The role of the ATP-binding cassette G1 (ABCG1) transporter in human pathophysiology is still largely unknown. Indeed, beyond its role in mediating free cholesterol efflux to HDL, the ABCG1 transporter equally promotes lipid accumulation in a triglyceride (TG)-rich environment through regulation of the bioavailability of lipoprotein lipase (LPL). Because both ABCG1 and LPL are expressed in adipose tissue, we hypothesized that ABCG1 is implicated in adipocyte TG storage and therefore could be a major actor in adipose tissue fat accumulation. Silencing of Abcg1 expression by RNA interference in 3T3-L1 preadipocytes compromised LPL-dependent TG accumulation during the initial phase of differentiation. Generation of stable Abcg1 knockdown 3T3-L1 adipocytes revealed that Abcg1 deficiency reduces TG storage and diminishes lipid droplet size through inhibition of Pparγ expression. Strikingly, local inhibition of adipocyte Abcg1 in adipose tissue from mice fed a high-fat diet led to a rapid decrease of adiposity and weight gain. Analysis of two frequent ABCG1 single nucleotide polymorphisms (rs1893590 [A/C] and rs1378577 [T/G]) in morbidly obese individuals indicated that elevated ABCG1 expression in adipose tissue was associated with increased PPARγ expression and adiposity concomitant to increased fat mass and BMI (haplotype AT>GC). The critical role of ABCG1 in obesity was further confirmed in independent populations of severe obese and diabetic obese individuals. This study identifies for the first time a major role of adipocyte ABCG1 in adiposity and fat mass growth and suggests that adipose ABCG1 might represent a potential therapeutic target in obesity.

The ATP-binding cassette G1 (ABCG1) transporter has been proposed to promote cellular cholesterol efflux to HDL (1), and targeted disruption of Abcg1 induced massive tissue neutral lipid accumulation in mice fed a high-fat/high-cholesterol diet (2). The precise role of ABCG1 is still a matter of debate, however, especially in human pathophysiology (3).

We recently reported that two frequent ABCG1 single nucleotide polymorphisms (SNPs) (rs1893590 and rs1378577) were significantly associated with plasma lipoprotein lipase (LPL) activity in the Regression Growth Evaluation Statin Study (REGRESS) population (4). Analysis of the relationship between the ABCG1 genotype and LPL led us to propose a mechanism by which ABCG1 controls macrophage LPL activity through modulation of membrane lipid rafts to promote intracellular lipid accumulation and foam cell formation in a triglyceride (TG)-rich context (4). Thus, beyond a role in sterol export to HDL, ABCG1 may equally contribute to intracellular fatty acid accumulation and lipid storage in metabolic situations associated with increased concentrations of circulating TG-rich lipoproteins. Consistent with a role of Abcg1 in lipid storage, random insertion of modified transposable elements of the P-family in Drosophila melanogaster identified the CG17646 locus, the Drosophila ortholog of Abcg1, as a candidate gene for TG storage (5). Moreover, total ablation of Abcg1 in mice fed a high-fat diet devoid of cholesterol (5) reduced TG accumulation in adipose and liver tissues. However, the cellular mechanisms underlying this phenotype, and more specifically the tissue-specific contribution of Abcg1, were not elucidated. Considered together, these data prompted us to evaluate the function of ABCG1 in adipocytes, which are professional cells for TG storage.

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

Our data demonstrate that silencing of Abcg1 expression in adipocytes reduced LPL activity and altered lipid homeostasis. Moreover, Abcg1 deficiency resulted in inhibition of Pparγ expression and alteration of adipocyte maturation. In vivo, local lentiviral-mediated targeting of Abcg1 in adipose tissue rapidly reduced adiposity and high-fat diet-induced weight gain in mice. More strikingly, we observed that the ABCG1 genotype in humans was associated with fat mass formation and obesity in independent populations of obese individuals, thereby highlighting the critical role of ABCG1 in the context of human obesity. Taken together, this study suggests that ABCG1 in adipose tissue might represent a future therapeutic target in metabolic disorders associated with obesity.

Morbidly Obese Population

Middle-aged (42.0 ± 0.04 years old) morbidly obese patients (BMI 45.5 ± 0.07 kg/m2; n = 1320) of Caucasian origin (male-to-female ratio 0.33) were recruited at the Department of Nutrition at the Pitié-Salpêtrière Hospital, Paris, France (9). Patients were phenotyped for a series of bioclinical variables. Body composition in this population was evaluated by biphotonic absorptiometry (dual-energy X-ray absorptiometry), as previously described (10). All subjects gave their informed written consent to participate in the genetic study (a clinical research contract), which was approved by the local ethic committee.

The severe obese and diabetic obese populations are described in the Supplementary Data. In a subset of patients who were candidates for bariatric surgery (patients who had the ABCG1 AT or GC haplotype), after an overnight fast pieces of subcutaneous periumbilical adipose tissue were sampled by needle biopsy under local anesthesia (1% xylocaine). Biopsy samples were washed and stored in RNAlater preservative solution (Qiagen) at −80°C until analysis. Total RNA was extracted from adipose tissue biopsies using the RNeasy total RNA minikit (Qiagen). Total RNA concentration and quality were confirmed using an Agilent 2100 bioanalyzer (Agilent Technologies).

Genotyping

ABCG1 SNPs (rs1893590 and rs1378577) were genotyped using the TaqMan SNP genotyping assay (Applied Biosystems). Hardy-Weinberg equilibrium was respected for both ABCG1 SNPs in the obese populations studied.

Culture and Differentiation of Adipocytes

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

Silencing of Abcg1 expression was performed by application of small interfering RNA (siRNA) oligonucleotides (Dharmacon) targeted to the cDNA sequence of the murine Abcg1 gene (NM_009593). Transfection of 3T3-L1 preadipocytes and differentiated adipocytes with siRNA was achieved using the Nucleofector technology (Lonza) according to the manufacturer’s protocol. For each experiment, 2 × 106 cells and 100 pmol siRNA were diluted in 100 μL of V solution and processed with the A-033 program.

Human preadipocytes (Promocell, Heidelberg, Germany) were cultured and differentiated as recommended by the manufacturer. Differentiation efficiency was validated by quantifying the induction of adipocyte marker mRNA levels (ADIPOQ, LEP, and PPARγ).

