PNPLA2 influences secretion of triglyceride-rich lipoproteins by human hepatoma cells

Abstract
Patatin-like phospholipase domain containing proteins (PNPLAs) are involved in triglyceride hydrolysis and lipid-droplet homeostasis in mice, but the physiological significance of the PNPLAs for triglyceride metabolism in human hepatocytes is unclear. Here we investigate the roles of PNPLA2, PNPLA3 and PNPLA4 in triglyceride metabolism of human Huh7 and HepG2 hepatoma cells using gene-specific inhibition methods. siRNA inhibition of PNPLA3 or PNPLA4 is not associated with changes in triglyceride hydrolysis, secretion of triglyceride-rich lipoproteins (TRLs) or triglyceride accumulation. However, PNPLA2 siRNA inhibition, both in the absence or presence of oleate- containing medium, or treatment with the PNPLA2 inhibitor Atglistatin, reduced intracellular triglyceride hydrolysis and decreased TRL secretion. In contrast, PNPLA2 inhibition showed no effects on lipid-droplet homeostasis, which is the primary physiological function of PNPLA2 in non- hepatic tissues. Moreover, confocal microscopy analysis found no clear evidence for the localization of PNPLA2 around lipid-droplets. However, significant co-localization of PNPLA2 with the endoplasmic reticulum marker PDI was found in HepG2 and Huh7 cells with Rcoloc values of 0.61±0.06 and 0.81±0.05, respectively. In conclusion, PNPLA2 influences TRL secretion but is not involved in lipid-droplet homeostasis in human hepatoma cells, a physiological role that is quite distinct from the metabolic function of PNPLA2 in non-hepatic tissues.

Introduction
Hepatic steatosis, the most common form of liver disease, is a precursor to fibrosis, cirrhosis and liver cancer (1) and is tightly linked to obesity, type 2 diabetes mellitus and cardiovascular disease (2,3). The hepatic triglyceride content defines steatosis and it is generally assumed that disturbances in hepatic triglyceride metabolism are an integral part of the etiology of hepatic steatosis (4). A potential cause of steatosis is defects in the hydrolysis of triglycerides stored in hepatocytes, a metabolic process that is not only of importance for hepatic lipid-droplet homeostasis, but also influences the secretion of triglyceride-rich lipoproteins (TRLs) by the liver (5,6). Unfortunately, little is known about the lipase(s) responsible for cellular triglyceride hydrolysis in human hepatocytes.In 2004, three groups independently identified a member of the patatin-like phospholipase domain containing protein (PNPLA) family, PNPLA2 (also known as adipose triglyceride lipase, ATGL), as the enzyme responsible for the catalysis of the initial step of triglyceride lipolysis in adipocytes, converting triglycerides to diacylglycerols (7-9). PNPLA2 knockout (KO) mice exhibited increased adipose tissue mass (9) as a consequence of impaired rates of adipocyte triglyceride hydrolysis (10). Because of its high expression and prominent role in adipose metabolism, subsequent research has largely focused on the role of PNPLA2 in adipose tissue (11). However, PNPLA2 is also expressed, albeit at lower levels, in non-adipose tissues such as heart and liver. Cardiac muscle triglyceride content was markedly increased in PNPLA2-KO mice (9,10). In contrast, only partial reductions in triglyceride-hydrolase activity and modest increases in triglyceride content were observed in the liver of PNPLA2-KO mice (9,10). Subsequent studies evaluated the role of hepatic PNPLA2 in triglyceride metabolism using methods to modulate PNPLA2-activity specifically in the liver.

