adipocyte atp-binding cassette g1 promotes triglyceride...

Eric Frisdal,1,2,3 Soazig Le Lay,4 Henri Hooton,2,5 Lucie Poupel,3 Maryline Olivier,1,2 Rohia Alili,2,3,5

Wanee Plengpanich,1,6 Elise F. Villard,1,2,3 Sophie Gilibert,1,2,3 Marie Lhomme,3

Alexandre Superville,1,2,3 Lobna Miftah-Alkhair,1 M. John Chapman,1,2 Geesje M. Dallinga-Thie,7

Nicolas Venteclef,2,3,5 Christine Poitou,2,3,5,8 Joan Tordjman,2,3,5 Philippe Lesnik,1,2,3

Anatol Kontush,1,2,3 Thierry Huby,1,2,3 Isabelle Dugail,2,3,5 Karine Clement,2,3,5,8 Maryse Guerin,1,2,3 andWilfried Le Goff1,2,3

Adipocyte ATP-BindingCassette G1 PromotesTriglyceride Storage, Fat MassGrowth, and Human ObesityDiabetes 2015;64:840–855 | DOI: 10.2337/db14-0245

The role of the ATP-binding cassette G1 (ABCG1) trans-porter in human pathophysiology is still largely unknown.Indeed, beyond its role in mediating free cholesterol effluxto HDL, the ABCG1 transporter equally promotes lipidaccumulation in a triglyceride (TG)-rich environmentthrough regulation of the bioavailability of lipoproteinlipase (LPL). Because both ABCG1 and LPL are expressedin adipose tissue, we hypothesized that ABCG1 is impli-cated in adipocyte TG storage and therefore could bea major actor in adipose tissue fat accumulation. Silenc-ing of Abcg1 expression by RNA interference in 3T3-L1preadipocytes compromised LPL-dependent TG accumu-lation during the initial phase of differentiation. Generationof stable Abcg1 knockdown 3T3-L1 adipocytes revealedthat Abcg1 deficiency reduces TG storage and diminisheslipid droplet size through inhibition of Pparg expression.Strikingly, local inhibition of adipocyte Abcg1 in adiposetissue from mice fed a high-fat diet led to a rapiddecrease of adiposity and weight gain. Analysis oftwo frequent ABCG1 single nucleotide polymor-phisms (rs1893590 [A/C] and rs1378577 [T/G]) in mor-bidly obese individuals indicated that elevated ABCG1expression in adipose tissue was associated with in-creased PPARg expression and adiposity concomitant

to increased fat mass and BMI (haplotype AT>GC). Thecritical role of ABCG1 in obesity was further confirmedin independent populations of severe obese and dia-betic obese individuals. This study identifies for the firsttime a major role of adipocyte ABCG1 in adiposity andfat mass growth and suggests that adipose ABCG1might represent a potential therapeutic target in obesity.

The ATP-binding cassette G1 (ABCG1) transporter hasbeen proposed to promote cellular cholesterol efflux toHDL (1), and targeted disruption of Abcg1 induced mas-sive tissue neutral lipid accumulation in mice fed a high-fat/high-cholesterol diet (2). The precise role of ABCG1 isstill a matter of debate, however, especially in humanpathophysiology (3).

We recently reported that two frequent ABCG1 single nu-cleotide polymorphisms (SNPs) (rs1893590 and rs1378577)were significantly associated with plasma lipoprotein lipase(LPL) activity in the Regression Growth Evaluation StatinStudy (REGRESS) population (4). Analysis of the relation-ship between the ABCG1 genotype and LPL led us to pro-pose a mechanism by which ABCG1 controls macrophageLPL activity through modulation of membrane lipid rafts

1INSERM, UMR_S1166, Team 4, Paris, France2Université Pierre et Marie Curie-Paris 6, Paris, France3Institute of Cardiometabolism and Nutrition, Pitié-Salpêtrière Hospital, Paris,France4INSERM, U1063, Angers, France5INSERM, U872, Nutriomique Team 7, Cordeliers Research Center, Paris, France6King Chulalongkorn Memorial Hospital, Thai Red Cross Society, Patumwan,Bangkok, Thailand7Laboratory of Vascular Medicine, AMC Amsterdam, Amsterdam, the Netherlands8Heart and Metabolism, Assistance-Publique Hôpitaux de Paris, Pitié-SalpêtrièreHospital, Paris, France

Corresponding author: Wilfried Le Goff, [email protected].

Received 13 February 2014 and accepted 10 September 2014.

This article contains Supplementary Data online at http://diabetes.diabetesjournals.org/lookup/suppl/doi:10.2337/db14-0245/-/DC1.

© 2015 by the American Diabetes Association. Readers may use this article aslong as the work is properly cited, the use is educational and not for profit, andthe work is not altered.

See accompanying article, p. 689.

840 Diabetes Volume 64, March 2015

OBESITY

STUDIES

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to promote intracellular lipid accumulation and foam cellformation in a triglyceride (TG)-rich context (4). Thus,beyond a role in sterol export to HDL, ABCG1 may equallycontribute to intracellular fatty acid accumulation andlipid storage in metabolic situations associated with in-creased concentrations of circulating TG-rich lipoproteins.Consistent with a role of Abcg1 in lipid storage, randominsertion of modified transposable elements of the P-familyin Drosophila melanogaster identified the CG17646 locus,the Drosophila ortholog of Abcg1, as a candidate gene forTG storage (5). Moreover, total ablation of Abcg1 in micefed a high-fat diet devoid of cholesterol (5) reduced TGaccumulation in adipose and liver tissues. However, thecellular mechanisms underlying this phenotype, and morespecifically the tissue-specific contribution of Abcg1, werenot elucidated. Considered together, these data promptedus to evaluate the function of ABCG1 in adipocytes, whichare professional cells for TG storage.

ABCG1 is expressed in adipocytes and in adipose tissueof mice that develop diet-induced obesity (5,6). Moreover,adipose tissue is a major source of LPL (7), which criticallycontrols TG accumulation by generating free fatty acidsfrom circulating lipoproteins (8).

Our data demonstrate that silencing of Abcg1 expres-sion in adipocytes reduced LPL activity and altered lipidhomeostasis. Moreover, Abcg1 deficiency resulted in in-hibition of Pparg expression and alteration of adipocytematuration. In vivo, local lentiviral-mediated targeting ofAbcg1 in adipose tissue rapidly reduced adiposity andhigh-fat diet-induced weight gain in mice. More strikingly,we observed that the ABCG1 genotype in humans wasassociated with fat mass formation and obesity in indepen-dent populations of obese individuals, thereby highlightingthe critical role of ABCG1 in the context of human obesity.Taken together, this study suggests that ABCG1 in adi-pose tissue might represent a future therapeutic target inmetabolic disorders associated with obesity.

RESEARCH DESIGN AND METHODS

Morbidly Obese PopulationMiddle-aged (42.0 6 0.04 years old) morbidly obesepatients (BMI 45.5 6 0.07 kg/m2; n = 1320) of Caucasianorigin (male-to-female ratio 0.33) were recruited at theDepartment of Nutrition at the Pitié-Salpêtrière Hospital,Paris, France (9). Patients were phenotyped for a series ofbioclinical variables. Body composition in this populationwas evaluated by biphotonic absorptiometry (dual-energyX-ray absorptiometry), as previously described (10). Allsubjects gave their informed written consent to partici-pate in the genetic study (a clinical research contract),which was approved by the local ethic committee.