Generation of Stable Abcg1 Knockdown 3T3-L1 Adipocytes

Control short hairpin RNAs (shRNAs) and validated oligonucleotides encoding shRNAs targeting the cDNA sequence of the murine Abcg1 gene (NM_009593) (R1 sense: 5′-GAT CCC CGG AAA GGT CTC CAA TCT CGT TCA AGA GAC GAG ATT GGA GAC CTT TCC TTT TTG GAA A-3′, and R1 antisense: 5′-AGC TTT TCC AAA AAG GAA AGG TCT CCA ATC TCG TCT CTT GAA CGA GAT TGG AGA CCT TTC CGG G-3′; R2 sense: 5′-GAT CCC CGA GAA GAC CTG CAC TGC GAT TCA AGA GAT CGC AGT GCA GGT CTT CTC TTT TTG GAA A-3′, and R2 antisense: 5′-AGC TTT TCC AAA AAG AGA AGA CCT GCA CTG CGA TCT CTT GAA TCG CAG TGC AGG TCT TCT CGG G-3′) were annealed and cloned into pSUPER, as previously described (11). The shRNA expression cassette then was transferred into the XhoI/EcoRI site of the pRVH1-puro retroviral vector, and recombinant knockdown (KD) viruses were generated using the human Phoenix gag-pol packaging cell line (obtained from the National Gene Vector Biorepository, Indianapolis, IN), as previously described (12). 3T3-L1 preadipocytes were plated in 6-well plates (1 × 105 cells per plate) in DMEM supplemented with 10% calf serum. After 48 h, the medium was aspirated and cells were infected with 1 mL of either supernatant from Phoenix cells containing control (R-Ctrl) or Abcg1 KD (R1 and R2) retroviral particles supplemented with 4 µg/mL of hexadimethrine bromide (Polybrene; Sigma) or lentiviral particles expressing control shRNA (L-Ctrl) or shRNAs targeting the cDNA sequence of the murine Abcg1 gene (NM_009593) (L1, L2, L3) (Sigma). Selection of virus-transduced 3T3-L1 preadipocytes was achieved by incubation with 4 µg/mL puromycin (Invitrogen) for 6 days. Stable control and Abcg1 KD 3T3-L1 clones then were trypsinized and reseeded into DMEM (4.5 g/L glucose) supplemented with 10% FBS and 4 µg/mL puromycin (Invitrogen) and differentiated into adipocytes, as described above.

RNA Extraction, Reverse Transcription, and Quantitative PCR

Total RNA from cell culture or tissues were extracted using the NucleoSpin RNA II kit (Macherey-Nagel) or TRIzol reagents (Euromedex), respectively, according to the manufacturer’s instructions. Reverse transcription and real-time quantitative PCR using a LightCycler LC480 (Roche) were performed as previously described (13). Expression of mRNA levels was normalized to the human non-POU domain–containing, octamer-binding housekeeping gene (NONO); human α-tubulin (TUBA) and human heat shock protein 90 kDa α (cytosolic); class B member 1 (HSP90AB1) or mouse hypoxanthine phosphoribosyltransferase 1 (Hprt1); mouse non-POU domain–containing, octamer-binding housekeeping gene (Nono); mouse heat shock protein 90 kDa α (cytosolic); class B member 1 (Hsp90ab1); mouse cyclophilin A (CycA); and mouse β-glucuronidase (Gusb) and mouse 18S ribosomal RNA (18S rRNA). Data were expressed as a fold change in mRNA expression relative to control values.

Adipocyte Diameter Measurements

Adipose tissue pieces were minced and immediately digested by 200 µg/mL collagenase (Sigma) for 30 min at 37°C. For measurement of cell size, adipocyte suspensions then were visualized under a light microscope attached to a camera (TriCCD; Sony, France) and computer interface. Adipocyte diameters were measured using PERFECT IMAGE software (Numeris, Nanterre, France). Mean adipocyte diameter and volume were defined as the median value of the distribution of adipocyte diameters of at least 250 cells, measured by the same investigator.

Quantification of Apolipoprotein E Secretion

Secreted apolipoprotein (apo)E in the culture media of 3T3-L1 adipocytes was quantified by ELISA (Cloud-Clone Corp., Houston, TX) according to the manufacturer’s instructions and normalized to cell protein levels.

LPL Activity Measurement and Cellular Lipid Quantification

LPL activity was determined with a 50-µL aliquot of culture medium using an LPL activity assay kit (Roar, New York, NY), according to the manufacturer’s instructions. Intracellular total lipase activity was measured using a lipase activity assay kit III (Sigma, Saint-Quentin Fallavier, France). Results were normalized to cell protein levels. When indicated, 3T3-L1 adipocytes were incubated for 16 h at 37°C with either 250 μmol/L sphingomyelin (SM) (from chicken egg yolk; Sigma) or 3 units/mL sphingomyelinase (from Staphylococcus aureus; Sigma) to enrich or deplete membranes in sphingomyelin, respectively, or with 1 mmol/L methyl-β-cyclodextrin (in PBS; Sigma) to remove free cholesterol at the plasma membrane. Quantification of cellular TG and total and free cholesterol mass was performed as previously described (14).

Quantification of SM Mass

Control and Abcg1 stable KD (SKD) adipocytes (day 10) were incubated for 16 h at 37°C with 0.2% BSA (free of endotoxins and free fatty acids) as an efficient acceptor for Abcg1-mediated SM efflux (15). Media were collected and cells were washed extensively with cold PBS. The extraction was adapted from that described by Ivanova et al. (16). In brief, cells (30 µg of enzymatically quantified phospholipids) were supplemented with 3.2 mL of methanol acidified with 0.1 N hydrogen chloride (1:1 v/v) containing 80 ng of phosphatidylcholine (PC) 16:0/16:0 d9 (9 deuterium) and 3 ng of lysophosphatidylcholine (LPC) 15:0. Cell media (600 μL) were supplemented with 1 mL of methanol/0.1N HCl (1:1 v/v), 66 µL of 1N HCl and 660 µL methanol containing 80 ng of PC 16:0/16:0 d9 (9 deuterium), and 3 ng of LPC 15:0. The mixtures were vortexed for 1 min. Blank and control samples were extracted in parallel with each batch to ensure quality control; each sample was corrected for blank readings. Chloroform was added to cells (800 µL) and media (1.16 mL), and the mixtures were vortexed for 1 min and centrifuged at 3,600g for 10 min at 4°C. Lower organic phases were dried in nitrogen, resuspended, and transferred into liquid chromatography–mass spectrometry (LC-MS) amber vials with inserts. LC-MS analysis and SM quantification were performed as previously described (17). The percentage of SM efflux was calculated as 100 × (medium SM)/(medium SM + cell SM).

Free Cholesterol Efflux Assays

Differentiated 3T3-L1 adipocytes were incubated for 24 h with the [3H]-cholesterol-labeled (1 µCi/mL) FBS (10%) in DMEM (4.5 g/L glucose) medium. Then the labeling medium was removed and cells were equilibrated in a serum-free medium containing 0.2% BSA for an additional 16-h period. Cellular free [3H]-cholesterol efflux to 20 μg/mL free ApoA-I (Sigma) or 30 µg/mL of HDL phospholipid was assayed in serum-free medium containing 0.2% BSA for a 4-h chase period. Finally, culture media were harvested and cleared of cellular debris by brief centrifugation. Cell radioactivity was determined by extraction in hexane–isopropanol (3:2), evaporation of the solvent, and liquid scintillation counting (Wallac Trilux 1450 Microbeta). The percentage of cholesterol efflux was calculated as 100 × (medium cpm)/(medium cpm + cell cpm).

