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Am J Physiol Cell Physiol. Oct 1, 2012; 303(7): C728–C742.
Published online Jun 27, 2012. doi:  10.1152/ajpcell.00448.2011
PMCID: PMC3469596

Direct interaction of Plin2 with lipids on the surface of lipid droplets: a live cell FRET analysis

Abstract

Despite increasing awareness of the health risks associated with excess lipid storage in cells and tissues, knowledge of events governing lipid exchange at the surface of lipid droplets remains unclear. To address this issue, fluorescence resonance energy transfer (FRET) was performed to examine live cell interactions of Plin2 with lipids involved in maintaining lipid droplet structure and function. FRET efficiencies (E) between CFP-labeled Plin2 and fluorescently labeled phosphatidylcholine, sphingomyelin, stearic acid, and cholesterol were quantitated on a pixel-by-pixel basis to generate FRET image maps that specified areas with high E (>60%) in lipid droplets. The mean E and the distance R between the probes indicated a high yield of energy transfer and demonstrated molecular distances on the order of 44–57 Å, in keeping with direct molecular contact. In contrast, FRET between CFP-Plin2 and Nile red was not detected, indicating that the CFP-Plin2/Nile red interaction was beyond FRET proximity (>100 Å). An examination of the effect of Plin2 on cellular metabolism revealed that triacylglycerol, fatty acid, and cholesteryl ester content increased while diacylglycerol remained constant in CFP-Plin2-overexpressing cells. Total phospholipids also increased, reflecting increased phosphatidylcholine and sphingomyelin. Consistent with these results, expression levels of enzymes involved in triacylglycerol, cholesteryl ester, and phospholipid synthesis were significantly upregulated in CFP-Plin2-expressing cells while those associated with lipolysis either decreased or were unaffected. Taken together, these data show for the first time that Plin2 interacts directly with lipids on the surface of lipid droplets and influences levels of key enzymes and lipids involved in maintaining lipid droplet structure and function.

Keywords: perilipin, ADRP, cyan fluorescent protein, triacylglycerols, phospholipids

intracellular lipid droplets, composed of a neutral lipid core surrounded by a phospholipid monolayer, are ubiquitously found in every cell type as storage vesicles for lipids. Larger triacylglycerol (TG)-rich lipid droplets (10–200 μm diameter) are present in adipocytes (52, 54) while smaller droplets (1 μm diameter or less) enriched in cholesteryl esters (CE) exist in steroidogenic cells (20). In other cell types such as fibroblasts, a mixed ratio of TG/CE can be found (7, 54). Understanding cell-specific regulation of lipid droplet reserves has become an active area of research, and cell type often defines the proteins associated with lipid droplets. The Plin family of proteins, related through sequence homology and affinity for lipid droplets, includes Plin1 (formerly known as perilipin) and Plin4 (S3–12), found primarily in adipocytes and steroidogenic cells; Plin2 (ADRP) and Plin3 (TIP47), both ubiquitously expressed in all cell types; and Plin5 (also known as OXPAT), a protein found in cells with high energy requirements (myocytes, hepatocytes) (reviewed in Ref. 11). Interestingly, intracellular location of these proteins varies and governs function. Plin2 and Plin1 are constitutively located on the surface of lipid droplets while Plin3, Plin4, and Plin5 can be found in both cytosolic and lipid droplet compartments (17, 52, 71, 72). While a large data set exists for Plin1, less is known regarding the physiological role of Plin2 (and other Plin family members) in maintaining lipid droplet structure and function. The present work was undertaken to examine how Plin2 participates in regulating intracellular lipid metabolism through protein-lipid interactions on the lipid droplet surface.1