The severe obese and diabetic obese populations aredescribed in the Supplementary Data. In a subset ofpatients who were candidates for bariatric surgery(patients who had the ABCG1 AT or GC haplotype), afteran overnight fast pieces of subcutaneous periumbilicaladipose tissue were sampled by needle biopsy under local

anesthesia (1% xylocaine). Biopsy samples were washed andstored in RNAlater preservative solution (Qiagen) at 280°Cuntil analysis. Total RNA was extracted from adipose tissuebiopsies using the RNeasy total RNA minikit (Qiagen). TotalRNA concentration and quality were confirmed usingan Agilent 2100 bioanalyzer (Agilent Technologies).

GenotypingABCG1 SNPs (rs1893590 and rs1378577) were genotypedusing the TaqMan SNP genotyping assay (Applied Biosys-tems). Hardy-Weinberg equilibrium was respected forboth ABCG1 SNPs in the obese populations studied.

Culture and Differentiation of AdipocytesThe 3T3-L1 preadipocytes (provided by Dr. J. Pairault,Paris) were maintained in DMEM supplemented with10% calf serum and 2 mmol/L glutamine. Differentiationof confluent preadipocytes was initiated with 0.25 mmol/Linsulin, 1.25 mmol/L dexamethasone, and 250 mmol/L3-isobutyl-methyl-1-xanthine in DMEM (4.5 g/L glucose)supplemented with 10% FBS. After 3 days, the culturemedium was switched to DMEM (4.5 g/L glucose) supple-mented with 10% FBS and 100 nmol/L insulin for 2 days.Then, 3T3 adipocytes were allowed to differentiate inDMEM (4.5 g/L glucose) containing 10% FBS, whichwas replaced every other day for 15 days.

Silencing of Abcg1 expression was performed byapplication of small interfering RNA (siRNA) oligonucleo-tides (Dharmacon) targeted to the cDNA sequence of themurine Abcg1 gene (NM_009593). Transfection of 3T3-L1preadipocytes and differentiated adipocytes with siRNA wasachieved using the Nucleofector technology (Lonza) accord-ing to the manufacturer’s protocol. For each experiment, 23106 cells and 100 pmol siRNA were diluted in 100 mL ofV solution and processed with the A-033 program.

Human preadipocytes (Promocell, Heidelberg, Ger-many) were cultured and differentiated as recommendedby the manufacturer. Differentiation efficiency was vali-dated by quantifying the induction of adipocyte markermRNA levels (ADIPOQ, LEP, and PPARg).

Generation of Stable Abcg1 Knockdown 3T3-L1AdipocytesControl short hairpin RNAs (shRNAs) and validated oli-gonucleotides encoding shRNAs targeting the cDNA se-quence of the murine Abcg1 gene (NM_009593) (R1 sense:59-GAT CCC CGG AAA GGT CTC CAA TCT CGT TCA AGAGAC GAG ATT GGA GAC CTT TCC TTT TTG GAA A-39,and R1 antisense: 59-AGC TTT TCC AAA AAG GAA AGGTCT CCA ATC TCG TCT CTT GAA CGA GAT TGG AGA CCTTTC CGG G-39; R2 sense: 59-GAT CCC CGA GAA GAC CTGCAC TGC GAT TCA AGA GAT CGC AGT GCA GGT CTTCTC TTT TTG GAA A-39, and R2 antisense: 59-AGC TTTTCC AAA AAG AGA AGA CCT GCA CTG CGA TCT CTTGAA TCG CAG TGC AGG TCT TCT CGG G-39) wereannealed and cloned into pSUPER, as previously described(11). The shRNA expression cassette then was transferredinto the XhoI/EcoRI site of the pRVH1-puro retroviral

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vector, and recombinant knockdown (KD) viruses weregenerated using the human Phoenix gag-pol packagingcell line (obtained from the National Gene Vector Biorepo-sitory, Indianapolis, IN), as previously described (12). 3T3-L1 preadipocytes were plated in 6-well plates (13 105 cellsper plate) in DMEM supplemented with 10% calf serum.After 48 h, the medium was aspirated and cells wereinfected with 1 mL of either supernatant from Phoenixcells containing control (R-Ctrl) or Abcg1 KD (R1 and R2)retroviral particles supplemented with 4 mg/mL of hexadi-methrine bromide (Polybrene; Sigma) or lentiviral particlesexpressing control shRNA (L-Ctrl) or shRNAs targeting thecDNA sequence of the murine Abcg1 gene (NM_009593)(L1, L2, L3) (Sigma). Selection of virus-transduced 3T3-L1preadipocytes was achieved by incubation with 4 mg/mLpuromycin (Invitrogen) for 6 days. Stable control andAbcg1 KD 3T3-L1 clones then were trypsinized andreseeded into DMEM (4.5 g/L glucose) supplementedwith 10% FBS and 4 mg/mL puromycin (Invitrogen) anddifferentiated into adipocytes, as described above.

RNA Extraction, Reverse Transcription, andQuantitative PCRTotal RNA from cell culture or tissues were extractedusing the NucleoSpin RNA II kit (Macherey-Nagel) orTRIzol reagents (Euromedex), respectively, according tothe manufacturer’s instructions. Reverse transcriptionand real-time quantitative PCR using a LightCyclerLC480 (Roche) were performed as previously described(13). Expression of mRNA levels was normalized to thehuman non-POU domain–containing, octamer-bindinghousekeeping gene (NONO); human a-tubulin (TUBA)and human heat shock protein 90 kDa a (cytosolic);class B member 1 (HSP90AB1) or mouse hypoxanthinephosphoribosyltransferase 1 (Hprt1); mouse non-POUdomain–containing, octamer-binding housekeeping gene(Nono); mouse heat shock protein 90 kDa a (cytosolic);class B member 1 (Hsp90ab1); mouse cyclophilin A (CycA);and mouse b-glucuronidase (Gusb) and mouse 18Sribosomal RNA (18S rRNA). Data were expressed as afold change in mRNA expression relative to controlvalues.

Adipocyte Diameter MeasurementsAdipose tissue pieces were minced and immediatelydigested by 200 mg/mL collagenase (Sigma) for 30 minat 37°C. For measurement of cell size, adipocyte suspen-sions then were visualized under a light microscope at-tached to a camera (TriCCD; Sony, France) and computerinterface. Adipocyte diameters were measured using PER-FECT IMAGE software (Numeris, Nanterre, France). Meanadipocyte diameter and volume were defined as the me-dian value of the distribution of adipocyte diameters of atleast 250 cells, measured by the same investigator.

Quantification of Apolipoprotein E SecretionSecreted apolipoprotein (apo)E in the culture media of3T3-L1 adipocytes was quantified by ELISA (Cloud-Clone

Corp., Houston, TX) according to the manufacturer’sinstructions and normalized to cell protein levels.

LPL Activity Measurement and Cellular LipidQuantificationLPL activity was determined with a 50-mL aliquot of cul-ture medium using an LPL activity assay kit (Roar, NewYork, NY), according to the manufacturer’s instructions.Intracellular total lipase activity was measured using a li-pase activity assay kit III (Sigma, Saint-Quentin Fallavier,France). Results were normalized to cell protein levels.When indicated, 3T3-L1 adipocytes were incubated for16 h at 37°C with either 250 mmol/L sphingomyelin(SM) (from chicken egg yolk; Sigma) or 3 units/mL sphin-gomyelinase (from Staphylococcus aureus; Sigma) to enrichor deplete membranes in sphingomyelin, respectively, orwith 1 mmol/L methyl-b-cyclodextrin (in PBS; Sigma) toremove free cholesterol at the plasma membrane. Quanti-fication of cellular TG and total and free cholesterol masswas performed as previously described (14).