Quantification of Lipid Rafts

Lipid rafts in differentiated adipocytes was detected following incubation with 1 μg/mL Alexa Fluor 594–conjugated cholera toxin subunit-β (Molecular Probes) for 15 min at 4°C, as previously described (4,6). Indeed, binding of cholera toxin subunit-β to the pentasaccharide chain of plasma membrane ganglioside GM1, which selectively partitions into lipid rafts (18), allows the reliable detection of lipid rafts in live cells. After washing twice with cold PBS, cells were detached from plates with trypsin and subjected to flow cytometry analysis on an LSR II FORTESSA SORP (BD Biosciences). When indicated, images were captured using a Zeiss Axio Imager M2 microscope with a 63× objective.

Injection of siRNA Targeting ABCG1 Expression in Adipose Tissue In Vivo

Four-week-old male C57BL/6 mice (Janvier, Le Genest Saint Isle, France) were fed a high-fat diet (45% fat; Brogaarden diet no. TD12451) for 4 weeks before the day of injection. On the day of injection, mice were weighed and anesthetized with isoflurane; anesthesia was maintained during the surgical procedure. A subabdominal incision was made and, using a 30-gauge needle, epididymal fat pads were injected with 100 μL of lentiviral particles (1.4 × 105 lentiviral transducing particles/mL) encoding either an shRNA designed to knock down mouse Abcg1 expression (Santa Cruz) or control shRNA lentiviral particles encoding an shRNA that will not lead to the degradation of any known cellular mRNA (Santa Cruz). Cell targeting of lentiviral particles into adipose tissue was visualized by injection of copGFP control lentiviral particles (Santa Cruz). Dispersion of the injected volume in the whole organ by this procedure was validated using a colored dye in preliminary experiments (Supplementary Fig. 2A). Then, injected epididymal fat pads were placed back in the subabdominal cavity and the incision was sutured. Mice were fed a high-fat diet (60% fat; Brogaarden diet no. TD12492) for an additional 4-week period until they were killed. Food intake was monitored for a 3-day period, and locomotion and activity were monitored over a 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. Mice were weighed and killed, and epididymal fat pads were isolated for RNA extraction, immunohistochemical analysis, and adipocyte diameter measurements. Liver and intestine were collected, weighed, and flash frozen. Plasma samples 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 Sorting

Epididymal adipose tissue was excised from mice, minced, and digested in Hanks’ balanced salt solution (Gibco, Invitrogen, Cergy Pontoise, France) with 2.5 mg/mL collagenase D (Roche, Boulogne Billancourt, France) for 30 min at 37°C and dissociated through a 200-μm pored cell strainer. After decanting, adipocytes (supernatant) were washed with a 10% sucrose solution and used for subsequent analyses. Stromal vascular fraction cells (bottom) were suspended in cold Hanks’ balanced salt solution containing 3% FBS and centrifuged at 1,500 rpm for 5 min. Recovered cells (200 µL) were stained with 1 μg/mL purified anti-CD16/32 (Becton Dickinson, Franklin lakes, NJ) for 10 min at 4°C and for an additional 30 min with the appropriate dilution of specific antibodies. The following panel 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), and anti-CD31 (clone 390). Propidium iodide (PI) was used as a viability marker. Adipose tissue macrophages (ATMs) were defined as PICD45+CD11b+F4/80+CD64+CD31 and endothelial cells as PICD31+. Cells were sorted using the MoFlo Astrios (Beckman Coulter, Villepinte, France) with Summit acquisition software and stored in RLT buffer (QIAgen, Courtaboeuf, France) at −80°C until used.

Immunofluorescence

A portion of epididymal adipose tissue was fixed in 10% formalin overnight at 4°C before being embedded in paraffin. Paraffin tissue sections (5-μm thick) were dewaxed with xylene and graded ethanol, and antigens were unmasked by heating the sections in 10 mmol/L citrate buffer (pH 6.0) at 750 W for 15 min in a domestic microwave. Then, sections were washed twice in PBS and saturated with 3% bovine serum before being stained with primary antibodies against Abcg1 (Novus, Littleton, CO) and Perilipin (Progen, Heidelberg, Germany). Alexa Fluor 568–conjugated anti–guinea pig or Alexa Fluor 568– and 488–conjugated anti-rabbit were used as secondary antibodies (Life Technologies, Saint Aubin, France). Immunostained sections were examined on a Zeiss Axio Imager M2 microscope. Microscopy images were captured using AxioCam digital microscope cameras and AxioVision image processing software (Carl Zeiss Vision, Germany). The specificity of antibodies was tested with their isotype controls.

Western Blot Analysis

Cell proteins were extracted using 200 μL M-PER reagent (Pierce) containing protease inhibitors and were subsequently separated on a 4–12% Bis-Tris gel (Invitrogen). Proteins (25 µg/lane) were transferred to nitrocellulose and the membrane was blocked with casein blocker solution for 1 h. Membranes then were incubated overnight at 4°C with a rabbit anti-Abcg1 (NB400–132; Novus), anti-Pparγ (C26H12) (2435, Cell Signaling), or anti-Fabp4 (2120, Cell Signaling) or with guinea pig anti-perilipin (GP29, Progen) antibody diluted at 1:500 and revealed with either IRDye 800CW-conjugated goat anti-rabbit or donkey anti–guinea pig (Li-Cor) at 1:10,000 for 1 h. Detection was performed using an Odyssey infrared imaging system (Li-Cor).

Statistical Analysis

Linkage disequilibrium between both SNPs was calculated with Haploview 4.1 software. Associations between phenotypes and genotypes were tested with multivariate linear regression models. All phenotypes were transformed to log10 before testing for associations. Genotype–phenotype association tests were performed with R, version 2.8.2. Haplotypes and phenotypes were associated with Hapstat 3.0 software. All models were adjusted for age and sex; models testing associations with C-reactive protein and adiponectin also were adjusted for BMI. Finally, models testing associations with TGs, total cholesterol, HDL, lipoprotein(a), ApoA-I, and apoB also were adjusted for BMI and medical treatment for dyslipidemia. HOMA index was calculated using the Homa2 method (http://www.dtu.ox.ac.uk/Homacalculator/index.php), which led to the calculation of three different HOMAs (HOMA2S, HOMA2B, and HOMA of insulin resistance).

Data are shown as mean ± SEM. Experiments were performed in triplicate, and values correspond to the mean from at least three independent experiments. Comparisons of two groups were performed by a two-tailed Student t test, and comparisons of three or more groups were performed by ANOVA with Newman–Keuls posttest. All statistical analyses were performed using Prism software from GraphPad, Inc. (San Diego, CA).