Adenovirus- mediated knockdown of hepatic PNPLA2 with short hairpin RNA resulted in steatosis in mice and decreased triglyceride hydrolysis in primary murine hepatocyte cultures (12). In addition, mice with liver-specific inactivation of Pnpla2 developed a progressive form of hepatic steatosis (13). Moreover, liver overexpression of PNPLA2 in obese mice reduced hepatic steatosis (14,15). Overall, these studies suggest that PNPLA2 plays an important role in hepatic lipid metabolism, at least in mouse models. To the best of our knowledge, no study has verified the physiological role of PNPLA2 in human hepatocytes.PNPLA2 mRNA levels are low in human liver as compared to those of PNPLA3, an enzyme with in vitro triglyceride-hydrolase activity (7) and an established genetic risk-factor for hepatic steatosis and its sequelae (16). Moreover, human hepatocytes express PNPLA4, another PNPLA family-member with in vitro triglyceride-hydrolase activity that is not expressed in rodents (7). Here, we used siRNA inhibition methods to determine the physiological functions of PNPLA2, PNPLA3 and PNPLA4 in triglyceride metabolism in the human hepatoma Huh7 and HepG2 cell-lines. We found that PNPLA2, but not PNPLA3 or PNPLA4, influences secretion of TRLs, but is not involved in lipid-droplet homeostasis of human hepatoma cells. Confocal microscopy analysis found no clear evidence for the localization of PNPLA2 around lipid-droplets, while significant co-localization of PNPLA2 with the endoplasmic reticulum marker PDI was observed.Huh7 and HepG2 cell lines were purchased from the Japanese Cancer Research Resources Bank and ATCC, respectively. Cells were maintained at 37oC and 5% CO2 and cultured in low glucose Dulbecco’s Modified Eagle’s Medium containing L-glutamine, supplemented with 10% fetal bovine serum (FBS), 50 U/mL penicillin and 50 μg/mL streptomycin (PEST).

The medium was changed twice/week and cells were subcloned weekly. The universal mycoplasma detection kit (ATCC) was used at three-monthly intervals to screen for mycoplasma infection of the cells.For the siRNA silencing experiments, cells of 60-70% confluency were transfected with gene-specific or control siRNA oligonucleotides (Supplemental Table S2) using Lipofectamine RNAiMax (Thermo Fisher Scientific) as transfection reagent. For each gene, two different siRNA probes were used for all experiments. The medium was replaced after 24 hours with culture medium containing 14C-glycerol (PerkinElmer, see section below entitled “Assays”); 24 hours later the cell-medium was collected and the cells were harvested and analyzed.For the Atglistatin experiments, cells were first incubated for 24 hours with cell-medium, alternatively transfected with gene-specific or control siRNA oligonucleotides as described above. After 24 hours the medium was replaced with culture medium containing 14C-glycerol supplemented with 50 μM Atglistatin (Cayman Chemical) or DMSO; 24 hours later the cells were harvested and analyzed.For the oleic acid experiments, cells were first transfected with siRNA oligonucleotides as described above. The medium was replaced after 24 hours with culture medium containing 14C-glycerol (PerkinElmer), 5% FBS, 50 U/mL PEST and 0.2 mM oleic acid conjugated to fatty acid free BSA (Sigma). The cells were harvested after 24-hour incubation.Whole-cell protein lysates were extracted using RIPA buffer containing EDTA (Thermo Fisher Scientific), protease inhibitors (Halt Protease Inhibitor Cocktail, Thermo Fisher Scientific) and phosphatase inhibitors (Phosphatase Inhibitor Cocktail 2, Sigma). Ten μg of protein were loaded on10% mini-Protean TGX gels (Biorad) and transferred to activated PVDF membrane (Biorad), blocked in 3% BSA and incubated overnight with 1:1000 diluted PNPLA2 (R&D) or PNPLA3 (Aviva) primary antibody followed by incubation with HRP-conjugated secondary antibodies against sheep and mouse (Biorad). Proteins were visualized using Pierce ECL Western Blotting Substrate (Thermo Fisher Scientific) and analyzed with a LAS-1000 Imager (Fujifilm). The same blots were subsequently incubated with 1:3500 diluted -actin antibody (Biorad), visualized and analyzed as a loading control. See Supplemental Table S1 for more details regarding the antibodies used in this study.