Quantification of SM MassControl and Abcg1 stable KD (SKD) adipocytes (day 10)were incubated for 16 h at 37°C with 0.2% BSA (free ofendotoxins and free fatty acids) as an efficient acceptor forAbcg1-mediated SM efflux (15). Media were collected andcells were washed extensively with cold PBS. The extractionwas adapted from that described by Ivanova et al. (16). Inbrief, cells (30 mg of enzymatically quantified phospholi-pids) were supplemented with 3.2 mL of methanol acidifiedwith 0.1 N hydrogen chloride (1:1 v/v) containing 80 ng ofphosphatidylcholine (PC) 16:0/16:0 d9 (9 deuterium) and 3ng of lysophosphatidylcholine (LPC) 15:0. Cell media (600mL) were supplemented with 1 mL of methanol/0.1N HCl(1:1 v/v), 66 mL of 1N HCl and 660 mL methanol contain-ing 80 ng of PC 16:0/16:0 d9 (9 deuterium), and 3 ng ofLPC 15:0. The mixtures were vortexed for 1 min. Blank andcontrol samples were extracted in parallel with each batchto ensure quality control; each sample was corrected forblank readings. Chloroform was added to cells (800 mL)and media (1.16 mL), and the mixtures were vortexed for1 min and centrifuged at 3,600g for 10 min at 4°C. Lowerorganic phases were dried in nitrogen, resuspended, andtransferred into liquid chromatography–mass spectrometry(LC-MS) amber vials with inserts. LC-MS analysis and SMquantification were performed as previously described (17).The percentage of SM efflux was calculated as 100 3(medium SM)/(medium SM + cell SM).

Free Cholesterol Efflux AssaysDifferentiated 3T3-L1 adipocytes were incubated for 24 hwith the [3H]-cholesterol-labeled (1 mCi/mL) FBS (10%)in DMEM (4.5 g/L glucose) medium. Then the labeling me-dium was removed and cells were equilibrated in a serum-free medium containing 0.2% BSA for an additional 16-hperiod. Cellular free [3H]-cholesterol efflux to 20 mg/mL freeApoA-I (Sigma) or 30 mg/mL of HDL phospholipid wasassayed in serum-free medium containing 0.2% BSA for

842 Role of Adipocyte ABCG1 in Fat Mass Formation Diabetes Volume 64, March 2015

a 4-h chase period. Finally, culture media were harvestedand cleared of cellular debris by brief centrifugation. Cellradioactivity was determined by extraction in hexane–isopropanol (3:2), evaporation of the solvent, and liquidscintillation counting (Wallac Trilux 1450 Microbeta). Thepercentage of cholesterol efflux was calculated as 100 3(medium cpm)/(medium cpm + cell cpm).

Quantification of Lipid RaftsLipid rafts in differentiated adipocytes was detectedfollowing incubation with 1 mg/mL Alexa Fluor 594–conjugated cholera toxin subunit-b (Molecular Probes)for 15 min at 4°C, as previously described (4,6). Indeed,binding of cholera toxin subunit-b to the pentasaccharidechain of plasma membrane ganglioside GM1, which selec-tively partitions into lipid rafts (18), allows the reliabledetection of lipid rafts in live cells. After washing twicewith cold PBS, cells were detached from plates with trypsinand subjected to flow cytometry analysis on an LSR IIFORTESSA SORP (BD Biosciences). When indicated,images were captured using a Zeiss Axio Imager M2 mi-croscope with a 633 objective.

Injection of siRNA Targeting ABCG1 Expression inAdipose Tissue In VivoFour-week-old male C57BL/6 mice (Janvier, Le GenestSaint Isle, France) were fed a high-fat diet (45% fat;Brogaarden diet no. TD12451) for 4 weeks before the dayof injection. On the day of injection, mice were weighedand anesthetized with isoflurane; anesthesia was main-tained during the surgical procedure. A subabdominalincision was made and, using a 30-gauge needle, epidid-ymal fat pads were injected with 100 mL of lentiviralparticles (1.4 3 105 lentiviral transducing particles/mL)encoding either an shRNA designed to knock down mouseAbcg1 expression (Santa Cruz) or control shRNA lentiviralparticles encoding an shRNA that will not lead to thedegradation of any known cellular mRNA (Santa Cruz).Cell targeting of lentiviral particles into adipose tissue wasvisualized by injection of copGFP control lentiviral par-ticles (Santa Cruz). Dispersion of the injected volume inthe whole organ by this procedure was validated usinga colored dye in preliminary experiments (SupplementaryFig. 2A). Then, injected epididymal fat pads were placedback in the subabdominal cavity and the incision wassutured. Mice were fed a high-fat diet (60% fat; Brogaardendiet no. TD12492) for an additional 4-week period untilthey were killed. Food intake was monitored for a 3-dayperiod, and locomotion and activity were monitored overa 24-h period using the Activmeter actimetry system(Bioseb, Chaville, France). On the day they were killed,blood samples were collected in Microvette tubes (Sarstedt)by retroorbital bleeding under isoflurane anesthesia. Micewere weighed and killed, and epididymal fat pads wereisolated for RNA extraction, immunohistochemical analy-sis, and adipocyte diameter measurements. Liver and in-testine were collected, weighed, and flash frozen. Plasmasamples were analyzed with an Autoanalyzer (Konelab 20)

using reagent kits from Roche (total cholesterol), Diasys(free cholesterol, free fatty acids), and ThermoElectron(HDL cholesterol, TGs, and glucose).

Adipose Tissue Cell SortingEpididymal adipose tissue was excised from mice, minced,and digested in Hanks’ balanced salt solution (Gibco,Invitrogen, Cergy Pontoise, France) with 2.5 mg/mL col-lagenase D (Roche, Boulogne Billancourt, France) for 30min at 37°C and dissociated through a 200-mm pored cellstrainer. After decanting, adipocytes (supernatant) werewashed with a 10% sucrose solution and used for sub-sequent analyses. Stromal vascular fraction cells (bottom)were suspended in cold Hanks’ balanced salt solution con-taining 3% FBS and centrifuged at 1,500 rpm for 5 min.Recovered cells (200 mL) were stained with 1 mg/mL puri-fied anti-CD16/32 (Becton Dickinson, Franklin lakes, NJ)for 10 min at 4°C and for an additional 30 min with theappropriate dilution of specific antibodies. The followingpanel of antibodies was used: anti-CD45 (clone 30-F11),anti-CD11b (clone M1/70), anti-F4/80 (clone BM8), anti-CD64 (clone 290322), anti-CMH II (clone M5/114), andanti-CD31 (clone 390). Propidium iodide (PI) was used as aviability marker. Adipose tissue macrophages (ATMs) weredefined as PI2CD45+CD11b+F4/80+CD64+CD312 and en-dothelial cells as PI2CD31+. Cells were sorted using theMoFlo Astrios (Beckman Coulter, Villepinte, France) withSummit acquisition software and stored in RLT buffer(QIAgen, Courtaboeuf, France) at 280°C until used.

