1.0

0.5

0.5 0.0 Con

B

C 30

60

120

Con EtOH EtOH +SLS +βL

EtOH

PPARα

PPARα

8

3.5

** 6

*

4 2

0

30

60

120

βL (min)

Relative Intensity

Relative Intensity

EtOH EtOH +SLS +βL

βL

βL (min) 0

2.0

1.5

0.0

0

PPAR /p84

PPARα/p84 2.5

Relative Intensity

Con

3.0

*

2.5 2.0 1.5 1.0 0.5 0.0 βL

EtOH Fig. 7. βL enhanced PPARα expression and transcriptional activity (A) Rats were fed with ethanol-containing liquid diets for 30 days (n = 2; control) prior to feed daily with vehicle for the next 30 days (n = 2; EtOH + SLS) or with 150 mg/kg βL for the next 30 days (n = 2; EtOH + βL). Liver lysates were resolved by SDS-PAGE and analyzed by immunoblotting using antiPPARα antibodies and anti-PPARγ antibodies (top panel). Control for equal loading was checked by immunoblot analysis with anti-p84 antibodies. Relative density was also obtained by densitometry of the corresponding immunoblot data (bottom panel). Statistical differences of protein amount were determined by comparing the values for p84 control protein at each lane. *, p b 0.05. **, p b 0.01. (B) H4IIE cells were treated with 10 μM βL for the indicated times. Nuclear extracts were prepared for electrophoretic mobility shift assays. (C) H4IIE cells were treated either without or with 50 mM ethanol prior to stimulation of 10 μM βL for 60 min. Nuclear extracts were analyzed with radio-labeled probe for PPARα by using electrophoretic mobility shift assay. Arrows indicate PPARα (top panel). Relative density was obtained by densitometry of the corresponding signal of electrophoretic mobility shift assay data (bottom panel). The results represent the mean ± S.D. of three independent experiments. *, p b 0.05. **, p b 0.01.

βL administration appear to have an influence on the proper distribution of NADH reducing equivalents in mitochondria for energy production. This may indicate that βL might transiently increase AMP levels and activate AMPK signaling to compensate for cellular energy depletion [13,35,36]. Consistent with this hypothesis, the transport of lipid to mitochondria (Fig. 3A) and lipid oxidation (Fig. 3C) was significantly enhanced in the peripheral tissues of βL-administrated animals.

PPARα coordinates fatty acid metabolism in the liver when it is dimerized with retinoid X receptors (RXR) by controlling a set of genes containing peroxisome proliferator response elements (PPREs), which are involved in free fatty acid transport and oxidation [37]. These include membrane transporters such as carnitine palmitoyltransferase I, apolipoprotein genes, and several components of the mitochondrial and peroxisomal fatty acid oxidation pathways [38].

Fig. 6. Effects of βL on NAD+/NADH ratio and AMPK signaling (A) H4IIE cells were treated with either vehicle, 10 μM βL, ethanol + SLS or ethanol + βL for the indicated time. The changes of NADH in each condition were measured as described in the Materials and methods. (B) Dose dependency of βL effects on NADH levels was measured in H4IIE cells. The results represent the mean ± S.D. of three independent experiments. *, p b 0.05. **, p b 0.01. (C) H4IIE cells were treated with 10 μM βL, 1 mM metformin, or 1 mM AICAR for 30 min. Cell lysates were resolved by SDS-PAGE and analyzed by immunoblotting using anti-phospho-specific AMPK antibodies and anti-AMPK antibodies (top panel). Relative density was obtained by densitometry of the corresponding immunoblot data (bottom panel). Statistical differences of phospho-AMPK were determined by comparing the values for total AMPK at each lane. *, p b 0.05. **, p b 0.01. (D) Rats were fed with ethanol-containing liquid diets for 30 days (n = 4; control) prior to feed daily with vehicle for the next 30 days (n = 6; EtOH + SLS) or with 150 mg/kg βL for the next 30 days (n = 6; EtOH + βL). Liver lysates were analyzed with the corresponding antibodies. (E) H4IIE cells were treated with 10 μM βL for the indicated time. Cell lysates were analyzed by immunoblotting using the indicated antibodies. Control for equal loading was checked by immunoblot analysis with anti-actin antibodies. Results are representative of three independent experiments (top panel). Relative density was also obtained by densitometry of the corresponding immunoblot data (bottom panel). Statistical differences of phospho-protein were determined by comparing the values for total protein at each lane. *, p b 0.05. **, p b 0.01.