RNA was isolated with E.Z.N.A. Total RNA Kit 1 (Omega Bio-tek) and cDNA was synthesized with a High-Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific). Taqman assays (Supplemental Table S2) and AB7500 sequence detection system (Applied Biosystems) were used for relative mRNA expression analysis. Relative expression was assessed using the comparative ΔΔCT method and adjusted for the endogenous control RPLP0. The results of the gene expressions studies were verified using the relative standard curve method according to the Applied Biosystems guidelines.The triglyceride quantification colorimetric/fluorometric kit (Biovision) was used to measure the cellular triglyceride content. Cell protein concentration was quantified using the Pierce BCA assay (Thermo Fisher Scientific). Triglyceride secretion was quantified in cell-culture medium following 24- hour incubation of the cells with cell-culture medium containing 14C-glycerol (PerkinElmer) at a final concentration of 2.85 mCi/mL (19). The 14C-labelled lipids were extracted from the cell-culture medium and separated by TLC using Hexane:Diethylether:Acetic acid (80:10:1). The radioactivity associated with the 14C-triglycerides was quantified using a scintillation counter. The apolipoprotein B (APOB) in the cell-culture medium was quantified by ELISA (ALerCHEK).Triglyceride-hydrolase activity was measured essentially as described (18). Briefly, 24 hours after siRNA transfection of the hepatoma cells the cell-culture medium was switched to cell-culturemedium containing 1.0 μCi of 14C-palmitic acid (PerkinElmer). After 24 hours incubation, the cell- culture medium was changed to cell-culture medium containing 5 μM Triascin C (Sigma), an inhibitor of the long-chain fatty acyl CoA synthetase isoproteins 1, 3, 4 and 5.

Cells were harvested at the indicated time intervals and cellular 14C-triglyceride radioactivity was quantified as described above.Hepatoma cells were cultured overnight on glass coverslips (Marienfeld) and fixed in 4% PFA (Histolab) for 20 minutes at room temperature (RT). For the lipid-droplet analysis, the hepatoma cells were stained with either 1:100 diluted BODIPY493/503 (Molecular Probes, Life Technologies) or HCS LipidTOX Red (Invitrogen) and mounted with Vectashield mounting medium containing DAPI (Vector Laboratories). Selected sample regions were imaged with Leica SP5 confocal microscope, equipped with a 63×1.4 lens and diode and argon lasers. Image stacks consisted of a Z-stack of 15 to 20 optical slices taken at 0.15-0.30 μm intervals to enhance the spatial signal allocation. Lipid-droplet area was quantified in every experiment by analyzing 10 randomly chosen fields, each of 30–50 cells. The number and area of the lipid-droplets in each field were quantified using the ImageJ cell counter and Particle Analysis Plugin (Fiji), while the number of cells was calculated manually. The average lipid-droplet area/cell and lipid-droplet number/cell were calculated by dividing the overall lipid- droplet area and the total number of lipid-droplets by the number of cells in the same field, respectively.For the co-localization experiments, hepatoma cells were cultured on glass coverslips and fixed either with PFA (for PDI) or with methanol (for PLIN2), followed by permeabilization and blocking with 10% goat serum (Vector Laboratories). The cells were subsequently incubated for one hour with 1:50 diluted PNPLA2 monoclonal antibody (Novus Biologicals) conjugated with Alexa Fluor 488 using the Zenon Alexa Fluor 488 Mouse IgG2b Labeling Kit (Thermo Fisher Scientific) at RT, followed by overnight incubation with either 1:100 diluted PDI (Enzo) or 1:100 diluted PLIN2 (R&D Systems) monoclonal antibodies. The coverslips were then incubated with goat anti-mouse Alex Fluor 594 antibody, mounted with Vectashield (Vector Laboratories) medium containing 4′,6-diamidino-2- phenylindole (DAPI) and stored at 4 oC. Image stacks were obtained as described above. The averageco-localization was calculated using the JACoP plug in of the ImageJ program in 10-15 images obtained for each condition. The relationships between the lipid-droplets and PNPLA2 and PLIN2 were analyzed in the absence or presence of 0.4 mM oleic acid conjugated to fatty acid free BSA in cell culture medium for 8 hours, followed by fixation, permeabilization, blocking and overnight incubation with 1:100 diluted PNPLA2 monoclonal antibody. The cells were subsequently incubated with goat anti-mouse Alex Fluor 488 antibody and 1:1000 HCS LipidTOX Red (Invitrogen) for 1 hour, mounted and analyzed as previously described.Statistical analysis and graphic design was conducted using GraphPad Prism v 6.01 software. Data is presented as mean ± SD unless stated otherwise. Statistical significance was evaluated by Student’s t- test with a threshold value of p = 0.05. Bonferroni correction was used to adjust the threshold p-values for multiple testing in the gene-expression studies.