RNA Interference–Mediated Abcg1 Targeting Decreases LPL-Dependent Cell TG Storage in Preadipocytes

Analysis of Abcg1 expression during the course of differentiation of 3T3-L1 preadipocytes into mature adipocytes indicated that Abcg1 mRNA levels were increased by ∼40-fold during adipocyte differentiation (Fig. 1A). In agreement with a role for Abcg1 in TG storage, the accumulation of TGs was significantly correlated with Abcg1 mRNA levels (r2 = 0.9; P < 0.0001; Fig. 1B).

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 of the Abcg1 KD in 3T3-L1 adipocytes was assessed by quantification of mRNA (C) and protein concentrations (D). Levels of Lpl mRNA 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, ×63). Cellular TG content was quantified during maturation (from day 0 to day 4) of control and Abcg1 KD 3T3-L1 preadipocytes into adipocytes (H). I: Impact of a 24-h treatment with either tetrahydrolipstatin (THL) or increasing doses of bovine LPL (bLPL) on cellular TG mass and secreted LPL activity during control adipocyte differentiation (day 6). Data are shown as mean ± SEM. Experiments were performed in triplicate. *P < 0.05 and **P < 0.005 vs. control cells.

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 of the Abcg1 KD in 3T3-L1 adipocytes was assessed by quantification of mRNA (C) and protein concentrations (D). Levels of Lpl mRNA 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, ×63). Cellular TG content was quantified during maturation (from day 0 to day 4) of control and Abcg1 KD 3T3-L1 preadipocytes into adipocytes (H). I: Impact of a 24-h treatment with either tetrahydrolipstatin (THL) or increasing doses of bovine LPL (bLPL) on cellular TG mass and secreted LPL activity during control adipocyte differentiation (day 6). Data are shown as mean ± SEM. Experiments were performed in triplicate. *P < 0.05 and **P < 0.005 vs. control cells.

Close modal

We previously reported that macrophage ABCG1 promotes TG storage by modulating LPL activity (4). To test whether the same mechanism operates in adipocytes, Abcg1 expression was silenced in 3T3-L1 preadipocytes using specific siRNAs that did not alter adipocyte differentiation and viability. Inhibition of Abcg1 expression at both 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 (−81%; P < 0.05) in media from Abcg1 KD adipocytes (Fig. 1F) compared with control cells. Moreover, decreased LPL activity was associated with changes in cell surface membrane properties; the binding of cholera toxin β subunit, which preferentially associates with lipid rafts, increased in cells transfected with Abcg1 RNA interference, as visualized by fluorescence microscopy and quantified by flow cytometry (+24%; P < 0.05; Fig. 1G).

Although the silencing of Abcg1 in preadipocytes (Abcg1 KD) was initiated before the addition of the adipocyte differentiation cocktail (day 0), a marked reduction in intracellular TG accumulation occurred during subsequent adipocyte conversion (−22% after 4 days of differentiation; P < 0.05; Fig. 1H). Remarkably, addition of tetrahydrolipstatin, an inhibitor of LPL activity, over a period of 24 h compromised TG accumulation in control cells, indicating the dependence of TG secretion on LPL in preadipocytes (Fig. 1I, day 6). Conversely, the addition of increasing amounts of exogenous recombinant bovine LPL (bLPL) led to dose-dependent increases in intracellular TG mass in control 3T3-L1 preadipocytes that paralleled those of LPL activity in culture media (Fig. 1I) (day 6). Together, these data indicate that TG accumulation during the initial phase of adipocyte differentiation is LPL dependent and is compromised by Abcg1 invalidation.

Impaired Maturation in Stable Abcg1 KD 3T3-L1 Adipocytes

To explore the impact of prolonged inhibition of Abcg1 expression on adipocyte maturation, stable, fully differentiated Abcg1 KD 3T3-L1 adipocytes were generated using either lentiviral or retroviral particles expressing shRNAs targeted to distinct regions of the Abcg1 mRNA (Fig. 2).

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 expressing control shRNA or shRNAs targeting the cDNA sequence of mouse Abcg1 gene (L1, L2, L3 and R1, R2, respectively) following 10 days of differentiation. A similar gene expression pattern was observed in all the Abcg1 KD adipocytes generated. Decreased expression of Pparγ and Pparγ target genes in stable Abcg1 KD adipocytes clones compared with stable control adipocytes. Data are shown as mean ± SEM. Experiments were performed in triplicate. *P < 0.05, **P < 0.005, and ***P < 0.0005 vs. respective 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 expressing control shRNA or shRNAs targeting the cDNA sequence of mouse Abcg1 gene (L1, L2, L3 and R1, R2, respectively) following 10 days of differentiation. A similar gene expression pattern was observed in all the Abcg1 KD adipocytes generated. Decreased expression of Pparγ and Pparγ target genes in stable Abcg1 KD adipocytes clones compared with stable control adipocytes. Data are shown as mean ± SEM. Experiments were performed in triplicate. *P < 0.05, **P < 0.005, and ***P < 0.0005 vs. respective control cells.

Close modal

SKD of Abcg1, either by lentiviruses (L1 to L3 vs. L-Ctrl) or by retroviruses (R1-R2 vs. R-Ctrl), led to a marked reduction in Pparγ, Fabp4, C/ebpα, Perilipin, Cd36, and Hsl mRNA expression (Fig. 2B–G). The expression of some other genes, including Fas or C/ebpβ, which also are known to participate in adipocyte differentiation, remained unaffected. Notably, expression of Abca1 in adipocytes was recently reported to influence adipocyte lipid homeostasis (19). This observation may be important because Abcg1 deficiency in mouse macrophages was proposed to be compensated by an increase in Abca1 expression (20). However, such elevation of Abca1 expression was not observed in our conditions when Abcg1 was knocked down in 3T3-L1 adipocytes (Fig. 2J). The reduction of Pparγ, Perilipin, and Fabp4 expression was confirmed at the protein level (Fig. 3A–E). Strikingly, lipid accumulation was reduced markedly in Abcg1 SKD adipocytes compared with stable 3T3-L1 control cells (Fig. 3F) and was confirmed by a decrease in lipid droplet diameter (−46%; P < 0.05; Fig. 3G) and lower intracellular TG storage (−45%; P < 0.005; Fig. 3H). Moreover, 24-h treatment with exogenous bLPL partially rescued TG storage in Abcg1 SKD adipocytes, whereas total restoration was observed when bLPL was added throughout the course of adipocyte maturation (from day 0 to day 10; Fig. 3I).

Figure 3

Stable Abcg1 KD compromises 3T3-L1 adipocyte lipid storage. A: Total protein concentrations were assessed by Western blot analysis. Quantification of Abcg1 (B), Pparγ (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 SKD 3T3-L1 adipocytes visualized by microscopy (magnification, ×20). 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 impaired TG 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 of the 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 ± SEM. Experiments were performed in triplicate. prot, protein. *P < 0.05 and **P < 0.005 vs. control cells.