Despite the discovery over 20 years ago of Plin2 as an early marker for adipocyte differentiation (43), the physiological relevance of Plin2 in cellular metabolism remains unclear. Initial views considered Plin2 (and Plin1) to act as a protein barrier against lipolysis (50, 52). However, there is currently no evidence to suggest that Plin2 participates in the lipolytic process which involves the actions of Plin1, adipose triglyceride lipase (ATGL), hormone-sensitive lipase (HSL), and comparative gene identification-58 (CGI-58) working in concert to release stored TG from the lipid droplet neutral core (11, 15, 16, 30). Upon hormonal stimulation, HSL and Plin1 are phosphorylated by cAMP-dependent protein kinase A, causing HSL to translocate to the lipid droplet surface and Plin1 to release CGI-58 which then activates ATGL to start the lipolytic process (reviewed in Ref. 75). Based on studies with mice deficient in ATGL and HSL, the primary lipase responsible for TG hydrolysis is ATGL while HSL targets diacylglycerols (DG) (60) and also acts as a cholesterol esterase (33) converting CE to free cholesterol (Chol). In the last step of TG hydrolysis, the final fatty acid (FA) and glycerol are released by monoacylglycerol lipase (MAGL) (2). The following enzymes are also involved in the biosynthesis of TG, CE, and phospholipids: 1) monoacylglycerol acyltransferase (MGAT), involved in the first step of TG synthesis in the monoacylglycerol (MG) pathway to TG (reviewed in Ref. 62); 2) acyl-coenzyme A:diacylglycerol acyltransferase (DGAT) enzymes 1 and 2, involved in the second step of the MG pathway combining a fatty acyl-CoA and DG molecule to form TG (reviewed in Ref. 74); 3) glycerol-3-phosphate acyltransferases (GPAT), existing as four isoforms derived from separate genes that act as the rate-limiting enzyme in the glycerol phosphate or Kennedy pathway to TG synthesis (70); 4) 3-hydroxy-3-methylglutaryl-CoA reductase (HMGR), involved in the rate-limiting step of the mevalonate pathway that produces cholesterol (19); 5) acyl-coenzyme A:cholesterol acyltransferase-1 (ACAT-1), an enzyme that catalyzes the formation of CE from Chol and long-chain fatty-acyl-CoA (22); 6) phosphocholine cytidylyltransferase (CCT), the rate-limiting enzyme to PC synthesis (46); and 7) SM synthase (SMS), existing as two isoforms and the last enzyme in the SM biosynthetic pathway involved in converting PC to SM (24). It should be noted that several of the proteins involved in lipolysis including Plin1, ATL, and HSL (but not Plin2 or Plin3) were recently found localized in rigid, highly organized areas of the lipid droplet phospholipid monolayer, presumably to help facilitate the lipolytic process through protein sequestration (66), much like areas of the plasma membrane where FA/Chol uptake and efflux occur (18, 25, 49, 57). The lack of Plin2 in the lipolytic complex was consistent with the fact that there is no evidence to suggest that Plin2 participates, or can replace Plin1's lipolytic function, even in perilipin-knockout mice where Plin2 replaced Plin1 on the surface of lipid droplets (67). There is, however, some indication that Plin2 can regulate access of ATGL to the lipid droplet surface in different cells (10, 50), but it is unclear whether CGI-58 is involved in the process. These results, along with the fact that Plin2 exhibits high-affinity binding of lipids such as Chol (6, 7, 27), FA (7, 61), and phospholipids (54) gives rise to the hypothesis that Plin2 may play an important role in regulating lipid exchange at the lipid droplet surface to maintain lipid homeostasis. In keeping with this, several studies including the present work have shown that increased expression of Plin2 promotes cellular lipid accumulation (29, 48, 50). Conversely, knockdown of Plin2 in macrophages decreased cellular lipids and lipid droplet size and number (48). In mice deficient in Plin2, liver lipids were reduced without changes in triacylglycerol synthesis, very-low-density lipoprotein secretion, fatty acid uptake, synthesis, or β-oxidation (21). Instead, a twofold increase in microsomal triacylglycerol was observed, suggesting an inability to direct lipid stores from the ER to lipid droplets (21). However, since this Plin2-deficient mouse model showed expression of an active truncate (designated Δ2,3-ADPH) that could target to lipid droplets (59), the full phenotype of Plin2 gene ablation remains to be determined. In the present work, new insights into the role of Plin2 in lipid droplet metabolism were demonstrated by both live cell FRET imaging and an examination of lipid content and protein expression in CFP-Plin2-overexpressing cells. First, CFP-Plin2 was shown to directly interact with several fluorescently labeled lipids on the surface of lipid droplets in living cells. Given the surface proximity and ubiquitous nature of Plin2 in all cell types, along with its high binding affinity for several lipid droplet lipids, these results suggested a role for Plin2 in binding to, and directing lipid exchange across, the lipid droplet surface. Second, levels of key enzymes and lipids involved in maintaining lipid droplet structure and function were influenced by Plin2 overexpression. Protein expression levels of enzymes involved in TG, CE, and phospholipid synthesis were significantly increased while levels of enzymes associated with lipolysis were either decreased or unaffected in CFP-Plin2-overexpressing cells. In summary, the data presented herein provide novel insights into the role of Plin2 in cellular metabolism and suggest that Plin2 influences protein-lipid interactions on the lipid droplet surface to promote lipogenesis.

