Role of adipocyte-derived lipoprotein lipase in adipocyte hypertrophy
© Gonzales and Orlando; licensee BioMed Central Ltd. 2007
Received: 30 May 2007
Accepted: 30 October 2007
Published: 30 October 2007
A major portion of available fatty acids for adipocyte uptake is derived from lipoprotein lipase (LPL)-mediated hydrolysis of circulating lipoprotein particles. In vivo studies aimed at identifying the precise role of adipocyte-derived LPL in fat storage function of adipose tissue have been unable to provide conclusive evidence due to compensatory mechanisms that activate endogenous fatty acid synthesis. To address this gap in knowledge, we have measured the effect of reducing adipocyte LPL expression on intracellular lipid accumulation using a well-established cultured model of adipocyte differentiation.
siRNA specific for mouse LPL was transfected into 3T3-L1 adipocytes. Expression of LPL was measured by quantitative real-time PCR and cell surface-associated LPL enzymatic activity was measured by colorimetric detection following substrate (p-nitrophenyl butyrate) hydrolysis. Apolipoprotein CII and CIII expression ratios were also measured by qRT-PCR. Intracellular lipid accumulation was quantified by Nile Red staining.
During differentiation of 3T3-L1 pre-adipocytes, LPL mRNA expression increases 6-fold resulting in a 2-fold increase in cell surface-associated LPL enzymatic activity. Parallel to this increase in LPL expression, we found that intracellular lipids increased ~10-fold demonstrating a direct correlation between adipocyte-derived LPL expression and lipid storage. We next reduced LPL expression in adipocytes using siRNA transfections to directly quantify the contributions of adipocyte-derived LPL to lipid storage, This treatment reduced LPL mRNA expression and cell surface-associated LPL enzymatic activity to ~50% of non-treated controls while intracellular lipid levels were reduced by 80%. Exogenous addition of purified LPL (to restore extracellular lipolytic activity) or palmitate (as a source of free fatty acids) to siRNA-treated cells restored intracellular lipid levels to those measured for non-treated controls. We also found that adipocytes express apolipoprotein CII and CIII and, in addition, the apoCII/apoCIII ratio remains largely unchanged in cells with reduced LPL expression.
We provide evidence that adipocyte-derived LPL is required for efficient fatty acid uptake and storage, and that adipocytes express their own source of apoCII and apoCIII for regulating extracellular LPL activity. These findings demonstrate that adipocytes are capable of producing the necessary enzymatic components and co-factors for efficient lipid storage independent of vascular sources.
The adipocyte plays a crucial role in metabolic regulation, serving as a storage depot for fatty acids and as an endocrine cell to manage energy utilization and feeding behavior [1, 2]. The mass of adipose tissue is maintained by a well-controlled balance of cell proliferation (hyperplasia) and increase in fat cell size (hypertrophy). Increases in adipocyte hypertrophy result from the uptake and assimilation of extracellular fatty acids into cytosolic triacylglycerol-rich lipid droplets. The primary sources of these extracellular fatty acids are those that are 1) associated with circulating albumin or 2) hydrolyzed from triacylglycerol-rich lipoprotein particles such as chylomicrons or very low density lipoproteins (VLDL). Since chylomicrons are short-lived fatty acid carriers present only during the post-prandial period, it is accepted that VLDL particles represent the major source of circulating fatty acids in the form of triacylglycerols. Triacylglycerols are the major component of VLDL and provide a concentrated source of esterified fatty acids for peripheral tissues.
The availability of VLDL-derived fatty acids for energy needs and adipocyte storage is dependent upon hydrolytic activity of lipoprotein lipase (LPL) [3–6], and to a lesser extent, endothelial lipase [7, 8]. To date, two models have been proposed describing how LPL mediates lipolysis of circulating lipoproteins to release fatty acids for adipocyte uptake . In the first model, LPL originating in adipose or muscle tissue is transported across the endothelial barrier where it remains tethered to the lumenal surface through its affinity for heparan sulfate proteoglycans (HSPG). In this position, LPL can mediate release of fatty acids from circulating lipoproteins for diffusion-based transport to the adipocyte surface or newly released fatty acids are transported through their association with circulating albumin. Alternatively, largely intact lipoproteins can cross the capillary endothelial barrier by either transcytosis [9–11] or through leaky cell-cell junctions. Lipolysis itself is known to cause capillary leakage and can contribute to transendothelial transport [12, 13].