Figure 3

Stable Abcg1 KD compromises 3T3-L1 adipocyte lipid storage. A: Total protein concentrations were assessed by Western blot analysis. Quantification of Abcg1 (B), Pparγ (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 SKD 3T3-L1 adipocytes visualized by microscopy (magnification, ×20). 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 impaired TG 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 of the 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 ± SEM. Experiments were performed in triplicate. prot, protein. *P < 0.05 and **P < 0.005 vs. control cells.

Close modal

Consistent with the well-established role of Abcg1 in cholesterol transport, Abcg1-deficient adipocytes exhibited reduced capacity to promote free cholesterol efflux to HDL (−31%; P < 0.005; Fig. 3J), even if free cholesterol mass was decreased in cells (−60%; P < 0.005; Fig. 3K). Such a decrease in intracellular cholesterol concentrations seemed to be accompanied by increased amounts of mRNA of genes involved in cholesterol synthesis in Abcg1 SKD adipocytes (Supplementary Fig. 1A–D).

Accumulation of SM in Stable Abcg1 KD Adipocytes Reduces LPL-Dependent TG Storage

ABCG1 was reported to promote export of not only free cholesterol but also phospholipids such as SM (15), which was described to inhibit LPL activity (21,22). Therefore, we next address the hypothesis that an accumulation of SM, found in large amounts in lipid raft domains, was responsible for the reduced LPL activity and the subsequent impaired TG storage in Abcg1 SKD adipocytes. Quantification of SM mass by LC-MS revealed that stable Abcg1 KD in 3T3-L1 adipocytes was accompanied by a reduced SM efflux (−14%; P < 0.0005; Fig. 4A), which led to a marked increase of intracellular SM content (48%; P < 0.0005; Fig. 4B) in those cells compared with control adipocytes. More strikingly, depletion of SM by sphingomyelinase in Abcg1 SKD adipocytes restored LPL activity to a level comparable to that observed in control adipocytes (Fig. 4C) and promoted TG storage (Fig. 4D). The contribution of SM in this mechanism was further strengthened by the observation that enriching control adipocytes with SM led to impaired TG storage, similar to Abcg1 SKD adipocytes (Fig. 4D). Treatment with 1 mmol/L methyl-β-cyclodextrin for 16 h, which removes cholesterol but not SM from the plasma membrane (23,24), had no effect on intracellular TG levels in Abcg1 SKD adipocytes (Supplementary Fig. 1E), suggesting that free cholesterol in lipid rafts was not responsible for the reduced TG storage in those cells. Finally, because Abcg1 deficiency may be associated with increased apoE secretion (25), which may affect LPL activity (26), apoE secretion from control and Abcg1 SKD adipocytes was examined. As shown in Supplementary Fig. 1F, no difference in apoE secretion was detected in Abcg1 SKD adipocytes compared with control cells.

Figure 4

Increased SM content in Abcg1 KD adipocytes is associated with altered LPL-dependent TG storage. SM efflux to BSA (A) and intracellular SM mass (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 ± SEM. Experiments were performed in triplicate. prot, protein. *P < 0.05 and ***P < 0.0005 vs. untreated control cells. #P < 0.05 vs. untreated Abcg1 SKD cells.

Figure 4

Increased SM content in Abcg1 KD adipocytes is associated with altered LPL-dependent TG storage. SM efflux to BSA (A) and intracellular SM mass (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 ± SEM. Experiments were performed in triplicate. prot, protein. *P < 0.05 and ***P < 0.0005 vs. untreated control cells. #P < 0.05 vs. untreated Abcg1 SKD cells.

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Taken together, those results support the mechanism through which Abcg1 deficiency led to reduced SM efflux and a concomitant increase in SM content at the plasma membrane, likely associated with lipid rafts, which decreases LPL activity and subsequent TG storage.

In Vivo Silencing of Abcg1 Expression Locally in Adipose Tissue Attenuates Fat Storage Upon High-Fat Diet

To further investigate the in vivo role of Abcg1 adipocytes, adipose tissue Abcg1 was silenced by local delivery of lentiviral particles encoding either control shRNAs (L-Ctrl) or Abcg1 shRNAs (L-Abcg1) in epididymal adipose tissue. Injected C57BL/6 mice were maintained on a high-fat diet according to the experimental design presented in Fig. 5A. Expression of Abcg1 in adipocytes in adipose tissue from mice fed a high-fat diet (week 4) was visualized by immunofluorescence (Fig. 5B). Injection of lentiviral particles expressing GFP alone into epididymal adipose tissue confirmed that this strategy was efficient in targeting adipocytes from adipose tissue (W4; Fig. 5C and D). In an independent control experiment, a marked decrease of Abcg1 staining was observed in an epididymal fat pad injected with L-Abcg1 4 weeks following injection compared with an epididymal fat pad injected with L-Ctrl (Fig. 5E), thus validating the KD of Abcg1 in adipose tissue. Such reduced Abcg1 expression in L-Abcg1 fat pads was highly reproducible when tested in multiple tissue samples treated simultaneously under identical experimental conditions. Quantification of adipose tissue Abcg1 mRNA indicated that a significant reduction of Abcg1 expression (−45%; P < 0.05; Fig. 5F) was observed in injected fat pads of L-Abcg1 mice, which mostly reflected the specific silencing of Abcg1 expression in adipocytes (−47%; P < 0.0005) and to a lesser degree in ATMs; the expression of Abcg1 in those cells was approximately threefold less abundant than in adipocytes (Fig. 5G). No effect was observed in endothelial cells isolated from adipose tissue (Fig. 5G) or in other organs such as intestine (Fig. 5H) or liver (Fig. 5I).

Figure 5

KD of adipose tissue Abcg1 following shRNA lentiviral local delivery. A: Scheme of the experimental procedure. B: Abcg1 expression in epididymal adipose tissue from a C57BL/6 mouse was visualized by fluorescence microscopy (magnification, ×63). Arrows indicate Abcg1 expression (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 epididymal adipose 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, ×400). E: Visualization of Abcg1 (red) by fluorescence microscopy (magnification, ×63) in epididymal fat pads following the local injection in the epididymal adipose tissue of lentiviral particles encoding either an shRNA control or an shRNA inhibiting mouse Abcg1 expression. Nuclei were counterstained with DAPI (blue). Quantification of Abcg1 mRNA levels in adipose tissue (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 in L-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 lentiviral particles encoding either a shRNA inhibiting mouse Abcg1 expression (L-Abcg1) or an shRNA control (L-Ctrl). G: Quantification of Abcg1 mRNA levels in adipocytes (mean cycle threshold: 27.95 in L-Ctrl), adipose tissue macrophages (ATMs) (mean cycle threshold: 31.01 in L-Ctrl), and endothelial 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 mean ± SEM (n = 11 mice per group). *P < 0.05 and ***P < 0.0005 vs. L-Ctrl.