MATERIALS AND METHODS

Materials.

Lipid standards were purchased from Nu-Chek Prep (Elysian, MN) and Avanti (Alabasta, AL). Silica Gel G and Silica Gel 60 thin layer chromatography (TLC) plates were from Analtech (Newark, DE) and EM Industries (Darmstadt, Germany), respectively. Rabbit polyclonal antiserum to Plin2 was prepared in house as described previously (5). The following antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA): affinity-purified goat polyclonal antibodies raised against mouse ATGL, mouse GPAT, and human MGAT, and rabbit polyclonal antibodies raised against human DGAT, human HSL, and human SMS. Rabbit polyclonal antiserum raised against human CCT was purchased from Epitomics (Burlingame, CA). Affinity-purified rabbit polyclonal antibodies raised against HMGR, ACAT-1, and LDLR were purchased from Biovision Research Products (Milpitas, CA). The rabbit monoclonal antibody against SR-B1 was from Novus Biologicals (Littleton, CO). Rabbit polyclonal antiserum to GFP and the following probes were purchased from Invitrogen Molecular Probes (Eugene, OR): 6-((N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino)-hexanoyl)sphingosyl-phosphocholine (NBD-sphingomyelin); [22-(N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino)-23,24-bisnor-5-cholen-3b-ol] (NBD-cholesterol); 12-N-methyl-(7-nitrobenz-2-oxa-1,3-diazo)aminostearic acid (NBD-stearate); 1-acyl-2-(N-4- nitrobenzo-2-oxa-1,3-diazole)-aminocaproyl phosphatidylcholine (NBD-PC); and 7-diethylamino-3,4-benzophenoxazine-2-one (Nile red). All reagents and solvents used were of the highest grade available and were cell culture tested.

Cells in culture.

Mouse-transfected L cell fibroblasts were cultured at 37°C under 5% CO2 in Higuchi medium (36) containing 10% fetal bovine serum (Sigma, St. Louis, MO). Fluorescence imaging and FRET experiments were performed with cells seeded at a density of 50,000 cells/chamber on Lab-Tek chambered coverglass slides (Nunc, Naperville, IL) and cultured overnight before use. The plasmid expressing CFP-Plin2 was generated by subcloning the complete coding sequence of mouse Plin2 into vector pECFP-N1 from BD Biosciences Clontech (Palo Alto, CA) using unique restriction sites (XhoI, KpnI) introduced by PCR. The CFP-Plin2 expression plasmid was sequenced to verify identity and fidelity and then stably transfected into mouse L cell fibroblasts using Superfect from Qiagen (Valencia, CA) according to the manufacturer's instructions. Twenty-four hours after transfection, cells were placed on medium containing G418 (700 μg/ml of medium) to allow selection of resistant clones. Clones surviving G418 treatment were selected and screened by Western blot analysis and fluorescence imaging to ensure stable expression of CFP-Plin2. Mock transfectants (clones transfected with vector DNA without insert) were generated in parallel with the expression clones and were designated as controls. Because there was no significant difference between untransfected control cells and mock-transfected control cells over the parameters considered in the present work, data for both were combined and designated as controls.

Western blot analysis.

Relative expression of proteins related to lipid storage was determined by Western blot analysis in CFP-Plin2-overexpressing cells versus control cells as described previously (4, 54). Briefly, homogenates samples (5–10 μg protein) from CFP-Plin2-overexpressing cells and control cells were run on tricine gels (12%) and then transferred by electroblotting to 0.45-μm nitrocellulose paper (Sigma). Blots were blocked in 3% gelatin in TBST (10 mM Tris·HCl, pH 8, 100 mM NaCl, 0.05% Tween 20) for 1 h at room temperature, washed 2× with TBST, and incubated overnight at 1:1,000 dilutions in 1% gelatin TBST with primary polyclonal rabbit antibodies against Plin2, GFP (specific for all variations of GFP), GPAT, CCT, SMS, HSL, MGAT, DGAT, ATGL, LDLR, SR-B1, HMGR, or ACAT-1. After, the blot was washed (3×, TBST) and incubated at room temperature for 2 h with alkaline-phosphatase conjugates of either anti-goat or anti-rabbit IgG in 1% gelatin TBST. Then the blot was washed (3×, TBST) and developed with Sigma Fast 5-bromo-4chloro-3-indolyl phosphate/nitro blue tetrazolium tablets (Sigma) according to the manufacturer's protocol. To ensure equal protein loading on the gels, expression levels of the housekeeping gene β-actin was also determined to normalize protein expression. Briefly, the proteins of interest and β-actin were resolved by size on tricine gels as described above, and, after the transfer step, membranes were cut lengthwise into two blots to detect expression levels of the protein of interest or β-actin. Relative protein expression was determined as integrated density values based on densitometric analysis of image files acquired using a single-chip charge-coupled device (CCD) video camera and a computer workstation (IS-500 system from Alpha Innotech, San Leandro, CA) using ImageJ (available from National Institutes of Health, Bethesda, MD; http://rsbweb.nih.gov/ij/download.html).