ImmunofluorescenceA portion of epididymal adipose tissue was fixed in 10%formalin overnight at 4°C before being embedded in paraf-fin. Paraffin tissue sections (5-mm thick) were dewaxed withxylene and graded ethanol, and antigens were unmasked byheating the sections in 10 mmol/L citrate buffer (pH 6.0) at750 W for 15 min in a domestic microwave. Then, sectionswere washed twice in PBS and saturated with 3% bovineserum before being stained with primary antibodies againstAbcg1 (Novus, Littleton, CO) and Perilipin (Progen, Heidel-berg, Germany). Alexa Fluor 568–conjugated anti–guineapig or Alexa Fluor 568– and 488–conjugated anti-rabbitwere used as secondary antibodies (Life Technologies, SaintAubin, France). Immunostained sections were examined ona Zeiss Axio Imager M2 microscope. Microscopy imageswere captured using AxioCam digital microscope camerasand AxioVision image processing software (Carl Zeiss Vi-sion, Germany). The specificity of antibodies was testedwith their isotype controls.

Western Blot AnalysisCell proteins were extracted using 200 mL M-PER reagent(Pierce) containing protease inhibitors and were subse-quently separated on a 4–12% Bis-Tris gel (Invitrogen).Proteins (25 mg/lane) were transferred to nitrocellulose andthe membrane was blocked with casein blocker solutionfor 1 h. Membranes then were incubated overnight at 4°Cwith a rabbit anti-Abcg1 (NB400–132; Novus), anti-Pparg




(C26H12) (2435, Cell Signaling), or anti-Fabp4 (2120, CellSignaling) or with guinea pig anti-perilipin (GP29, Progen)antibody diluted at 1:500 and revealed with either IRDye800CW-conjugated goat anti-rabbit or donkey anti–guinea pig (Li-Cor) at 1:10,000 for 1 h. Detection wasperformed using an Odyssey infrared imaging system(Li-Cor).

Statistical AnalysisLinkage disequilibrium between both SNPs was calculatedwith Haploview 4.1 software. Associations between phe-notypes and genotypes were tested with multivariatelinear regression models. All phenotypes were trans-formed to log10 before testing for associations. Geno-type–phenotype association tests were performed withR, version 2.8.2. Haplotypes and phenotypes were associ-ated with Hapstat 3.0 software. All models were adjustedfor age and sex; models testing associations with C-reactiveprotein and adiponectin also were adjusted for BMI. Fi-nally, models testing associations with TGs, total cholesterol,HDL, lipoprotein(a), ApoA-I, and apoB also were adjustedfor BMI and medical treatment for dyslipidemia. HOMAindex was calculated using the Homa2 method (http://www.dtu.ox.ac.uk/Homacalculator/index.php), which ledto the calculation of three different HOMAs (HOMA2S,HOMA2B, and HOMA of insulin resistance).

Data are shown as mean 6 SEM. Experiments wereperformed in triplicate, and values correspond to themean from at least three independent experiments. Com-parisons of two groups were performed by a two-tailedStudent t test, and comparisons of three or more groupswere performed by ANOVA with Newman–Keuls posttest.All statistical analyses were performed using Prism soft-ware from GraphPad, Inc. (San Diego, CA).

RESULTS

RNA Interference–Mediated Abcg1 TargetingDecreases LPL-Dependent Cell TG Storage inPreadipocytesAnalysis of Abcg1 expression during the course ofdifferentiation of 3T3-L1 preadipocytes into matureadipocytes indicated that Abcg1 mRNA levels were in-creased by ;40-fold during adipocyte differentiation(Fig. 1A). In agreement with a role for Abcg1 in TG stor-age, the accumulation of TGs was significantly correlatedwith Abcg1 mRNA levels (r2 = 0.9; P , 0.0001; Fig. 1B).

We previously reported that macrophage ABCG1promotes TG storage by modulating LPL activity (4). Totest whether the same mechanism operates in adipocytes,Abcg1 expression was silenced in 3T3-L1 preadipocytesusing specific siRNAs that did not alter adipocyte differ-entiation and viability. Inhibition of Abcg1 expression atboth mRNA and protein levels in 3T3-L1 preadipocytes(Fig. 1C and D) had no effect on Lpl mRNA expression(Fig. 1E). We observed a marked reduction of LPL activity(281%; P , 0.05) in media from Abcg1 KD adipocytes(Fig. 1F) compared with control cells. Moreover, decreasedLPL activity was associated with changes in cell surface

membrane properties; the binding of cholera toxin b sub-unit, which preferentially associates with lipid rafts, in-creased in cells transfected with Abcg1 RNA interference,as visualized by fluorescence microscopy and quantified byflow cytometry (+24%; P , 0.05; Fig. 1G).

Although the silencing of Abcg1 in preadipocytes(Abcg1 KD) was initiated before the addition of theadipocyte differentiation cocktail (day 0), a marked re-duction in intracellular TG accumulation occurred duringsubsequent adipocyte conversion (222% after 4 days ofdifferentiation; P , 0.05; Fig. 1H). Remarkably, additionof tetrahydrolipstatin, an inhibitor of LPL activity, overa period of 24 h compromised TG accumulation in controlcells, indicating the dependence of TG secretion on LPL inpreadipocytes (Fig. 1I, day 6). Conversely, the addition ofincreasing amounts of exogenous recombinant bovineLPL (bLPL) led to dose-dependent increases in intra-cellular TG mass in control 3T3-L1 preadipocytes thatparalleled those of LPL activity in culture media (Fig. 1I)(day 6). Together, these data indicate that TG accumu-lation during the initial phase of adipocyte differentia-tion is LPL dependent and is compromised by Abcg1invalidation.

Impaired Maturation in Stable Abcg1 KD 3T3-L1AdipocytesTo explore the impact of prolonged inhibition of Abcg1expression on adipocyte maturation, stable, fully differ-entiated Abcg1 KD 3T3-L1 adipocytes were generatedusing either lentiviral or retroviral particles expressingshRNAs targeted to distinct regions of the Abcg1 mRNA(Fig. 2).

SKD of Abcg1, either by lentiviruses (L1 to L3 vs. L-Ctrl) or by retroviruses (R1-R2 vs. R-Ctrl), led to a markedreduction in Pparg, Fabp4, C/ebpa, Perilipin, Cd36, and HslmRNA expression (Fig. 2B–G). The expression of someother genes, including Fas or C/ebpb, which also areknown to participate in adipocyte differentiation,remained unaffected. Notably, expression of Abca1 in adi-pocytes was recently reported to influence adipocyte lipidhomeostasis (19). This observation may be important be-cause Abcg1 deficiency in mouse macrophages was pro-posed to be compensated by an increase in Abca1expression (20). However, such elevation of Abca1 expres-sion was not observed in our conditions when Abcg1 wasknocked down in 3T3-L1 adipocytes (Fig. 2J). The reduc-tion of Pparg, Perilipin, and Fabp4 expression was con-firmed at the protein level (Fig. 3A–E). Strikingly, lipidaccumulation was reduced markedly in Abcg1 SKD adipo-cytes compared with stable 3T3-L1 control cells (Fig. 3F)and was confirmed by a decrease in lipid droplet diameter(246%; P , 0.05; Fig. 3G) and lower intracellular TGstorage (245%; P , 0.005; Fig. 3H). Moreover, 24-h treat-ment with exogenous bLPL partially rescued TG storage inAbcg1 SKD adipocytes, whereas total restoration was ob-served when bLPL was added throughout the course ofadipocyte maturation (from day 0 to day 10; Fig. 3I).


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Consistent with the well-established role of Abcg1in cholesterol transport, Abcg1-deficient adipocytesexhibited reduced capacity to promote free cholesterolefflux to HDL (231%; P , 0.005; Fig. 3J), even if freecholesterol mass was decreased in cells (260%; P ,

0.005; Fig. 3K). Such a decrease in intracellular choles-terol concentrations seemed to be accompanied byincreased amounts of mRNA of genes involved in cho-lesterol synthesis in Abcg1 SKD adipocytes (Supplemen-tary Fig. 1A–D).