Figure 5

KD of adipose tissue Abcg1 following shRNA lentiviral local delivery. A: Scheme of the experimental procedure. B: Abcg1 expression in epididymal adipose tissue from a C57BL/6 mouse was visualized by fluorescence microscopy (magnification, ×63). Arrows indicate Abcg1 expression (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 epididymal adipose 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, ×400). E: Visualization of Abcg1 (red) by fluorescence microscopy (magnification, ×63) in epididymal fat pads following the local injection in the epididymal adipose tissue of lentiviral particles encoding either an shRNA control or an shRNA inhibiting mouse Abcg1 expression. Nuclei were counterstained with DAPI (blue). Quantification of Abcg1 mRNA levels in adipose tissue (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 in L-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 lentiviral particles encoding either a shRNA inhibiting mouse Abcg1 expression (L-Abcg1) or an shRNA control (L-Ctrl). G: Quantification of Abcg1 mRNA levels in adipocytes (mean cycle threshold: 27.95 in L-Ctrl), adipose tissue macrophages (ATMs) (mean cycle threshold: 31.01 in L-Ctrl), and endothelial 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 mean ± SEM (n = 11 mice per group). *P < 0.05 and ***P < 0.0005 vs. L-Ctrl.

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All mice were maintained on a high-fat diet for 4 weeks following injection, and weight gain was evaluated in the 2 groups. Mice injected with L-Abcg1 gained less weight than mice injected with L-Ctrl (−24%; P < 0.05; Fig. 6A) and had decreased epididymal fat mass (−26%; P < 0.05; Fig. 6B), suggesting less fat stored in adipose tissue. Indeed, mean adipocyte diameter was significantly smaller than that in epididymal adipose tissue from L-Ctrl mice (Fig. 6C). In agreement, 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 locomotor activity (Fig. 6F and G) indicated that reduced weight gain in locally injected mice did not result from an overt alteration of energy balance. Additional analysis of metabolic parameters in L-Abcg1 and L-Ctrl mice is presented in Supplementary Table 1. The genes with downregulated expression in stable Abcg1 SKD mouse adipocytes generated in culture were strikingly affected in Abcg1-silenced fat pads, namely, Pparγ, Perilipin, Fabp4, Cd36, Hsl, and C/ebpα (Fig. 6H–M), whereas that of C/ebpβ or Fas was not altered (Fig. 6N and O). Expression of inflammatory and insulin resistance genes in epididymal adipose tissue from L-Ctrl and L-Abcg1 mice is shown in Supplementary Fig. 2.

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) in epididymal adipose tissue were measured 4 weeks after the day of the injection (n = 10 mice per group). E: Food intake was measured in C57BL/6 mice fed a high-fat diet (60% fat) 4 weeks after the local injection of lentiviral particles encoding either an shRNA inhibiting mouse Abcg1 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 mean ± SEM. *P < 0.05, **P < 0.005, and ***P < 0.0005 vs. L-Ctrl.

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) in epididymal adipose tissue were measured 4 weeks after the day of the injection (n = 10 mice per group). E: Food intake was measured in C57BL/6 mice fed a high-fat diet (60% fat) 4 weeks after the local injection of lentiviral particles encoding either an shRNA inhibiting mouse Abcg1 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 mean ± SEM. *P < 0.05, **P < 0.005, and ***P < 0.0005 vs. L-Ctrl.

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

Higher Expression of ABCG1 in Human Adipose Tissue Is Associated With Increased Fat Mass and Corpulence

We demonstrated that adipocyte Abcg1 contributes to TG storage and adiposity and hypothesized that elevated expression of ABCG1 might be associated with increased fat mass in obese subjects. To evaluate this hypothesis, we took the opportunity to examine functional ABCG1 SNPs and their association with adipose tissue gene expression and body composition in obese individuals. Two frequent ABCG1 SNPs (rs1378577 and rs1893590) located in the human ABCG1 gene promoter were genotyped in a population of 1,320 middle-aged morbidly obese patients (BMI >40 kg/m2; mean BMI 45.5 ± 0.04 kg/m2). The relative allele frequencies for both ABCG1 SNPs (rs1893590, −204A/C: −0.73/0.27 and rs1378577, −134T/G: 0.78/0.22) were similar to those observed in the REGRESS cohort (4). As previously described (4), in vitro analysis of ABCG1 promoter activity according to ABCG1 haplotypes confirmed that the frequent AT haplotype was associated with higher transcriptional activity than the rare CG haplotype (Fig. 7A). Analysis of ABCG1 expression in biopsies of adipose tissue isolated from a subset of obese patients displaying either the AT or CG haplotypes revealed that mRNA levels of ABCG1 were 27% (P < 0.05) more elevated in adipose tissues from patients carrying the AT haplotype relative to those carrying the CG haplotype (Fig. 7B). It is notable that ABCG1 expression in primary human preadipocytes was significantly induced upon differentiation into adipocytes (Supplementary Fig. 3A).

Figure 7

Increased adipose tissue ABCG1 expression and increased fat mass and obesity in obese individuals carrying the AT haplotype. A: Human ABCG1 promoter activity according to the CG and AT haplotypes. HepG2 cells were transiently transfected with a construct containing the proximal 1056 bp of the human promoter with either the −204A/−134T (AT) haplotype or the −204C/−134G (CG) haplotype. Luciferase activity is expressed in relative luciferase units (RLU) after normalization for β-galactosidase activity. Values are means ± SEMs of 5 independent experiments performed in triplicate. *P < 0.0005. B–F: Quantification of mRNA levels isolated in adipose tissue biopsies from 10 morbidly obese women carrying either the AT or the CG haplotype. G: Correlation between adipocyte diameter and ABCG1 mRNA levels in adipose tissue from morbidly obese women (n = 20). Fat mass (H) and fat-free mass (I) in obese individuals carrying either the AT (n = 102) or the CG haplotype (n = 22). Data are shown as mean ± SEM. *P < 0.05 vs. CG haplotype, adjusted for age and sex. Association of the rs1378577 (−134T/G) and rs1893590 (−204A/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 ± 0.002 kg/m2). K: Amount of −204A/−134T (AT) haplotypes relative to BMI in obese individuals. AT/AT = 2. The effect of each SNP on BMI was analyzed by linear regression in an additive, 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 ± SEM. *P < 0.05.