Live cell colocalization studies.

For colocalization studies, digital images were acquired using a MRC-1024MP Laser Scanning Confocal Imaging System (LSCIS) with LaserSharp software equipped with three PMT for fluorescence detection in separate channels, a 15 mW krypton-argon laser (Kr+/Ar, American Laser, Salt Lake City, UT) with a 5 mW output measured at the microscope stage, a 408 nm violet diode laser (Power Technology, Little Rock, AR), and a Zeiss Axiovert 135 inverted epifluorescence microscope (Zeiss, Thornwood, NY) equipped with a ×63 Zeiss oil Apochromat objective. To determine subcellular localization, CFP-Plin2-overexpressing and control cells grown on chamber slides were rinsed twice with phosphate-buffered saline (PBS) and then incubated in PBS for 30 min at room temperature with Nile red or the following NBD-labeled lipid probes: Chol, stearic acid, PC, and SM. For probe excitation, the MRC-1024MP LSCIS utilized the 408 nm violet diode laser (for CFP) and the 488 nm line (for NBD-labels) or the 568 nm line (for Nile red) from the Kr+/Ar laser to acquire images of the cells by sequential excitation. During image acquisition, cells were exposed to the light source for minimal time periods to minimize photobleaching effects. Fluorescence emission of the CFP label was collected using a HQ470/15 nm filter (Chroma Technology, Bellow Falls, VT). Emission from the NBD-labeled lipids was collected through a HQ530/30 nm filter (Chroma Technology) while the Nile red fluorescence emission was collected through a HQ598/40 nm filter. Image files were analyzed using MetaMorph 7.5 software (Molecular Devices, Sunnyvale, CA). For colocalization experiments, LaserSharp 3.0 was used to identify either the CFP-labeled pixels (arbitrarily placed in the red channel) colocalized with NBD (green channel) or the CFP (green channel) with Nile red signals (red channel). The confocal images from the green and red channels were merged and appeared yellow where superimposition occurred (red and green are additive and yield yellow to orange in RGB color space). Pixel fluorograms were constructed and correlation coefficients generated from the fluorograms were derived from the following equations:

Cred=iRi,colociRiCgreen=iGi,colociGi

where ∑Ri,coloc is the sum of intensities of all red pixels which also have a green component; ∑Ri is the sum of intensities of all red pixels in the image; ΣGi,coloc is the sum of intensities of all green pixels which also have a red component; and ∑Gi is the sum of intensities of all the green pixels in the image.

Live cell FRET imaging.