304

S. Shin et al. / Cellular Signalling 26 (2014) 295–305

Ethanol Changes in NAD+/NADH ratio NAD+

ADH

β-Lapachone-H2

Oxidoreductases NADH

NAD+/NADH

ATP/AMP

β-Lapachone

P

NAD+

ALDH

β-Lapachone-H2

PPARα

Oxidoreductases NADH

Sirt-1

AMPK

Acetaldehyde

P

SREBP

β-Lapachone

CPT1 ACC Acetate

Acetyl-CoA

Malonyl-CoA β-oxidation

FAS FA synthesis

Improvement of Alcoholic fatty liver Fig. 8. Proposed model for the improvement of alcoholic fatty liver by βL. Ethanol metabolism generates the reduced form of NADH, which promotes steatosis by stimulating the synthesis of fatty acids and opposing their oxidation. βL altered NAD+/NADH redox potential in the liver, leading to activation of ADH and the reduction of ATP/AMP ratio. Subsequently, AMPK gets activated and inhibits ACC activity. It has been shown that AMPK can also block the SREBP function by phosphorylation events, leading to reduction of FAS activity. Furthermore βL-induced changes of NAD+/NADH ratio can also activate Sirt1, which is a NAD+ dependent deacetylase, resulting in the upregulation of PPARα. Thereby the enhancement of mitochondrial β-oxidation and inhibition of lipogenesis will improve the alcohol-induced fatty liver in patho-physiological condition. Therefore, the βL-mediated alteration of NAD+/NADH redox potential can be the new therapeutic benefit in clinical settings.

The transcriptional activity of PPARα is activated on binding of free fatty acid as well as a number of drugs such as nonsteroidal antiinflammatory drugs and fibrates [39]. This provides a feedback mechanism that increases the capacity of the liver to decrease fatty acids when the intracellular concentration of the fatty acids increases. Fasted PPARα knock-out mice have severe defect in their ability to oxidize free fatty acid in the liver, leading to fatty liver [40,41]. Therefore it was obvious to address the question whether PPARα is involved in the βL-mediated improvement of alcoholic fatty liver. As shown in Fig. 7B, βL can directly enhance the DNA-binding activity of PPARα toward PPRE of acyl-CoA oxidase gene [22]. Furthermore, ethanol-mediated inhibition of DNA-binding activity was completely recovered in the presence of βL (Fig. 7C). In addition, expression levels of PPARα were significantly increased in ethanol/βL treated animals (Fig. 7A), suggesting that PPARα plays an important role in βL-mediated improvement of fatty liver. In conclusion, the enhancement of mitochondrial β-oxidation and inhibition of lipogenesis by βL will improve the alcohol-induced fatty liver in pathophysiological conditions and may be of use as a potential new therapy for alcoholic liver disease. Acknowledgments We would like to thank Dr. JW Lee (Korea Basic Science Institute, Korea) for TEM analysis. We also thank Drs. HS Choi (Chunnam National University, Korea), GR Keon (Chungnam National University, Korea) and CH Lee (Korea Research Institute for Bioscience and Biotechnology, Korea) for critical comments on the manuscript. This work was financially supported by the National Research Foundation of Korea (NRF)