Results
Functional analysis of the roles of PNPLA2, PNPLA3 and PNPLA4 in hepatic triglyceride metabolism was performed in human hepatoma Huh7 and HepG2 cell-lines using transient transfection techniques with gene-specific siRNA probes. For each gene, two different siRNA probes were used for all experiments. Comparable results were obtained for both probes for each of the three PNPLA genes and most data are presented as the average results of the two gene-specific siRNA probes, but occasionally results of the two probes are shown separately. As shown in Figure 1A, substantial reductions in mRNA levels of PNPLA2, PNPLA3 and PNPLA4 are achieved, in the absence of compensatory increases in the expression of non-targeted PNPLA genes. Corresponding decreases in PNPLA2 and PNPLA3 protein concentrations after respective siRNA inhibition, evaluated using Western Blot analysis, are shown in Figures 1B and 1C. Unfortunately, no suitable PNPLA4 antibody for Western Blot analysis could be identified. No consistent changes in mRNA levels are noted for genes involved in triglyceride synthesis, lipid-droplet or TRL metabolism after gene-specific siRNA inhibition in Huh7 or HepG2 cells (Figure 1D and Supplemental Figure S1).We were unable to quantify the low triglyceride-hydrolase activity in Huh7 or HepG2 cells using an established technique for the quantification of triglyceride hydrolysis in adipocytes in vitro (17). We therefore developed a semi-quantitative in vivo triglyceride-hydrolysis assay using the approach of Hobbs and coworkers (18). In this assay, cellular triglycerides are first labelled with 14C-palmitate for 24-hours, followed by inhibition of de novo triglyceride synthesis with Triascin C and subsequent quantification of cellular 14C-labeled triglycerides after 4 and 8 hours. We found that PNPLA2 siRNA inhibition decreases triglyceride hydrolysis at both time points in Huh7 and HepG2 cells (Figure 2A). A similar effect was observed when 3H-oleate instead of 14C-palmitate was used for the analysis of triglyceride hydrolysis in Huh7 cell (Supplemental Figure S2). We subsequently quantified triglyceride hydrolysis using only the 8-hour time point and found that PNPLA2 inhibition, but notgene-specific inhibition of PNPLA3 or PNPLA4, leads to significant reductions of cellular triglyceride hydrolysis (Figure 2B).

Finally, we found no evidence that gene-specific inhibition of either PNPLA2, PNPLA3 or PNPLA4 influences the cellular uptake of 14C-palmitate or the incorporation of 14C- palmitate in triglycerides (a measure of triglyceride synthesis) in the hepatoma cell-lines (Supplemental Figure S3), two factors that can in theory influence the triglyceride hydrolysis assay. Overall, these results indicate that PNPLA2, but not PNPLA3 or PNPLA4, influences cellular triglyceride hydrolysis in human hepatoma cells. In agreement with the data from PNPLA2-KO mice (9,10) we find that PNPLA2 siRNA inhibition decreases the overall cellular triglyceride hydrolysis by only around 50%, despite effective inhibition of PNPLA2. This suggests that a substantial proportion of the triglyceride hydrolase activity in human hepatoma cells cannot be accounted for by PNPLA2 and must instead be attributed to the action of other triglyceride hydrolase enzyme(s).The effects of gene-specific inhibition of PNPLA2, PNPLA3 or PNPLA4 on the secretion of TRLs and cellular triglyceride accumulation by Huh7 and HepG2 cells were analyzed using methods described previously (19). Significant reductions in the secretion of triglycerides and APOB are observed in Huh7 and HepG2 cells following siRNA inhibition of PNPLA2 while no effects of PNPLA3 or PNPLA4 inhibition on TRL-secretion are found (Figure 2C). However, no effects of gene-specific inhibition of PNPLA2, PNPLA3 or PNPLA4 on cellular triglyceride content are observed in the hepatoma cell-lines (Figure 2D). We performed confocal microscopy studies to verify this observation. We did not identify significant effects of PNPLA2 inhibition on lipid-droplet area or the overall lipid-droplet size distribution in either Huh7 (Figures 3A, 3B and 3C) or HepG2 cells (Figures 3D, 3E and 3F). Thus, PNPLA2 inhibition reduces TRL-secretion, while no evidence is found for a significant role of PNPLA2 in lipid-droplet homeostasis in human hepatoma Huh7 and HepG2 cells.