Lipoprotein particles that are transported out of the vasculature represent a significant source of fatty acids for adipocyte storage. The release of fatty acids from these particles is likely to be dependent on adipocyte-derived LPL activity. One of our recent studies provides evidence that HSPG found on the surface of adipocytes are essential for lipid accumulation in adipocytes . Although the exact role played by these HSPG toward the uptake of fatty acids remains to be identified; we have hypothesized that HSPG serve as reaction centers for LPL-mediated triacylglycerol hydrolysis. Cell surface HSPG are known to provide high affinity binding sites for both LPL [15, 16] and apoE-enriched lipoproteins [17–19]. In this manner, HSPG can serve to co-localize enzyme and substrate, creating a focal reaction center to bring about the release of fatty acids in proximity of the adipocyte cell surface. Uptake of these liberated fatty acids can then be accomplished by fatty acid transport proteins, such as FAT/CD36  or FATP1 , which are known to be expressed by adipocytes [22–24]. A significant advantage of this mechanism is that lipoproteins, such as VLDL, can provide a concentrated supply of fatty acids to adipocytes for uptake as opposed to simply relying on free diffusion across the endothelial barrier.
An essential element necessary for this mechanism to be effective is adipocyte-derived LPL activity [25–27]. Studies examining the effects of LPL gene ablation in mice have attempted to define the role of adipocyte-derived LPL in fat storage function of adipose tissue . Homozygous LPL deficiency was found to be lethal in mice; however, muscle-specific expression of LPL in an LPL -/- background was able to rescue the lethal phenotype . In these crossbred mice, LPL expression is absent in adipose, yet little or no change was noted in adipose tissue mass. It was later shown that normal lipid storage was preserved in these mice by an increase in endogenous fatty acid synthesis . Thus, interpreting the exact role of adipose-derived LPL toward lipid storage in these studies was limited due to compensatory mechanisms that activate endogenous fatty acid synthesis . Because endogenous fatty acid synthesis in adipose is able to counterbalance the loss of adipose-derived LPL activity in vivo, animal studies have not been able to shed light on the quantitative role played by this source of LPL in lipid uptake and storage. In order to directly address this gap in knowledge, we have measured the effect of reducing adipocyte LPL expression on intracellular lipid accumulation using a well-established cultured model of adipocyte differentiation.
Cell culture and adipocyte differentiation
3T3-L1 cells were obtained from American Type Culture Collection (Manassas, VA) and grown in Dulbecco's modified Eagle's medium (DMEM) (Invitrogen, Carlsbad, CA) supplemented with 10% (v/v) fetal calf serum (Irvine Scientific, Santa Ana, CA), 1 mM sodium pyruvate, 0.1 mM non-essential amino acids, 2 mM L-glutamine, 100 μg/ml streptomycin sulfate, and 100 units/ml penicillin. Cells were cultured at 37°C with 10% CO2 and passaged twice weekly. To differentiate 3T3-L1 cells into adipocytes, cells were incubated with 250 nM dexamethasone, 450 μM 3-isobutyl-1-methylxanthine, and 167 nM insulin for 2 days, followed by 167 nM insulin for an additional 3 days.
siRNA Transfection and PCR analysis
Purified Silencer® pre-designed small interfering RNA (siRNA) directed toward mouse lipoprotein lipase (LPL) was synthesized by Ambion Inc. (Austin, TX). The target sequence for the siLPL is on exon 2, with a sense sequence of GCAAAUUUGCCCUAAGGACtt and an antisense sequence of GUCCUUAGGGCAAAUUUGCtt. Cells were transfected with siRNA (100 nM) using DharmaFECT™ 3 transfection reagent according to the manufacturer's specific protocol for 3T3-L1 cells (Dharmacon, Inc., Lafayette, CO). Total RNA was purified from cells using RNeasy (Qiagen, Valencia, CA) and converted to cDNA using TaqMan Reverse Transcriptase (Applied Biosystems, Branchburg, NJ). LPL, leptin, apoCII, apoCIII, and β-actin expression levels were measured by quantitative Real Time PCR analysis (qRT-PCR) of cDNA samples. Primers specific