To determine the molecular association between Plin2 and lipids on the lipid droplet surface, FRET analysis was performed by acceptor photobleaching on CFP-labeled Plin2 and NBD-labeled lipids or Nile red as described previously (44, 56, 73) using CFP (donor)-NBD (acceptor) and CFP (donor)-Nile red (acceptor) as FRET pairs. Cells were processed as for colocalization studies (described above). Qualitatively, FRET from the CFP donor to NBD acceptor molecules was detected upon excitation at 408 nm as the increased donor emission of CFP (through HQ470/15 nm band-pass filter) after photobleaching of the NBD acceptor at 488 nm since excitation overlap of CFP and NBD at 408 nm did not allow adequate separation. Nonlabeled cells were used to set the gain and black level in the CFP channel so that no cellular autofluorescence was detected in the donor CFP channel while maintaining maximum dynamic range with CFP-Plin2-labeled cells. In addition, to check for bleed through from the acceptor channel despite the use of a narrow HQ470/15 nm filter, cells not expressing CFP-Plin2 were labeled with NBD-labeled lipids or Nile red at 2× higher concentrations than were experimentally used with no observed fluorescence in the donor CFP channel upon 408 nm excitation. In the acceptor (NBD or Nile red) channel, gain and black level were set to suppress CFP fluorescence bleed through as determined by cells just before the addition of NBD-labeled lipids or Nile red. After control experiments were completed, FRET analysis was performed as follows: the fluorescence emission of the CFP donor through the HQ470/15 nm filter was recorded using 408 nm excitation before and after photobleaching of the acceptor where the photobleaching process occurred using either the 488 nm (NBD) or 568 nm (Nile red) line of the Kr+/Ar+ laser for a duration of 2–4 min. In addition, pre- and post-bleach images of the acceptor, NBD or Nile red, were acquired at attenuated excitation levels. Duration of photobleaching under high-intensity excitation was optimized to avoid damage to cells. As a control, these same photobleaching conditions were applied to CFP-Plin2-labeled cells without acceptor, which resulted in no measurable changes in the CFP-Plin2 fluorescence in the CFP channel under conditions used for acceptor depletion. FRET efficiency (E) was calculated from the following: E = 1 − (IDA/ID) where IDA is donor fluorescence intensity before acceptor (NBD-label) photobleaching and ID is the donor fluorescence intensity after acceptor photobleaching. An average E value was calculated from CFP fluorescence emission increase after photobleaching in 30–40 lipid droplets from a minimum of 20 cells. The intermolecular distance R between CFP-Plin2 and the NBD- or Nile red-labeled lipid was estimated from the calculated E according to E = 1/[1 − (R/Ro)6] where Ro for the CFP/NBD and CFP/Nile red FRET pair was previously determined as 42 Å and 48 Å, respectively. For the FRET efficiency images, analysis was performed in MetaMorph 7.5 (Molecular Devices). Images were filtered to remove randomized high frequency noise by using a low pass filter using 4 × 4 pixel setting. The filtered images of the donor emission before acceptor photobleaching were subtracted from the image after acceptor photobleaching. The resultant image was divided by the image of donor emission after acceptor photobleaching and multiplied by 100 to show the grayscale FRET efficiencies. A FRET overlay was created from this image and pseudo-colored in order to visualize regions of higher and lower FRET where blue indicated little to no efficiency and red to yellow represented efficiencies >60%.

Lipid analysis.

Lipids were extracted from CFP-Plin2 and control cell homogenates and resolved into individual lipid classes as described earlier (7, 54). In brief, samples extracted with n-hexane:2-propanol 3:2 (vol/vol) were resolved by lipid group into triacylglycerol (TG), cholesteryl esters (CE), cholesterol (Chol), and total phospholipids (PL) using Silica gel G TLC plates developed in petroleum ether-diethyl ether-methanol-acetic acid (90:7:2:0.5, vol/vol/vol/vol). For resolution of diacylglycerols (DG), the following solvent system was used: petroleum ether-diethyl ether-glacial acetic acid (280:120:4 vol/vol/vol/vol). Levels of Chol, DG, TG, CE were determined by the method of Marzo et al. (53). Polar lipids (PL) were eluted from the Silica gel G resin using chloroform:methanol:HCl (100:50:0.375, vol/vol/vol), dried under N2, and resuspended in chloroform. Half of each sample was used for total PL analysis and half was applied to Silica gel 60 TLC plates to resolve the following individual PL: sphingomyelin (SM), phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylinositol (PI), and phosphatidylserine (PS) using chloroform:methanol:water:acetic acid (150:112.5:6:10.5, vol/vol/vol/vol). No separation was observed between phosphatidic acid (PA) and PE. Individual PL spots were visualized (iodine vapor) and eluted from the Silica gel 60 resin using chloroform:methanol:HCl (100:50:0.375, vol/vol/vol) before analysis by the method of Marzo et al. (53). All lipids were identified by comparison to known standards. Protein concentration was determined by the method of Bradford (13) from the dried protein extract residue digested overnight in 0.2 M KOH. Lipids were stored under an atmosphere of N2 to limit oxidation and all glassware was washed with sulfuric acid-chromate before use.

Statistics.

All values are expressed as means ± SE. Statistical analysis was performed using analysis of variance (ANOVA) combined with the Newman-Keuls multiple comparisons test (GraphPad Prism, San Diego, CA). Values with P < 0.05 were considered statistically significant.