Figure 1—Abcg1 silencing in preadipocytes affects TG storage in an LPL-dependent manner. Graphs show the levels of Abcg1 mRNA (A)correlation between Abcg1 mRNA and cellular TG storage during adipocyte differentiation (from day [D] 0 to day 13) (B). The efficiency ofthe Abcg1 KD in 3T3-L1 adipocytes was assessed by quantification of mRNA (C ) and protein concentrations (D). Levels of LplmRNA levels(E ), secreted LPL activity (F ), and membrane lipid raft formation (G) were evaluated in control (Ctrl) and Abcg1 KD 3T3-L1 adipocyte. G,right: Representative photographs of lipid rafts visualized by fluorescence microscopy (magnification, 363). Cellular TG content wasquantified during maturation (from day 0 to day 4) of control and Abcg1 KD 3T3-L1 preadipocytes into adipocytes (H). I: Impact ofa 24-h treatment with either tetrahydrolipstatin (THL) or increasing doses of bovine LPL (bLPL) on cellular TG mass and secreted LPLactivity during control adipocyte differentiation (day 6). Data are shown as mean 6 SEM. Experiments were performed in triplicate. *P <0.05 and **P < 0.005 vs. control cells.




Figure 2—Gene expression profile of stable Abcg1 KD 3T3-L1 mature adipocytes. Quantification of protein (A) and mRNA levels(B–J) in different stable 3T3-L1 adipocytes generated following infection with lentiviral (L) or retroviral (R) particles expressingcontrol shRNA or shRNAs targeting the cDNA sequence of mouse Abcg1 gene (L1, L2, L3 and R1, R2, respectively) following10 days of differentiation. A similar gene expression pattern was observed in all the Abcg1 KD adipocytes generated. Decreasedexpression of Pparg and Pparg target genes in stable Abcg1 KD adipocytes clones compared with stable control adipocytes. Dataare shown as mean 6 SEM. Experiments were performed in triplicate. *P < 0.05, **P < 0.005, and ***P < 0.0005 vs. respectivecontrol cells.


Figure 3—Stable Abcg1 KD compromises 3T3-L1 adipocyte lipid storage. A: Total protein concentrations were assessed by Western blotanalysis. Quantification of Abcg1 (B), Pparg (C), perilipin (D), and Fabp4 (E) protein concentrations in stable control (Ctrl) and Abcg1 SKD 3T3-L1 adipocytes following 10 days of differentiation. F: Phase-contrast photographs representative of lipid droplets in control and Abcg1 SKD3T3-L1 adipocytes visualized by microscopy (magnification, 320). G: Measurement of lipid droplets size and quantification of cellular TGs (H)and free cholesterol masses (K) in stable control and Abcg1 SKD 3T3-L1 adipocytes following 10 days of differentiation. I: Rescue of impairedTG storage in Abcg1 SKD adipocytes upon incubation with exogenous bovine LPL (bLPL) for the last 24 h (days = 1) or all along the 10 days ofthe maturation period (days = 10). J: Cellular [3H]-cholesterol efflux to apoA-I or HDL from mature control and Abcg1 SKD adipocytes (day 10).Data are shown as mean 6 SEM. Experiments were performed in triplicate. prot, protein. *P < 0.05 and **P < 0.005 vs. control cells.


Accumulation of SM in Stable Abcg1 KD AdipocytesReduces LPL-Dependent TG StorageABCG1 was reported to promote export of not only freecholesterol but also phospholipids such as SM (15), whichwas described to inhibit LPL activity (21,22). Therefore,we next address the hypothesis that an accumulation ofSM, found in large amounts in lipid raft domains, wasresponsible for the reduced LPL activity and the subse-quent impaired TG storage in Abcg1 SKD adipocytes.Quantification of SM mass by LC-MS revealed that stableAbcg1 KD in 3T3-L1 adipocytes was accompanied by a re-duced SM efflux (214%; P , 0.0005; Fig. 4A), which ledto a marked increase of intracellular SM content (48%;P , 0.0005; Fig. 4B) in those cells compared with controladipocytes. More strikingly, depletion of SM by sphingo-myelinase in Abcg1 SKD adipocytes restored LPL activityto a level comparable to that observed in control adipo-cytes (Fig. 4C) and promoted TG storage (Fig. 4D). Thecontribution of SM in this mechanism was furtherstrengthened by the observation that enriching control adi-pocytes with SM led to impaired TG storage, similar toAbcg1 SKD adipocytes (Fig. 4D). Treatment with 1 mmol/Lmethyl-b-cyclodextrin for 16 h, which removes cholesterolbut not SM from the plasma membrane (23,24), had noeffect on intracellular TG levels in Abcg1 SKD adipocytes(Supplementary Fig. 1E), suggesting that free cholesterol inlipid rafts was not responsible for the reduced TG storage inthose cells. Finally, because Abcg1 deficiency may be asso-ciated with increased apoE secretion (25), which may affectLPL activity (26), apoE secretion from control and Abcg1SKD adipocytes was examined. As shown in SupplementaryFig. 1F, no difference in apoE secretion was detected inAbcg1 SKD adipocytes compared with control cells.

Taken together, those results support the mechanismthrough which Abcg1 deficiency led to reduced SM effluxand a concomitant increase in SM content at the plasmamembrane, likely associated with lipid rafts, whichdecreases LPL activity and subsequent TG storage.

In Vivo Silencing of Abcg1 Expression Locally inAdipose Tissue Attenuates Fat Storage Upon High-FatDietTo further investigate the in vivo role of Abcg1 adipocytes,adipose tissue Abcg1 was silenced by local delivery of lenti-viral particles encoding either control shRNAs (L-Ctrl) orAbcg1 shRNAs (L-Abcg1) in epididymal adipose tissue.Injected C57BL/6 mice were maintained on a high-fat dietaccording to the experimental design presented in Fig. 5A.Expression of Abcg1 in adipocytes in adipose tissue frommice fed a high-fat diet (week 4) was visualized by immuno-fluorescence (Fig. 5B). Injection of lentiviral particles express-ing GFP alone into epididymal adipose tissue confirmedthat this strategy was efficient in targeting adipocytesfrom adipose tissue (W4; Fig. 5C and D). In an independentcontrol experiment, a marked decrease of Abcg1 stainingwas observed in an epididymal fat pad injected withL-Abcg1 4 weeks following injection compared with anepididymal fat pad injected with L-Ctrl (Fig. 5E), thus val-idating the KD of Abcg1 in adipose tissue. Such reducedAbcg1 expression in L-Abcg1 fat pads was highly repro-ducible when tested in multiple tissue samples treatedsimultaneously under identical experimental conditions.Quantification of adipose tissue Abcg1 mRNA indicatedthat a significant reduction of Abcg1 expression (245%;P , 0.05; Fig. 5F) was observed in injected fat pads ofL-Abcg1 mice, which mostly reflected the specific silencingof Abcg1 expression in adipocytes (247%; P, 0.0005) andto a lesser degree in ATMs; the expression of Abcg1 inthose cells was approximately threefold less abundantthan in adipocytes (Fig. 5G). No effect was observed inendothelial cells isolated from adipose tissue (Fig. 5G) orin other organs such as intestine (Fig. 5H) or liver (Fig. 5I).