Incubation of Huh7 and HepG2 cells with 0.2 mM oleate-supplemented cell-medium increases, as expected, the secretion of triglyceride and APOB, and the cellular triglyceride content of the hepatoma cells as compared to cells incubated with cell-medium without oleate-supplement (Figures 4A and 4B). Nevertheless, Huh7 and HepG2 cells cultured in 0.2 mM oleate-supplemented cell-medium showed similar effects of PNPLA2 siRNA inhibition on PNPLA2 mRNA levels (Figure 4C), triglyceride secretion (Figure 4D) and cellular triglycerides content (Figure 4E) as compared to human hepatoma cells cultured without oleate-supplement (Figures 1A, 2C and 2D).Atglistatin is an inhibitor of PNPLA2 in mouse cells/tissues (20), but a recent study noted that Atglistatin is not an effective inhibitor of PNPLA2 in human Simpson-Golabi-Behmel-Syndrome adipocytes (21). Nevertheless, we found that incubation of human hepatoma Huh7 cells with 50 μM Atglistatin resulted in 30-40% reductions of total cellular triglyceride-hydrolysis activity (Supplemental Figure S4). We subsequently compared the effects of PNPLA2 siRNA inhibition and PNPLA2 inhibition with 50 μM Atglistatin on triglyceride-hydrolysis activity in both Huh7 and HepG2 cells (Figure 5A). It was found that both PNPLA2 siRNA inhibition and PNPLA2 inhibition with Atglistatin are associated with partial reductions in overall triglyceride-hydrolysis activity in both hepatoma cells lines. However, PNPLA2 siRNA inhibition is a slightly more effective method to reduce triglyceride-hydrolase activity in Huh7 and HepG2 cells as compared to PNPLA2 inhibition with Atglistatin (Figure 5A). We subsequently evaluated the effect of a combination of Atglistatin and PNPLA2 siRNA inhibition on triglyceride-hydrolase activity in the hepatoma cells. As shown in Figure 5A, no evidence was found that a combination of the two PNPLA2 inhibition methods leads to a greater inhibition of total cellular triglyceride-hydrolase activity as compared to PNPLA2 siRNA inhibition alone.

These observations indicate that both PNPLA2 inhibition techniques are able to inhibit PNPLA2-related triglyceride-hydrolase activity in the two human hepatoma cell-lines.No consistent changes in mRNA levels for genes involved in triglyceride synthesis, lipid-droplet or TRL metabolism after Atglistatin inhibition or a combination of Atglistatin and PNPLA2 siRNA inhibition were observed (Figure 5B and Supplemental Figure S5). As shown in Figure 5C, the twoPNPLA2 inhibition techniques reduce the secretion of triglycerides by the Huh7 and HepG2 cells by approximately 45% and 35%, respectively. Again, no additive effects of the combined PNPLA2 siRNA and Atglistatin inhibition on triglyceride secretion is observed when compared with PNPLA2 siRNA inhibition alone. The effects of the two inhibition methods on APOB secretion show a similar pattern (Figure 5C): a modest decrease of APOB secretion following Atglistatin inhibition, but no significant differences in the reduction of APOB secretion when the PNPLA2 siRNA inhibition and the combined PNPLA2 siRNA and Atglistatin inhibitions are compared in the two hepatoma cell-lines. In contrast, the two PNPLA2 inhibition strategies do not influence cellular triglyceride levels (Figure 5D). Confocal microscopy confirmed that neither Atglistatin inhibition alone nor the combination of PNPLA2 siRNA and Atglistatin inhibition influences lipid-droplet area or size distribution in Huh7 cells (Supplemental Figure S5). Overall, we find comparable qualitative effects of PNPLA2 siRNA and Atglistatin inhibition on TRL-secretion, while no effects of these inhibition strategies on lipid droplet homeostasis are detectable in either Huh7 or HepG2 cells.Subcellular localization studies in 3T3-L1 adipocytes demonstrated that PNPLA2 is located predominantly around lipid-droplets (4,22). Here, we employed confocal microscopy to determine the subcellular localization of PNPLA2 in human hepatoma Huh7 and HepG2 cells. PNPLA2 was visualized using a monoclonal antibody that showed specificity for PNPLA2 and no cross-reactivity with other proteins in Huh7 and HepG2 cells (Supplemental Figure S).