for LPL [GenBank:NM_008509] were designed to amplify 231 base pairs flanking intron 4. Primer sequences for LPL were: upstream, CTGCTGGCGTAGCAGGAAGT; downstream, GCTGGAAAGTGCCTCCATTG. Primers specific for leptin [GenBank:NM_008493] were designed to amplify 213 base pairs flanking intron 2. Primer sequences for leptin were: upstream, TGACACCAAAACCCTCATCA; downstream, TCATTGGCTATCTGCAGCAC. Primers specific for apoCII [GenBank: NM_009695] were designed to amplify 147 base pairs flanking intron 2. Primer sequences for apoCII were: upstream, GCAGGGCTCCCTCTTAAGTT; downstream, AAAATGCCTGCGTAAGTGCT. Primers specific for apoCIII [GenBank: NM_023114] were designed to amplify 99 base pairs flanking intron 3. Primer sequences for apoCIII were: upstream, GGCTGGATGGACAATCACTT; downstream, TGGTTGGTCCTCAGGGTTAG. Primers specific for β-actin [GenBank:NM_007393] were designed to amplify 287 base pairs flanking intron 3. Primer sequences for β-actin were: upstream, CCTGAACCCTAAGGCCAACC; downstream, CAGCTGTGGTGGTGAAGCTG. qRT-PCR was performed using ABsolute QPCR SYBR Green Mix (Fisher Scientific, Atlanta, GA) with the following cycling parameters: 1 cycle, 95°C, 15 min; 40 cycles, 95°C, 15 sec, 60°C, 1 min. β-actin mRNA levels were measured in parallel with identical cycling conditions and used to normalize values obtained for LPL, leptin, apoCII, and apoCIII expression. Relative quantitation was performed using the comparative CT method. For this analysis, the amount of target message (LPL, leptin, apoCII, or apoCIII in siRNA-treated adipocytes) was normalized to the internal reference (β-actin) and compared to the calibrator (LPL, leptin, apoCII, or apoCIII in untreated adipocytes).
For straight RT-PCR reactions of leptin, apoCII, apoCIII and β-actin, isolated mRNA was reverse transcribed as described above and cDNA amplification was performed using TaKaRa Ex Taq™ (Fisher Scientific, Atlanta, GA) with the following cycling parameters: 1 cycle, 95°C, 2 min; 40 cycles, 95°C, 30 sec, 60°C, 30 sec, 72 °C 1 min. Amplified products were separated on 3% agarose gels and stained with Gel Star® (Cambrex, Rockland, ME).
LPL enzymatic activity
Cells, grown in 6-well plates, were rinsed twice with phosphate buffered saline, pH 7 (PBS), and incubated with 200 μg/ml heparin (from porcine intestinal mucosa, 180 USP units/mg, Sigma, St. Louis, MO) prepared in PBS for 15 min at 25°C. Following incubation, the heparin solution was centrifuged (15,000 × g, 5 min, 4°C) to remove cell debris and mixed with an equal volume of 2 mM p-nitrophenyl butyrate (PNPB). Absorbance at 405 nm was recorded following a 10 min incubation using a Genesys UV Spectrophotometer. LPL activity is reported as the amount of p-nitrophenol product formed over the 10 min incubation. The molar extinction coefficient of p-nitrophenol at 405 nm is 11,000 M-1cm-1.
Palmitate (sodium salt), purified bovine LPL, and fatty acid-free albumin were purchased from Sigma. Palmitate-albumin complexes were prepared as follows. Palmitate was dissolved in 3% albumin (wt/v) to a final concentration of 10 mM in 1X Hank's Balanced Salt Solution. The mixture was slowly heated to 50°C and stirred overnight. The mixture was then cooled to 37°C, stirred for an additional 3 hours and sterile filtered through a 0.2 μm syringe filter. Protein was quantified using the BCA™ Protein Assay Kit (Pierce, Rockford, IL).
Nile Red staining and quantitation
Cells were rinsed twice with PBS, pH 7, and trypsinized with phenol red-free 0.05% Trypsin-EDTA (Invitrogen Corporation, Grand Island, NY) for 5 min at 37°C. Cells were fixed in suspension with 0.5% paraformaldehyde (v/v) in PBS for 10 min at 25°C. A stock solution of 1 mg/ml Nile Red (Sigma, St. Louis, MO) was prepared in acetone and added to the cell preparation to a final concentration of 5 μg/ml. Cells were incubated at 25°C for 30 min in the dark. Flow cytometry was performed using a FACScan flow cytometer (λexcitation 488 nm; λemission 585 nm). Nile Red fluorescence was quantitated using a Shimadzu RF-1501 spectrofluorophotometer (λexcitation 488 nm; λemission 560 nm).
All experimental protocols were done in triplicate. All error bars represent standard deviation of triplicate points.