All mice were maintained on a high-fat diet for 4 weeksfollowing injection, and weight gain was evaluated in the 2groups. Mice injected with L-Abcg1 gained less weight thanmice injected with L-Ctrl (224%; P, 0.05; Fig. 6A) and haddecreased epididymal fat mass (226%; P , 0.05; Fig. 6B),

Figure 4—Increased SM content in Abcg1 KD adipocytes is associated with altered LPL-dependent TG storage. SM efflux to BSA (A) andintracellular SMmass (B) in mature control (Ctrl) and Abcg1 SKD adipocytes (day 10). Secreted LPL activity (C) and intracellular TG mass (D)in adipocytes (day 10) enriched or depleted with either SM or sphingomyelinase (SMase) for 16 h. Data are shown as mean 6 SEM.Experiments were performed in triplicate. prot, protein. *P < 0.05 and ***P < 0.0005 vs. untreated control cells. #P < 0.05 vs. untreatedAbcg1 SKD cells.





Figure 5—KD of adipose tissue Abcg1 following shRNA lentiviral local delivery. A: Scheme of the experimental procedure. B: Abcg1 expressionin epididymal adipose tissue from a C57BL/6 mouse was visualized by fluorescence microscopy (magnification, 363). Arrows indicate Abcg1expression (green) in adipocytes (red, perilipin). Nuclei were counterstained with DAPI (blue). Recovery of green fluorescent protein (GFP)fluorescence in adipose tissue from one C57BL/6 mouse fed a high-fat diet (60% fat) 4 weeks following a local injection in the epididymaladipose tissue of lentiviral particles encoding either an shRNA control (C, left fat pad) or the full-length copGFP gene (D, right fat pad).Fluorescence was visualized by microscopy (magnification, 3400). E: Visualization of Abcg1 (red) by fluorescence microscopy (magnification,363) in epididymal fat pads following the local injection in the epididymal adipose tissue of lentiviral particles encoding either an shRNA control oran shRNA inhibiting mouse Abcg1 expression. Nuclei were counterstained with DAPI (blue). Quantification of Abcg1 mRNA levels in adiposetissue (mean cycle threshold: 25.18 in L-Ctrl) (F), intestine (mean cycle threshold: 31.78 in L-Ctrl) (H), and liver (mean cycle threshold: 29.95 inL-Ctrl) (I) from C57BL/6 mice fed a high-fat diet (60% fat) after 4 weeks following the local injection in the epididymal adipose tissue of lentiviralparticles encoding either a shRNA inhibiting mouse Abcg1 expression (L-Abcg1) or an shRNA control (L-Ctrl). G: Quantification of Abcg1mRNAlevels in adipocytes (mean cycle threshold: 27.95 in L-Ctrl), adipose tissue macrophages (ATMs) (mean cycle threshold: 31.01 in L-Ctrl), andendothelial cells (EC) (mean cycle threshold: 30.15 in L-Ctrl) isolated in adipose tissue from L-Abcg1 and L-Ctrl mice. Data are shown as mean6SEM (n = 11 mice per group). *P < 0.05 and ***P < 0.0005 vs. L-Ctrl.


suggesting less fat stored in adipose tissue. Indeed, meanadipocyte diameter was significantly smaller than that inepididymal adipose tissue from L-Ctrl mice (Fig. 6C). Inagreement, mRNA levels of leptin were decreased in adipose

tissue from L-Abcg1 mice compared with L-Ctrl mice(Fig. 6D). Analysis of food intake (Fig. 6E) and locomotoractivity (Fig. 6F and G) indicated that reduced weight gain inlocally injected mice did not result from an overt alteration

Figure 6—Abcg1 deficiency in adipose tissues affects the high-fat feeding response of mice. C57BL/6 mice fed a high-fat diet (40% fat)were injected with lentiviral particles encoding either an shRNA inhibiting murine Abcg1 expression (L-Abcg1) or an shRNA control (L-Ctrl)locally in the epididymal adipose tissue. Weight gain (A), epididymal fat mass (B), adipocyte diameter (C ), and mRNA levels (D, H–O) inepididymal adipose tissue were measured 4 weeks after the day of the injection (n = 10 mice per group). E: Food intake was measured inC57BL/6 mice fed a high-fat diet (60% fat) 4 weeks after the local injection of lentiviral particles encoding either an shRNA inhibiting mouseAbcg1 expression (L-Abcg1; n = 6) or an shRNA control (L-Ctrl; n = 6) in the epididymal adipose tissue. F and G: Locomotor activity in L-Ctrl(n = 8) and L-Abcg1 mice (n = 8) was monitored throughout the 4 weeks following the injection. Data are shown as mean6 SEM. *P< 0.05,**P < 0.005, and ***P < 0.0005 vs. L-Ctrl.


of energy balance. Additional analysis of metabolic par-ameters in L-Abcg1 and L-Ctrl mice is presented in Sup-plementary Table 1. The genes with downregulatedexpression in stable Abcg1 SKD mouse adipocytes gener-ated in culture were strikingly affected in Abcg1-silencedfat pads, namely, Pparg, Perilipin, Fabp4, Cd36, Hsl, andC/ebpa (Fig. 6H–M), whereas that of C/ebpb or Fas was notaltered (Fig. 6N and O). Expression of inflammatory andinsulin resistance genes in epididymal adipose tissue fromL-Ctrl and L-Abcg1 mice is shown in Supplementary Fig. 2.

Taken together, these results support a critical role ofAbcg1 in adipocyte lipid storage and indicate that localinhibition of Abcg1 in murine adipose tissue impaired fatstorage under high-fat diet.

Higher Expression of ABCG1 in Human Adipose TissueIs Associated With Increased Fat Mass andCorpulenceWe demonstrated that adipocyte Abcg1 contributes toTG storage and adiposity and hypothesized that elevatedexpression of ABCG1 might be associated with increasedfat mass in obese subjects. To evaluate this hypothesis, wetook the opportunity to examine functional ABCG1 SNPsand their association with adipose tissue gene expressionand body composition in obese individuals. Two frequentABCG1 SNPs (rs1378577 and rs1893590) located in the hu-man ABCG1 gene promoter were genotyped in a populationof 1,320 middle-aged morbidly obese patients (BMI .40kg/m2; mean BMI 45.5 6 0.04 kg/m2). The relative allelefrequencies for both ABCG1 SNPs (rs1893590, 2204A/C:20.73/0.27 and rs1378577, 2134T/G: 0.78/0.22) weresimilar to those observed in the REGRESS cohort (4). Aspreviously described (4), in vitro analysis of ABCG1 pro-moter activity according to ABCG1 haplotypes confirmedthat the frequent AT haplotype was associated withhigher transcriptional activity than the rare CG haplotype(Fig. 7A). Analysis of ABCG1 expression in biopsies ofadipose tissue isolated from a subset of obese patientsdisplaying either the AT or CG haplotypes revealed thatmRNA levels of ABCG1 were 27% (P , 0.05) more ele-vated in adipose tissues from patients carrying the AThaplotype relative to those carrying the CG haplotype(Fig. 7B). It is notable that ABCG1 expression in primaryhuman preadipocytes was significantly induced upon dif-ferentiation into adipocytes (Supplementary Fig. 3A).

In agreement with our data in Abcg1-deficient adipo-cytes, levels of mRNAs coding for genes involved inadipocyte differentiation (PPARg, CD36, PLIN1) were in-creased in adipose tissue from obese patients carryingthe AT haplotype compared with those carrying the CGhaplotype (Fig. 7C–E). By comparison, those of LPL wereunchanged (Fig. 7F). Adipose tissue mRNA levels of mac-rophage markers were not different between patientscarrying the AT haplotype and those carrying the GChaplotype (Supplementary Fig. 3B–D), suggesting thatthe number of macrophages present in adipose tissuewas similar.