Immunofluorescence staining of PNPLA2 showed a patchy/mottled distribution of PNPLA2 in the cytoplasm of the hepatoma cells (Figure ). However, no evidence was found for the enrichment of PNPLA2 around lipid droplets of hepatoma cells cultured in either 10% FBS-supplemented or oleate-supplemented cell-medium (Figure ). Nevertheless, in agreement with previous reports (23,24), immunofluorescence staining of perilipin2 (PLIN2) showed a preferential localization of PLIN2 around lipid-droplets in human hepatoma cells cultured in 10% FBS medium and an enhanced accumulation of PLIN2 around lipid- droplets in human hepatoma cells cultured in oleate-supplemented cell-medium (Supplemental Figure S7). Overall, these subcellular localization studies provided no clear evidence for the preferentiallocalization of PNPLA2 around lipid-droplets. Of note, we did not study the subcellular localization of PLIN1, while we found that the PLIN1 mRNA levels were below 2% of the PLIN2 mRNA concentration in Huh7 and HepG2 cells. Furthermore, we are not aware that anyone has been able to detect PLIN1 protein in human hepatoma cell-lines.The subcellular localization studies of PNPLA2 and PLIN2 shown in Figure and Online Figure S demonstrated that essentially all PNPLA2 and a substantial fraction of PLIN2 is found in the cytoplasma of the hepatoma cells cultured in 10% FBS cell-medium. In view of the lipophilic nature of PNPLA2 and the preferential association PLIN2 with lipid-droplets, we hypothesized that PNPLA2 and PLIN2 are both associated with small lipid-droplets present in the cytoplasm of the hepatoma cells. These lipid-droplets, because of their small size, are not detectable by conventional light- microscopy as distinct lipid-droplets, but are expected to show a diffuse staining pattern when performing confocal microscopy, explaining the patchy/mottled appearance of the PNPLA2-stained hepatoma cells shown in Figure . Indeed, PNPLA2 and PLIN2 exhibited comparable patchy/mottled distribution patterns in the cytoplasm of both hepatoma cell lines (Figure 7A). Moreover, partial co- localization was observed between PNPLA2 and PLIN2, with Rcoloc values (mean±SD) of 0.49±0.07 (n=10) and 0.±0.02 (n=13) for Huh7 and HepG2 cells, respectively. Note that these calculations are based on the overall distribution of PNPLA2 and PLIN2 in the hepatoma cells. Since a proportion of PLIN2, but not PNPLA2, is preferentially localized around large lipid droplets (Figure and Supplemental Figure S7), it can be expected that the Rcoloc values are higher for the cytoplasmic regions devoid of large lipid-droplets.The involvement of PNPLA2 in the regulation of TRL secretion suggests that PNPLA2 resides in a subcellular compartment that is part of, or in close proximity to the TRL-synthetic machinery present in the endoplasmic reticulum (ER). We therefore analyzed the co-localization of PNPLA2 with PDI, asubunit of the microsomal triglyceride-transfer complex (MTTP) involved in the assembly of TRLs and commonly used as an ER marker. As shown in Figure 7B, we found that both PNPLA2 as well as PDI exhibited a patchy/mottled pattern in the cytoplasm of the hepatoma cells. Moreover, a substantial co-localization of PNPLA2 with PDI was observed, with Rcoloc values (mean±SD) of 0.1±0.0 (n=10) and 0.81±0.05 (n=11) for Huh7 and HepG2 cells, respectively.

Discussion
This study evaluated the biological significance of PNPLA2, PNPLA3 and PNPLA4 for hepatic triglyceride metabolism in the human hepatoma Huh7 and HepG2 cell-lines, with particular emphasis on the roles of these proteins in triglyceride hydrolysis, TRL-secretion and cellular triglyceride accumulation. Gene-specific inhibition of PNPLA3 or PNPLA4 was not associated with changes in triglyceride hydrolysis, TRL-secretion or cellular triglyceride accumulation, indicating that PNPLA3 and PNPLA4 do not play major roles in these aspects of hepatic lipid metabolism. In contrast, PNPLA2 siRNA inhibition, both in the absence or presence of oleate-containing medium, or treatment with the PNPLA2 inhibitor Atglistatin, reduced intracellular triglyceride hydrolysis and decreased TRL secretion. However, no effects of PNPLA2 inhibition were observed on cellular triglyceride concentration or the number and size of the lipid-droplets. Subcellular localization studies found no clear evidence for associations between PNPLA2 and large lipid-droplets, while co-localization studies revealed that PNPLA2 is primarily found in close proximity to the TRL-synthetic machinery present in the ER of human hepatoma cells. Together, these observations suggest that the triglyceride- hydrolase activity of PNPLA2 contributes to the regulation of TRL-secretion by human hepatocytes, but does not influence lipid-droplet homeostasis of human hepatoma cells.