RESULTS

CFP-Plin2-expressing cells.

To analyze protein-lipid interactions on the surface of lipid droplets by laser scanning confocal microscopy (LSCM), a CFP-Plin2 expression construct was made by fusing the complete coding sequence of mouse Plin2 in-frame to the COOH terminus of the mammalian expression vector pECFP-N1 using the unique restriction sites XhoI and KpnI (Fig. 1A). The resulting plasmid was stably transfected into mouse L cell fibroblasts, and cells were screened for CFP fluorescence by LSCM. CFP-Plin2 overexpression was also determined by quantitative analysis of multiple Western blots (normalized to the housekeeping gene β-actin) loaded with CFP-Plin2 and control cell homogenates probed against anti-Plin2 (Fig. 1B) or anti-CFP (Fig. 1C) to show a respective 1.5- and 1.6-fold increase in CFP-Plin2 expression (Fig. 1D). The cyan variant of GFP was chosen for the CFP-Plin2 expression construct because CFP forms a strong FRET interaction with commercially available NBD-labeled lipids (Fig. 2A) that target to lipid droplets (6, 7, 27, 54) due to the strong overlap of the emission spectra of the cyan fluorescence (408 nm excitation, 475 nm emission) with the NBD excitation spectra (488 nm excitation, 530 nm emission). In addition, a strong overlap of CFP emission with the excitation of the lipid droplet stain Nile red (568 nm excitation, 598 nm emission) was also indicated (Fig. 2B).

Fig. 1.
Construct design and relative protein expression in CFP- Plin2-overexpressing cells. The CFP-Plin2 expression construct (A), which included the entire coding region of mouse Plin2 cDNA cloned into the unique XhoI and KpnI restriction sites of vector pECFP-N1, ...
Fig. 2.
Spectral overlap between CFP/NBD-probes and CFP/Nile red. The excitation and emission spectra of CFP (lines 1 and 2, respectively) of the CFP/NBD FRET pair (A) are shown superimposed with the respective excitation and emission spectra of NBD (lines 3 ...

Live cell targeting and FRET analysis of CFP-Plin2 with NBD-labeled PC and SM.

Although it has been shown that Plin2 binds PC and SM with high affinity (54), the extent and functional significance of Plin2/phospholipid interactions on the LD surface are not currently known. Therefore, the ability of CFP-Plin2 to interact with NBD-labeled phospholipids was examined by live cell laser scanning confocal microscopy (LSCM) in a series of experiments. First, simultaneous acquisition of confocal images of CFP-Plin2-overexpressing cells incubated with NBD-PC revealed areas of high intensity and overlap in lipid droplets, resulting in yellow-to-orange colocalized signals (Fig. 3A). The extent of colocalization was quantified in pixel fluorograms (Fig. 3B) where correlation coefficients indicated that 99% of the CFP-label (red channel) colocalized with the NBD-label (green channel), while 87% of the NBD stain colocalized with the CFP probe. These results were consistent with NBD-PC targeting mostly to lipid droplets. Similarly, CFP-Plin2-overexpressing cells were incubated with NBD-SM to show initial targeting of the NBD label to the plasma membrane, followed by lipid droplets after 8–10 min (Fig. 4A). Overlap of the CFP-labeled Plin2 with NBD-SM was observed as yellow to red pixels indicating colocalization of probes (Fig. 4A). From the pixel fluorogram (data not shown), 99% of the CFP probe colocalized with the NBD-SM, but only 52% of NBD-SM colocalized with CFP due to NBD-SM targeting to plasma membranes. Taken together, these results were consistent with strong spatial overlap occurring between Plin2 and phospholipids on the surface of lipid droplets, to within 2,200 Å.

Fig. 3.
Colocalization and FRET imaging between CFP-Plin2 and NBD-PC. CFP-Plin2-overexpressing cells were labeled with NBD-PC to determine colocalization and FRET efficiency between the fluorescently labeled Plin2 and lipid. The extent of colocalization (A) was ...
Fig. 4.
Colocalization and FRET imaging between CFP-Plin2 and NBD-labeled SM, Chol, and stearic acid. Colocalization (A, C, and E) and FRET efficiency images (B, D, and F) were generated for CFP-Plin2-overexpressing cells with NBD-SM (A and B), NBD-Chol (C and ...