ABCG1 expression in adipose tissue from obese patientswas positively correlated with the adipocyte diameter (r =0.51; P = 0.023; Fig. 7G). Importantly, obese individualscarrying the functional AT haplotype displayed signifi-cantly higher fat mass on dual-energy X-ray absorptiome-try, together with smaller fat-free mass (P, 0.05; Fig. 7Hand I), than those carrying the CG haplotype.

Moreover, the two ABCG1 SNPs were significantly asso-ciated with BMI; individuals carrying the 2204AA or the-134TT genotype displayed the highest BMI (P = 0.0034 andP = 0.011, respectively) (Fig. 7J). Haplotype analysis con-firmed that the AT haplotype (2204A/2134T) was signif-icantly associated with BMI (P = 0.006); moreover, BMIincreased in parallel with an increase in the amount ofthe AT haplotype (Fig. 7K). Association of both ABCG1SNPs with BMI was still observed after adjustment fordiabetes or HOMA index (Supplementary Table 2).ABCG1 SNPs rs1378577 and rs1893590 were not associ-ated with HOMA index after adjustment or not with BMI(data not shown). In addition, obese individuals carrying the2134TT genotype (rs1378577, the most frequent genotype)also displayed the highest fat mass index (P = 0.0242; Fig.7L); this effect was equally observed in subjects carrying the2204AA genotype (rs1893590, the most frequent geno-type). Of note, adiponectin and C-reactive protein concen-trations were significantly associated with both ABCG1 SNPsin this population (Supplementary Fig. 3). However no sig-nificant effect of ABCG1 SNPs on plasma lipid and apoconcentrations was observed (Supplementary Table 3).Thus elevated adipose expression of ABCG1 in obese indi-viduals carrying the AT haplotype is linked with increasedfat cell size, fat mass, and obesity and for the first time linksABCG1 genotype to obesity in humans.

Association of the ABCG1 genotype with obesity wasreplicated in two independent populations with distinctgrades of obesity: severely obese (35 , BMI , 40 kg/m2;mean BMI 39.2 6 8.5 kg/m2) and diabetic obese subjects(30 , BMI , 35 kg/m2; mean BMI 30.9 6 4.9 kg/m2).Genotyping of both ABCG1 SNPs (rs1378577 andrs1893590) in 216 type 2 diabetic obese subjects (30 ,BMI , 35 kg/m2; mean BMI 30.9 6 4.9 kg/m2) from theDiabetes Atorvastatin Lipid Intervention (DALI) Study (27)revealed that both ABCG1 SNPs were significantly associ-ated with waist-to-hip ratio (Supplementary Fig. 4A and B);individuals carrying the AT haplotype displayed a signifi-cantly higher waist-to-hip ratio (P , 0.005) than thosecarrying the CG haplotype (Supplementary Fig. 4C), con-firming the deleterious role of the AT haplotype in obesity.Finally, genotyping of the ABCG1 SNP rs1378577 in anindependent population of 595 severely obese subjects(35 , BMI , 40 kg/m2; mean BMI 39.2 6 8.5 kg/m2)(28) confirmed the association of the ABCG1 genotype withboth BMI and fat mass (Supplementary Fig. 4D and E).

DISCUSSION

In this study, we unraveled an unexpected role forAbcg1 in the control of lipid homeostasis in adipocytes.













Through a mechanism involving modulation of bothLPL activity and regulation of Pparg, Abcg1 seems to bea critical component in TG storage and adiposity. Thecentral role of Abcg1 in adipocytes is further strength-ened following the targeted silencing of its expressionlocally in adipose tissue in mice fed a high-fat diet,which led to a rapid reduction in adiposity and weightgain. Consistent with these findings, ABCG1 is associatedwith adiposity, fat mass, and obesity in obese individuals.

These results may initially seem to conflict with respectto the widely described role of Abcg1 in cellular freecholesterol efflux, where its inhibition leads to an increaserather than a decrease in intracellular lipid accumulation(2). In this way, Abcg1 can protect the cell from the ac-cumulation of free cholesterol, which is toxic to cell sur-vival. In agreement with such a role for Abcg1, wedemonstrated that silencing of Abcg1 expression in dif-ferentiated 3T3-L1 adipocytes was accompanied by

Figure 7—Increased adipose tissue ABCG1 expression and increased fat mass and obesity in obese individuals carrying the AT hap-lotype. A: Human ABCG1 promoter activity according to the CG and AT haplotypes. HepG2 cells were transiently transfected witha construct containing the proximal 1056 bp of the human promoter with either the 2204A/2134T (AT) haplotype or the 2204C/2134G (CG) haplotype. Luciferase activity is expressed in relative luciferase units (RLU) after normalization for b-galactosidase activity.Values are means6 SEMs of 5 independent experiments performed in triplicate. *P< 0.0005. B–F: Quantification of mRNA levels isolatedin adipose tissue biopsies from 10 morbidly obese women carrying either the AT or the CG haplotype. G: Correlation between adipocytediameter and ABCG1 mRNA levels in adipose tissue from morbidly obese women (n = 20). Fat mass (H) and fat-free mass (I) in obeseindividuals carrying either the AT (n = 102) or the CG haplotype (n = 22). Data are shown as mean 6 SEM. *P < 0.05 vs. CG haplotype,adjusted for age and sex. Association of the rs1378577 (2134T/G) and rs1893590 (2204A/C) ABCG1 SNP with BMI (J) and fat mass index(L) in a population of 1,320 middle-aged, severely morbidly obese patients (BMI = 45.47 6 0.002 kg/m2). K: Amount of 2204A/2134T (AT)haplotypes relative to BMI in obese individuals. AT/AT = 2. The effect of each SNP on BMI was analyzed by linear regression in anadditive, dominant, and recessive manner. The best model fitting the data is shown (dominant). All models were adjusted for age and sex.Data are shown as mean 6 SEM. *P < 0.05.


a significant reduction in free cholesterol efflux to HDL,thereby indicating that this mechanism is also operativein fat cells. However, in contrast to free cholesterol in theplasma membrane, the large amount of free cholesterolassociated with lipid droplets, which is closely propor-tional to TG storage in adipocytes and fat cell size, isnot mobilized for efflux (29). Mobilization of free choles-terol in lipid droplets therefore provides an alternativepathway for the protection of adipocytes from toxicity.This study supports the notion that although the role ofAbcg1 in cellular free cholesterol efflux mechanisms iscrucial in maintaining tissue lipid homeostasis in acholesterol-rich environment (2), Abcg1 contributes to cel-lular lipid (mostly TGs) accumulation and storage in high-fat metabolic states (4,5). Thus, Abcg1 deficiency (30–32)as well as expression of human ABCG1 in mice (2,33) fedan atherogenic diet enriched in cholesterol highlights therole of Abcg1 in protecting tissues, especially the lung,from lipid accumulation without any apparent changesin adipose tissue mass or adiposity. By contrast, and con-sistent with the mechanism described in the currentstudy, Abcg1 KO mice fed a high-fat diet devoid of cho-lesterol did not accumulate lipids in tissues but ratherexhibited reduced adipose tissue formation and were pro-tected against diet-induced obesity (5).