Previous studies demonstrated that PNPLA2 plays a major role in the hydrolysis of triglycerides in adipocytes and other non-hepatic tissues (11). Correspondingly, PNPLA2 is preferentially localized around lipid-droplets in 3T3-L1 adipocytes (9,22). However, the current study provides two lines of evidence against a role of PNPLA2 in the hydrolysis of triglycerides stored in lipid-droplets of human hepatoma cells. First, PNPLA2 siRNA inhibition, both in the absence or presence of oleate-containing medium, or treatment with the PNPLA2 inhibitor Atglistatin, was found to have no effect on cellular triglyceride concentrations or lipid-droplet size and distribution of the hepatoma cells. Second, co- localization studies found no clear evidence for the localization of PNPLA2 on the surface of the large lipid-droplets visualized by confocal microscopy. These observations largely exclude a role for PNPLA2 in the hydrolysis of triglyceride molecules in large lipid-droplets in human hepatoma cells and thereby indicate that yet unidentified enzyme(s) are involved in this process. To the best of our knowledge, this is the first report demonstrating that PNPLA2 influences TRL secretion by human hepatoma cells. There is long-standing evidence that approximately half of the fatty acids used for TRL synthesis are derived from triglyceride molecules stored in hepatocytes (5,6). It is therefore conceivable that the triglyceride-hydrolase activity of PNPLA2 is responsible for mobilizing fatty acids from stored triglycerides, which are subsequently used for TRL synthesis, but the nature of these temporary triglyceride storage-sites is elusive. Our co-localization studies in human hepatoma cells demonstrated that PNPLA2 is predominantly localized near the ER-marker PDI. We also observed co-localization of PNPLA2 with PLIN2, a protein that is best known as a lipid-droplet associated protein. However, only a minor fraction of PLIN2 was actually associated with the large lipid-droplets in our co-localization studies. Most of the PLIN2 protein was located in the cytoplasm of the hepatoma cells and the cytoplasmic PLIN2 was largely responsible for the observed co- localization with PNPLA2. Together, these co-localization studies suggest that PNPLA2 and cytoplasmic PLIN2 are both located near the ER.

It is tempting to speculate that PNPLA2 and cytoplasmic PLIN2 are constituents of lipid-droplets that are loosely associated with the ER, but are too small to be visualized as distinct lipid-droplets by confocal microscopy. Small lipid-droplets, designated initial lipid-droplets (iLDs), were recently identified as a specific lipid-droplet subclass that is not detectable by conventional light-microscopy and is metabolically distinct from so-called expanding lipid droplets (eLDs), which are large lipid-droplets that can be visualized with confocal microscopy (4,25). An attractive hypothesis is that the small iLDs in human hepatoma cells act as a temporarily storage site for triglycerides synthesized in the ER and subsequently acquire/activate PNPLA2 for the generation of fatty acids for TRL formation, but more detailed studies are required to substantiate this hypothesis. The strength of our study is that various aspects of hepatic triglyceride metabolism are analyzed using complementary methods. For example, two different hepatoma cell-lines (HepG2 and Huh7) are employed, different siRNA inhibitors and Atglistatin inhibition are used, TRL secretion is analyzed by two different parameters (APOB and 14C-triglyceride measurements) and cellular triglyceride accumulation is estimated by triglyceride concentration measurement and confocal microscopy analysis. A limitation of our study is the use of human hepatoma cell-lines given their secretion of relatively dense, triglyceride-poor particles as compared to the VLDL lipoproteins secreted in vivo by mammalian liver (26), although it is generally thought that the overall mechanism of triglyceride metabolism and TRL secretion is retained in these cells (reviewed in 27).

In summary, this study demonstrates that PNPLA2 inhibition of human hepatoma Huh7 and HepG2 cells is associated with decreased triglyceride hydrolysis and reduced TRL secretion, while no changes in the cellular triglyceride content is observed. In contrast, the primary physiological function of PNPLA2 in non-hepatic tissues, such as adipose tissue, is the hydrolysis of triglycerides stored in lipid-droplets. This suggests that PNPLA2 plays a metabolic role in human hepatoma cells that is quite distinct from its function in non-hepatic tissues.