grant funded by the Korea Government (MEST) (nos. 2007-0054932, 2012R1A1A2004714, 2012M3A9B6055302), and by a grant of the Korea Healthcare Technology R&D Project, Ministry for Health, Welfare & Family Affairs, Republic of Korea (HI10C0573). DPB is supported by funding from DEL Northern Ireland, Action Medical Research UK, BBSRC UK and Diabetes UK. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.cellsig.2013.11.020. References [1] E. Albano, Expert. Rev. Gastroenterol. Hepatol. 2 (2008) 749–759. [2] V. Purohit, D. Russo, P.M. Coates, Alcohol (Fayetteville N.Y.) 34 (2004) 3–8. [3] H.K. Seitz, C.S. Lieber, F. Stickel, M. Salaspuro, H.P. Schlemmer, Y. Horie, Alcohol. Clin. Exp. Res. 29 (2005) 1276–1281. [4] A. Reuben, Curr. Opin. Gastroenterol. 22 (2006) 263–271. [5] M.S. Bentle, K.E. Reinicke, Y. Dong, E.A. Bey, D.A. Boothman, Cancer Res. 67 (2007) 6936–6945. [6] K. Schaffner-Sabba, K.H. Schmidt-Ruppin, W. Wehrli, A.R. Schuerch, J.W. Wasley, J. Med. Chem. 27 (1984) 990–994. [7] E. Blanco, E.A. Bey, Y. Dong, B.D. Weinberg, D.M. Sutton, D.A. Boothman, J. Gao, J. Control. Release 122 (2007) 365–374. [8] E.K. Choi, K. Terai, I.M. Ji, Y.H. Kook, K.H. Park, E.T. Oh, R.J. Griffin, B.U. Lim, J.S. Kim, D.S. Lee, D.A. Boothman, M. Loren, C.W. Song, H.J. Park, Neoplasia New York N.Y. 9 (2007) 634–642. [9] H.J. Park, K.J. Ahn, S.D. Ahn, E. Choi, S.W. Lee, B. Williams, E.J. Kim, R. Griffin, E.A. Bey, W.G. Bornmann, J. Gao, H.J. Park, D.A. Boothman, C.W. Song, Int. J. Radiat. Oncol. Biol. Phys. 61 (2005) 212–219. [10] X. Sun, Y. Li, W. Li, B. Zhang, A.J. Wang, J. Sun, K. Mikule, Z. Jiang, C.J. Li, Cell Cycle Georgetown Tex. 5 (2006) 2029–2035.

S. Shin et al. / Cellular Signalling 26 (2014) 295–305 [11] M.S. Bentle, K.E. Reinicke, E.A. Bey, D.R. Spitz, D.A. Boothman, J. Biol. Chem. 281 (2006) 33684–33696. [12] M.S. Bentle, E.A. Bey, Y. Dong, K.E. Reinicke, D.A. Boothman, J. Mol. Hist. 37 (2006) 203–218. [13] J.H. Hwang, D.W. Kim, E.J. Jo, Y.K. Kim, Y.S. Jo, J.H. Park, S.K. Yoo, M.K. Park, T.H. Kwak, Y.L. Kho, J. Han, H.S. Choi, S.H. Lee, J.M. Kim, I. Lee, T. Kyung, C. Jang, J. Chung, G.R. Kweon, M. Shong, Diabetes 58 (2009) 965–974. [14] S.Y. Kim, N.H. Jeoung, C.J. Oh, Y.K. Choi, H.J. Lee, H.J. Kim, J.Y. Kim, J.H. Hwang, S. Tadi, Y.H. Yim, K.U. Lee, K.G. Park, S. Huh, K.N. Min, K.H. Jeong, M.G. Park, T.H. Kwak, G.R. Kweon, K. Inukai, M. Shong, I.K. Lee, Circ. Res. 104 (1) (2009) 842–850. [15] W. Oh, L. Abu-Elheiga, P. Kordari, Z. Gu, T. Shaikenov, S.S. Chirala, S.J. Wakil, Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 1384–1389. [16] T. Nakamura, K. Yoshimoto, Y. Nakayama, Y. Tomita, A. Ichihara, Proc. Natl. Acad. Sci. U. S. A. 80 (1983) 7229–7233. [17] M. Fischer, M. You, M. Matsumoto, D.W. Crabb, J. Biol. Chem. 278 (2003) 27997–28004. [18] M.A. Jay, J. Ren, Curr. Diabetes Rev. 3 (2007) 33–39. [19] T.K. Li, S.J. Yin, D.W. Crabb, S. O'Connor, V.A. Ramchandani, Alcohol. Clin. Exp. Res. 25 (2001) 136–144. [20] Y. Ido, Antioxid. Redox Signal. 9 (2007) 931–942. [21] W. Ying, Front. Biosci. 11 (2006) 3129–3148. [22] A. Galli, J. Pinaire, M. Fischer, R. Dorris, D.W. Crabb, J. Biol. Chem. 276 (2001) 68–75. [23] C.S. Lieber, D.P. Jones, L.M. Decarli, J. Clin. Invest. 44 (1965) 1009–1021. [24] D.W. Crabb, Recent Dev. Alcohol. 11 (1993) 207–230. [25] N. Grunnet, J. Kondrup, Alcohol. Clin. Exp. Res. 10 (1986) 64S–68S. [26] M.P. Salaspuro, S. Shaw, E. Jayatilleke, W.A. Ross, C.S. Lieber, Hepatol. Baltim. Md. 1 (1981) 33–38.