Our data support a mechanism by which Abcg1promotes TG storage through the requirement of bioactiveLPL (4); addition of exogenous LPL totally rescued theimpaired TG storage observed in Abcg1-deficient adipo-cytes. Our data led us to propose a mechanism by whichAbcg1-mediated export of SM contributes to sustainingLPL activity and TG storage in adipocytes. Indeed, LPLhydrolyzes triacylglycerol-rich lipoproteins, allowing thesubsequent release and uptake of fatty acids for intracellu-lar TG synthesis. LPL is expressed during the early stagesof adipocyte differentiation, when cell–cell contact occurs(34), and adipocyte-derived LPL was reported to be a keydeterminant for efficient TG storage and adipocyte hyper-trophy (8). In agreement with this mechanism, silencing ofAbcg1 expression in 3T3-L1 adipocytes was accompaniedby marked reduction in TG storage—an effect observed inthe early stages of adipocyte differentiation. Beyond itsrole in TG hydrolysis, LPL also was reported to facilitatelipoprotein uptake and thereby contribute to cellular cho-lesterol accumulation through this mechanism (14,35).Consistent with such a role for LPL, intracellular concen-trations of free cholesterol were markedly reduced in Abcg1KD adipocytes, a finding in agreement with the reducedlipid droplet size observed in Abcg1 SKD adipocytes. Therole of ABCG1 in adiposity and obesity through its actionon adipocyte LPL activity is supported by studies of humansubjects indicating that adipose tissue LPL activity is in-creased in obesity (36). Furthermore, several variants inthe LPL gene have been associated with obesity (37,38)and ob/ob mice with a specific Lpl deficiency in adiposetissue have displayed reduced weight and fat mass com-pared with control mice (39).

Alteration of Pparg and Pparg-targeted gene expres-sion likely results from the lesser abundance of intracel-lular cholesterol and fatty acid derivatives delivered byLPL hydrolysis upon silencing of Abcg1 expression in adi-pocytes. Fatty acid derivatives are ligands for Pparg acti-vation, and Pparg expression in adipocytes was reportedto be induced by its own activators and/or ligands, such asfatty acids (40–42), and cholesterol derivatives throughactivation of the liver X receptor (43). In agreement withthis mechanism, some fatty acid derivatives, whose de-livery into cells is mediated by LPL, are activators of Pparg(41). Interestingly, overexpression of a dominant negativemutant of Pparg that lacks the 16 COOH2 terminal aminoacids in 3T3-L1 adipocytes led to a reduction in the rate offree fatty acid uptake, TG storage, and adipocyte size and,more interestingly, to a decrease in the expression of theperilipin, Fabp4, Cd36, Hsl, and C/ebpa genes (44)—a phe-notype similar to that observed in the current study whenAbcg1 expression was silenced in those cells. However, themodest reduction of adipocyte diameter observed in Abcg1KD epididymal adipose tissue, in comparison with themore pronounced decrease of the tissue weight, leads usto suggest that the decreased Pparg expression could alsoalter adipocyte differentiation and adipocyte cell number invivo.

The participation of Pparg in the overall mechanism bywhich Abcg1 contributes to adiposity and obesity is sup-ported by the observation that the adipose-specific Ppargknockout mice displayed diminished weight gain and wereprotected against high-fat diet–induced obesity (45).Taken together, our findings indicate that modulationof LPL activity by Abcg1 in adipocytes might act as anintracellular signaling pathway that controls adipocytegrowth through activation of Pparg and contributes tothe formation of fat mass and the development of obesityin humans. Moreover, we reported that a higher expres-sion of ABCG1 in adipose tissue from obese individualscarrying the AT haplotype was associated with increasedPPARg, adiposity, fat mass, and BMI. However, al-though targeted deletion of Pparg in murine adiposetissue led to impaired growth of adipose tissue, itmust be kept in mind that those mice also exhibiteddeleterious metabolic consequences such as lipid accu-mulation in liver and in muscle and potentially insulinresistance (45,46).

In a previous study, Buchman et al. (5) first linkedAbcg1 to obesity in a mouse model in which Abcg1 wasknocked out in the whole body. Thus, ablation of Abcg1 inAbcg12/2mice reduces adipose cell size and hepatic steatosisand protects against diet-induced obesity. Although themechanisms underlying those effects were not elucidatedin this earlier study, Buchman et al. proposed that resis-tance of Abcg12/2 mice to diet-induced obesity likelyresulted in increased energy expenditure compared withAbcg1+/+ animals. In addition, a slight reduction of foodintake in Abcg12/2 mice also was reported by Buchmanet al.; this could contribute to the protective effect of Abcg1


deficiency. Our findings indicate that the contribution ofAbcg1 in adiposity and weight gain results from the criticalrole of adipocyte Abcg1 in adipose tissue, as indicated bythe targeted silencing of Abcg1 expression by RNA inter-ference locally in adipose tissue in mice fed a high-fat diet.Moreover, energy expenditure as well as food intake wasnot altered upon inhibition of Abcg1 expression in adiposetissue, which further reinforces the specific contribution ofadipose tissue Abcg1 in those effects.

Although silencing of Abcg1 expression was restricted toadipose tissue, we observed that the injection of lentiviralparticles locally in adipose tissue not only targets adipo-cytes but also equally targets macrophages present in thistissue. Although Abcg1 seems to be expressed more inadipocytes than in adipose tissue macrophages under ourexperimental conditions, this point may be critical becauseAbcg1 promotes LPL-mediated lipid accumulation inmacrophages in a TG-rich environment (4). However,a recent study reported that macrophage Lpl does notcontribute to adiposity and weight gain (47). Epididymalfat mass, lipid droplet size, and gene expression levels inadipose tissue, as well as body weight, were not alteredin macrophage Lpl knockout mice compared with con-trol mice.

Although further investigations are required to studythe impact of adipocyte Abcg1 deficiency in adipose tissue,our findings nonetheless support the contention thatABCG1 might represent an interesting pharmacologicaltarget in obesity, notably by reducing fat mass growth andweight gain in obese patients or in individuals prone todeveloping morbid obesity.

Acknowledgments. The authors are indebted to the patients for theircooperation. The INSERM U872 team thanks Assistance-Publique Hôpitaux deParis, Programme Hospitalier de recherche clinique (PHRC 1996 and 2002), forsupporting the genetic DNA bank on obesity.Funding. INSERM and UPMC (Parinov program) provided generous sup-port for these studies. M.O., E.F.V., and A.S. have received a ResearchFellowship from the French Ministry of Research and Technology. W.P. hasreceived a junior research fellowship from the French Embassy in Thailand.W.L.G. has received a PNRC award from INSERM. The ethics committee (ComitéProtection des personnes no. 1 Hôtel-Dieu) provided the ethics agreements. Thiswork was supported by the Fondation de France (to P.L., T.H., and W.L.G.) andby the French National Agency through the national program “Investissementsd’avenir” (ANR-10-IAHU-05).Duality of Interest. M.O., M.G., and W.L.G. are inventors of a patent (PCT/EP2011/073140), which covers the use of the ABCG1 gene as a marker anda target gene for treating obesity. No other potential conflicts of interest relevantto this article were reported.Author Contributions. E.F., S.L.L., H.H., L.P., M.O., R.A., W.P., E.F.V.,S.G., M.L., A.S., L.M.-A., M.J.C., G.M.D.-T., N.V., C.P., J.T., I.D., P.L., A.K., T.H.,M.G., and W.L.G. contributed to the experimental work and/or analyzed data.E.F., S.L.L., I.D., T.H., K.C., M.G., and W.L.G. developed the study. C.P. and K.C.recruited patients, performed phenotyping, and constituted the biobank. Allauthors wrote the manuscript. W.L.G. conceived, designed, and supervised thestudy. W.L.G. is the guarantor of this work and, as such, had full access to all thedata in the study and takes responsibility for the integrity of the data andthe accuracy of the data analysis.

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