Figure 7

Increased adipose tissue ABCG1 expression and increased fat mass and obesity in obese individuals carrying the AT haplotype. A: Human ABCG1 promoter activity according to the CG and AT haplotypes. HepG2 cells were transiently transfected with a construct containing the proximal 1056 bp of the human promoter with either the −204A/−134T (AT) haplotype or the −204C/−134G (CG) haplotype. Luciferase activity is expressed in relative luciferase units (RLU) after normalization for β-galactosidase activity. Values are means ± SEMs of 5 independent experiments performed in triplicate. *P < 0.0005. B–F: Quantification of mRNA levels isolated in adipose tissue biopsies from 10 morbidly obese women carrying either the AT or the CG haplotype. G: Correlation between adipocyte diameter and ABCG1 mRNA levels in adipose tissue from morbidly obese women (n = 20). Fat mass (H) and fat-free mass (I) in obese individuals carrying either the AT (n = 102) or the CG haplotype (n = 22). Data are shown as mean ± SEM. *P < 0.05 vs. CG haplotype, adjusted for age and sex. Association of the rs1378577 (−134T/G) and rs1893590 (−204A/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 ± 0.002 kg/m2). K: Amount of −204A/−134T (AT) haplotypes relative to BMI in obese individuals. AT/AT = 2. The effect of each SNP on BMI was analyzed by linear regression in an additive, 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 ± SEM. *P < 0.05.

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In agreement with our data in Abcg1-deficient adipocytes, levels of mRNAs coding for genes involved in adipocyte differentiation (PPARγ, CD36, PLIN1) were increased in adipose tissue from obese patients carrying the AT haplotype compared with those carrying the CG haplotype (Fig. 7C–E). By comparison, those of LPL were unchanged (Fig. 7F). Adipose tissue mRNA levels of macrophage markers were not different between patients carrying the AT haplotype and those carrying the GC haplotype (Supplementary Fig. 3B–D), suggesting that the number of macrophages present in adipose tissue was similar.

ABCG1 expression in adipose tissue from obese patients was positively correlated with the adipocyte diameter (r = 0.51; P = 0.023; Fig. 7G). Importantly, obese individuals carrying the functional AT haplotype displayed significantly higher fat mass on dual-energy X-ray absorptiometry, together with smaller fat-free mass (P < 0.05; Fig. 7H and I), than those carrying the CG haplotype.

Moreover, the two ABCG1 SNPs were significantly associated with BMI; individuals carrying the −204AA or the-134TT genotype displayed the highest BMI (P = 0.0034 and P = 0.011, respectively) (Fig. 7J). Haplotype analysis confirmed that the AT haplotype (−204A/−134T) was significantly associated with BMI (P = 0.006); moreover, BMI increased in parallel with an increase in the amount of the AT haplotype (Fig. 7K). Association of both ABCG1 SNPs with BMI was still observed after adjustment for diabetes or HOMA index (Supplementary Table 2). ABCG1 SNPs rs1378577 and rs1893590 were not associated with HOMA index after adjustment or not with BMI (data not shown). In addition, obese individuals carrying the −134TT 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 the −204AA genotype (rs1893590, the most frequent genotype). Of note, adiponectin and C-reactive protein concentrations were significantly associated with both ABCG1 SNPs in this population (Supplementary Fig. 3). However no significant effect of ABCG1 SNPs on plasma lipid and apo concentrations was observed (Supplementary Table 3). Thus elevated adipose expression of ABCG1 in obese individuals carrying the AT haplotype is linked with increased fat cell size, fat mass, and obesity and for the first time links ABCG1 genotype to obesity in humans.

Association of the ABCG1 genotype with obesity was replicated in two independent populations with distinct grades of obesity: severely obese (35 < BMI < 40 kg/m2; mean BMI 39.2 ± 8.5 kg/m2) and diabetic obese subjects (30 < BMI < 35 kg/m2; mean BMI 30.9 ± 4.9 kg/m2). Genotyping of both ABCG1 SNPs (rs1378577 and rs1893590) in 216 type 2 diabetic obese subjects (30 < BMI < 35 kg/m2; mean BMI 30.9 ± 4.9 kg/m2) from the Diabetes Atorvastatin Lipid Intervention (DALI) Study (27) revealed that both ABCG1 SNPs were significantly associated with waist-to-hip ratio (Supplementary Fig. 4A and B); individuals carrying the AT haplotype displayed a significantly higher waist-to-hip ratio (P < 0.005) than those carrying the CG haplotype (Supplementary Fig. 4C), confirming the deleterious role of the AT haplotype in obesity. Finally, genotyping of the ABCG1 SNP rs1378577 in an independent population of 595 severely obese subjects (35 < BMI < 40 kg/m2; mean BMI 39.2 ± 8.5 kg/m2) (28) confirmed the association of the ABCG1 genotype with both BMI and fat mass (Supplementary Fig. 4D and E).

In this study, we unraveled an unexpected role for Abcg1 in the control of lipid homeostasis in adipocytes. Through a mechanism involving modulation of both LPL activity and regulation of Pparγ, Abcg1 seems to be a critical component in TG storage and adiposity. The central role of Abcg1 in adipocytes is further strengthened following the targeted silencing of its expression locally in adipose tissue in mice fed a high-fat diet, which led to a rapid reduction in adiposity and weight gain. Consistent with these findings, ABCG1 is associated with adiposity, fat mass, and obesity in obese individuals.

These results may initially seem to conflict with respect to the widely described role of Abcg1 in cellular free cholesterol efflux, where its inhibition leads to an increase rather than a decrease in intracellular lipid accumulation (2). In this way, Abcg1 can protect the cell from the accumulation of free cholesterol, which is toxic to cell survival. In agreement with such a role for Abcg1, we demonstrated that silencing of Abcg1 expression in differentiated 3T3-L1 adipocytes was accompanied by a significant reduction in free cholesterol efflux to HDL, thereby indicating that this mechanism is also operative in fat cells. However, in contrast to free cholesterol in the plasma membrane, the large amount of free cholesterol associated with lipid droplets, which is closely proportional to TG storage in adipocytes and fat cell size, is not mobilized for efflux (29). Mobilization of free cholesterol in lipid droplets therefore provides an alternative pathway for the protection of adipocytes from toxicity. This study supports the notion that although the role of Abcg1 in cellular free cholesterol efflux mechanisms is crucial in maintaining tissue lipid homeostasis in a cholesterol-rich environment (2), Abcg1 contributes to cellular lipid (mostly TGs) accumulation and storage in high-fat metabolic states (4,5). Thus, Abcg1 deficiency (3032) as well as expression of human ABCG1 in mice (2,33) fed an atherogenic diet enriched in cholesterol highlights the role of Abcg1 in protecting tissues, especially the lung, from lipid accumulation without any apparent changes in adipose tissue mass or adiposity. By contrast, and consistent with the mechanism described in the current study, Abcg1 KO mice fed a high-fat diet devoid of cholesterol did not accumulate lipids in tissues but rather exhibited reduced adipose tissue formation and were protected against diet-induced obesity (5).