305

[27] C.P. Day, S.J. Yeaman, Biochim. Biophys. Acta 1215 (1994) 33–48. [28] N. Grunnet, J. Kondrup, J. Dich, Biochem. J. 228 (1985) 673–681. [29] R.A. Davis, J.R. Boogaerts, R.A. Borchardt, M. Malone-McNeal, J. ArchambaultSchexnayder, J. Biol. Chem. 260 (1985) 14137–14144. [30] A.A. Nanji, D. Zakim, in: D. Zakim (Ed.), Hepatology: A Text Book of Liver Disease, Saunders, Philadelphia, 1996, pp. 911–913. [31] M.J. Eckardt, S.E. File, G.L. Gessa, K.A. Grant, C. Guerri, P.L. Hoffman, H. Kalant, G.F. Koob, T.K. Li, B. Tabakoff, Alcohol. Clin. Exp. Res. 22 (1998) 998–1040. [32] M. Kopun, P. Propping, Eur. J. Clin. Pharmacol. 11 (1977) 337–344. [33] W.F. Bosron, T. Ehrig, T.K. Li, Sem. Liver Dis. 13 (1993) 126–135. [34] J.J. Pink, S.M. Planchon, C. Tagliarino, M.E. Varnes, D. Siegel, D.A. Boothman, J. Biol. Chem. 275 (2000) 5416–5424. [35] J.A. Baur, K.J. Pearson, N.L. Price, H.A. Jamieson, C. Lerin, A. Kalra, V.V. Prabhu, J.S. Allard, G. Lopez-Lluch, K. Lewis, P.J. Pistell, S. Poosala, K.G. Becker, O. Boss, D. Gwinn, M. Wang, S. Ramaswamy, K.W. Fishbein, R.G. Spencer, E.G. Lakatta, D. Le Couteur, R.J. Shaw, P. Navas, P. Puigserver, D.K. Ingram, R. de Cabo, D.A. Sinclair, Nature 444 (2006) 337–342. [36] J. Palming, K. Sjoholm, M. Jernas, T.C. Lystig, A. Gummesson, S. Romeo, L. Lonn, M. Lonn, B. Carlsson, L.M. Carlsson, J. Clin. Endocrinol. Metab. 92 (2007) 2346–2352. [37] V. Purohit, B. Gao, B.J. Song, Alcohol. Clin. Exp. Res. 33 (2009) 191–205. [38] F.P. Kuhajda, G.V. Ronnett, Curr. Opin. Investig. Drugs 8 (2007) 312–317. [39] G. Chinetti-Gbaguidi, J.C. Fruchart, B. Staels, Curr. Atheroscler. Rep. 7 (2005) 396–401. [40] T. Hashimoto, W.S. Cook, C. Qi, A.V. Yeldandi, J.K. Reddy, M.S. Rao, J. Biol. Chem. 275 (2000) 28918–28928. [41] T.C. Leone, C.J. Weinheimer, D.P. Kelly, Proc. Natl. Acad. Sci. U. S. A. 96 (1999) 7473–7478.