LPL expression and cell-surface enzymatic activity of 3T3-L1 preadipocytes and differentiated adipocytes
LPL expression by adipocytes is necessary for intracellular lipid accumulation
Exogenous addition of LPL or palmitate restores intracellular fat storage in siRNA treated adipocytes
ApoCII and apoCIII expression in adipocytes
Since apoCII and apoCIII have opposite effects on LPL activity, it is generally accepted that the state of LPL activity depends on the ratio of apoCII/apoCIII expression. A high ratio of apoCII/apoCIII promotes LPL activity, whereas a low apoCII/apoCIII ratio inhibits LPL function. Since this ratio can significantly affect extracellular LPL activity and in turn fatty acid uptake and storage by adipocytes, we determined apoCII and apoCIII expression levels in adipocytes treated with or without siRNA for LPL. Although expression of both regulators was largely unaffected by LPL-siRNA treatment, we did find a small but reproducible effect on their expression as a result of diminished LPL expression (Fig. 6B). ApoCII expression increased by ~20%, while apoCIII expression decreased by ~20%.
The generation of fatty acids from dietary sources and their uptake by cells are essential processes for efficient energy metabolism and storage. A general role for LPL in these processes has been confirmed from LPL gene ablation studies in mice which results in a lethal phenotype . Naturally occurring LPL gene mutations have also been reported in the human population that lead to severe hypertriglyceridemia . However, in light of these studies which have established a critical systemic role for LPL in overall fatty acid handling, investigations carried out to evaluate the direct contributions of adipocyte-specific LPL toward lipid storage have not yielded conclusive evidence because of compensatory increases in endogenous fatty acid synthesis . The resulting limitations of published in vivo studies are likely due to hormonal regulation responsible for adipose mass preservation . To circumvent this problem and obtain direct quantitative evidence of the role of adipocyte-derived LPL in lipid storage, we used a well established in vitro culture model of adipocyte differentiation . Differentiation of this culture model to full adipocytes resulted in a 6-fold increase in LPL mRNA expression. This increase in LPL mRNA translated into a 2-fold increase in cell surface-associated LPL enzymatic activity and, importantly, ~10-fold increase in intracellular lipid storage. Using siRNA specific for LPL, we were able to reduce LPL expression by ≥ 50% during the course of adipocyte differentiation. This reduction in adipocyte LPL expression resulted in a parallel decrease in lipid storage of ~80%, which was completely reversed by the exogenous addition of either purified LPL or palmitate. These results clearly establish that adipocyte-derived LPL plays a quantitatively important role in intracellular lipid accumulation.
It is now well established that LPL requires apoCII as a co-factor for efficient triacylglycerol hydrolysis [39, 40]. This fact is highlighted by the observation that mutations in the apoCII gene lead to abnormally high levels of circulating triacylglycerols . Following its synthesis and secretion by hepatocytes, apoCII associates with circulating lipoproteins and stimulates LPL-mediated hydrolysis once in contact with LPL that is associated with the vascular wall. ApoCII is a multidomain protein with binding sites for both LPL and lipid components of circulating lipoprotein particles [42, 43]. The dual binding domains of apoCII are thought to anchor LPL to the particle surface while permitting substrate access to its active site . In contrast, apoCIII is a potent inhibitor of LPL activity [44, 45]. Because of this, the expression levels, or ratio, of apoCII to apoCIII can be a major influence on extracellular LPL activity . ApoCII and apoCIII are known to carry out their functions in the circulation largely while bound to lipoproteins. In the extravascular environment, we questioned if other non-hepatic sources, such as adipocytes, contribute to expression of apoCII and apoCIII and in doing so provide their own source of LPL regulators to manage extravascular lipid hydrolysis. This is indeed the case for differentiated adipocytes as RT-PCR analysis revealed expression of both apoCII and apoCIII. We also examined the apoCII/apoCIII expression ratio by qRT-PCR analysis and found potential regulation of this ratio in adipocytes that appears to be influenced by LPL expression levels; decreased LPL expression following siRNA treatment resulted in a small but reproducible increase in apoCII and decrease in apoCIII expression. This change in the apoCII/apoCIII ratio may reflect a compensatory mechanism whereby reduced levels of LPL are counterbalanced by an increase in the apoCII/apoCIII ratio. Increasing the apoCII/apoCIII ratio may serve to ensure more effective activation of the remaining pool of extracellular LPL.
We thank Mark B. Carter (University of New Mexico Cytometry research group) for his technical help with the flow cytometric analysis. This work was supported by an American Heart Association Grant-in-aid no. 0555647Z (to R.A.O.). The images presented in this study were generated in the Fluorescence Microscopy Facility which is supported by NCRR 1 S10 RR14668, NSF MCB9982161, NCRR P20 RR11830, NCI R24 CA88339, NCRR S10 RR19287, NCRR S10 RR016918, the University of New Mexico Health Sciences Center, and the University of New Mexico Cancer Center.
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