Our data support a mechanism by which Abcg1 promotes TG storage through the requirement of bioactive LPL (4); addition of exogenous LPL totally rescued the impaired TG storage observed in Abcg1-deficient adipocytes. Our data led us to propose a mechanism by which Abcg1-mediated export of SM contributes to sustaining LPL activity and TG storage in adipocytes. Indeed, LPL hydrolyzes triacylglycerol-rich lipoproteins, allowing the subsequent release and uptake of fatty acids for intracellular TG synthesis. LPL is expressed during the early stages of adipocyte differentiation, when cell–cell contact occurs (34), and adipocyte-derived LPL was reported to be a key determinant for efficient TG storage and adipocyte hypertrophy (8). In agreement with this mechanism, silencing of Abcg1 expression in 3T3-L1 adipocytes was accompanied by marked reduction in TG storage—an effect observed in the early stages of adipocyte differentiation. Beyond its role in TG hydrolysis, LPL also was reported to facilitate lipoprotein uptake and thereby contribute to cellular cholesterol accumulation through this mechanism (14,35). Consistent with such a role for LPL, intracellular concentrations of free cholesterol were markedly reduced in Abcg1 KD adipocytes, a finding in agreement with the reduced lipid droplet size observed in Abcg1 SKD adipocytes. The role of ABCG1 in adiposity and obesity through its action on adipocyte LPL activity is supported by studies of human subjects indicating that adipose tissue LPL activity is increased in obesity (36). Furthermore, several variants in the LPL gene have been associated with obesity (37,38) and ob/ob mice with a specific Lpl deficiency in adipose tissue have displayed reduced weight and fat mass compared with control mice (39).

Alteration of Pparγ and Pparγ-targeted gene expression likely results from the lesser abundance of intracellular cholesterol and fatty acid derivatives delivered by LPL hydrolysis upon silencing of Abcg1 expression in adipocytes. Fatty acid derivatives are ligands for Pparγ activation, and Pparγ expression in adipocytes was reported to be induced by its own activators and/or ligands, such as fatty acids (4042), and cholesterol derivatives through activation of the liver X receptor (43). In agreement with this mechanism, some fatty acid derivatives, whose delivery into cells is mediated by LPL, are activators of Pparγ (41). Interestingly, overexpression of a dominant negative mutant of Pparγ that lacks the 16 COOH− terminal amino acids in 3T3-L1 adipocytes led to a reduction in the rate of free fatty acid uptake, TG storage, and adipocyte size and, more interestingly, to a decrease in the expression of the perilipin, Fabp4, Cd36, Hsl, and C/ebpα genes (44)—a phenotype similar to that observed in the current study when Abcg1 expression was silenced in those cells. However, the modest reduction of adipocyte diameter observed in Abcg1 KD epididymal adipose tissue, in comparison with the more pronounced decrease of the tissue weight, leads us to suggest that the decreased Pparγ expression could also alter adipocyte differentiation and adipocyte cell number in vivo.

The participation of Pparγ in the overall mechanism by which Abcg1 contributes to adiposity and obesity is supported by the observation that the adipose-specific Pparγ knockout mice displayed diminished weight gain and were protected against high-fat diet–induced obesity (45). Taken together, our findings indicate that modulation of LPL activity by Abcg1 in adipocytes might act as an intracellular signaling pathway that controls adipocyte growth through activation of Pparγ and contributes to the formation of fat mass and the development of obesity in humans. Moreover, we reported that a higher expression of ABCG1 in adipose tissue from obese individuals carrying the AT haplotype was associated with increased PPARγ, adiposity, fat mass, and BMI. However, although targeted deletion of Pparγ in murine adipose tissue led to impaired growth of adipose tissue, it must be kept in mind that those mice also exhibited deleterious metabolic consequences such as lipid accumulation in liver and in muscle and potentially insulin resistance (45,46).

In a previous study, Buchman et al. (5) first linked Abcg1 to obesity in a mouse model in which Abcg1 was knocked out in the whole body. Thus, ablation of Abcg1 in Abcg1−/− mice reduces adipose cell size and hepatic steatosis and protects against diet-induced obesity. Although the mechanisms underlying those effects were not elucidated in this earlier study, Buchman et al. proposed that resistance of Abcg1−/− mice to diet-induced obesity likely resulted in increased energy expenditure compared with Abcg1+/+ animals. In addition, a slight reduction of food intake in Abcg1−/− mice also was reported by Buchman et al.; this could contribute to the protective effect of Abcg1 deficiency. Our findings indicate that the contribution of Abcg1 in adiposity and weight gain results from the critical role of adipocyte Abcg1 in adipose tissue, as indicated by the targeted silencing of Abcg1 expression by RNA interference locally in adipose tissue in mice fed a high-fat diet. Moreover, energy expenditure as well as food intake was not altered upon inhibition of Abcg1 expression in adipose tissue, which further reinforces the specific contribution of adipose tissue Abcg1 in those effects.

Although silencing of Abcg1 expression was restricted to adipose tissue, we observed that the injection of lentiviral particles locally in adipose tissue not only targets adipocytes but also equally targets macrophages present in this tissue. Although Abcg1 seems to be expressed more in adipocytes than in adipose tissue macrophages under our experimental conditions, this point may be critical because Abcg1 promotes LPL-mediated lipid accumulation in macrophages in a TG-rich environment (4). However, a recent study reported that macrophage Lpl does not contribute to adiposity and weight gain (47). Epididymal fat mass, lipid droplet size, and gene expression levels in adipose tissue, as well as body weight, were not altered in macrophage Lpl knockout mice compared with control mice.

Although further investigations are required to study the impact of adipocyte Abcg1 deficiency in adipose tissue, our findings nonetheless support the contention that ABCG1 might represent an interesting pharmacological target in obesity, notably by reducing fat mass growth and weight gain in obese patients or in individuals prone to developing morbid obesity.

See accompanying article, p. 689.

Acknowledgments. The authors are indebted to the patients for their cooperation. The INSERM U872 team thanks Assistance-Publique Hôpitaux de Paris, Programme Hospitalier de recherche clinique (PHRC 1996 and 2002), for supporting the genetic DNA bank on obesity.

Funding. INSERMhttp://dx.doi.org/10.13039/501100001677 and UPMC (Parinov program) provided generous support for these studies. M.O., E.F.V., and A.S. have received a Research Fellowship from the French Ministry of Research and Technology. W.P. has received a junior research fellowship from the French Embassy in Thailand. W.L.G. has received a PNRC award from INSERM. The ethics committee (Comité Protection des personnes no. 1 Hôtel-Dieu) provided the ethics agreements. This work was supported by the Fondation de Francehttp://dx.doi.org/10.13039/501100004431 (to P.L., T.H., and W.L.G.) and by the French National Agency through the national program “Investissements d’avenir”http://dx.doi.org/10.13039/501100001665 (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 and a target gene for treating obesity. No other potential conflicts of interest relevant to 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. All authors wrote the manuscript. W.L.G. conceived, designed, and supervised the study. W.L.G. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

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Supplementary data