To obtain higher spatial resolution, live cell FRET was performed as described in materials and methods and as previously described (44, 56, 73). FRET imaging allowed estimations of intermolecular distances between Plin2 and lipid molecules to within 10–100 Å. Experimentally, FRET was measured as the increase in donor (CFP) emission upon photobleaching of the acceptor (NBD). For the CFP-Plin2/NBD-PC FRET pair, emission of the CFP label was imaged at 408 nm excitation (Fig. 3C). Cells were then incubated with NBD-PC and imaged after excitation at 488 nm (Fig. 3E, cells in grayscale). Next, cells were photobleached by repeated scanning at 488 nm until no NBD signal was detected (Fig. 3F). The same photobleached area was then excited at 408 nm (CFP excitation), and a post-bleach CFP image was acquired (Fig. 3D). Many lipid droplets exhibited higher fluorescence intensity after photo-bleaching than observed in the pre-bleach image (Fig. 3C), indicating that CFP-Plin2 and the labeled lipids were in direct proximity. Therefore, the efficiency (E) of the FRET assay was determined experimentally as an image overlay (Fig. 3E, color overlay) as described in materials and methods to identify areas of the cell where FRET occurred. Images of the CFP-Plin2/NBD-PC-labeled cells were made from filtered images of the donor emission before photobleaching, subtracted from images after acceptor photobleaching. The resultant image was divided by the image of donor emission after acceptor photo-bleaching and multiplied by 100 to show the grayscale FRET efficiencies. A FRET overlay was created from this image and pseudo-colored as shown by the inset color scale to visualize regions of higher and lower FRET (Fig. 3E). With CFP-Plin2/NBD-PC labeled cells, areas of high intensity identified by morphology and lipid stain as lipid droplets showed E equal to 60% or greater (red to yellow on the FRET inset color scale) while the calculated mean E for the CFP-Plin2/NBD-PC pair derived from E = 1 − (IDA/ID) was 31 ± 5% (Table 1). Then, from E = 1/[1 − (R/Ro)6], the mean intermolecular distance R between the CFP-Plin2 and NBD-PC FRET pair was calculated as 57 ± 2 Å where Ro for CFP/NBD was equal to 42 Å. In similar fashion, FRET imaging was performed on CFP-Plin2-overexpressing cells incubated with NBD-SM (Fig. 4B), where the mean E and R were calculated as 60 ± 3% and 44 ± 1 Å, respectively (Table 1). Taken together, these results indicate that, in living cells, Plin2 forms a close physical association with phospholipids present in the LD surface monolayer.

Table 1.
FRET efficiency E and distance R between CFP-Plin2 and NBD-labeled lipids

Specificity of CFP-Plin2 targeted interactions determined by live cell targeting and FRET analysis of CFP-Plin2-overexpressing cells incubated with Nile red.

To determine the specificity of CFP-protein/NBD-lipid-targeted interactions measured by FRET, and to also illustrate the limits of optical spectroscopy, colocalization and FRET experiments were repeated with Nile red, another fluorescently labeled molecule that targets lipid droplets (12, 32). As with the NBD-labeled experiments, simultaneous acquisition of confocal images of CFP-Plin2-overexpressing cells incubated with Nile red showed colocalization within lipid droplets (Fig. 5A, yellow lipid droplets), indicating that Nile red molecules interacted with CFP-Plin2 in lipid droplets in living cells to within the limits of optical spectroscopy (2,200 Å). Since CFP also forms a strong FRET pair with Nile red (Fig. 2B), a FRET study with CFP-Plin2-overexpressing cells incubated with Nile red was performed as described earlier. In brief, the emission image of CFP-Plin2-overexpressing cells incubated with Nile red, excited at 408 nm, was compared before and after acceptor photobleaching at 568 nm (Nile red excitation) to remove the Nile red signal (Fig. 5F). The post-bleach image intensity of CFP (Fig. 5D) was similar to that of the CFP pre-bleach intensity (Fig. 5C), indicating that little to no FRET interaction occurred between Nile red and CFP-Plin2. With no change in intensity, the mean E equaled 0 [from E = 1 − (IDA/ID)] and distance R between the probes became >120 Å. Since R was greater than 2 × Ro, no energy transfer between Plin2 and Nile red was indicated (47, 73). These results were more clearly visualized by calculating FRET efficiencies over the entire image as described in materials and methods. Moreover, examination of n > 25 cells showed that the FRET efficiency between Plin2 and Nile red was <0.5% (Fig. 5E). Thus, despite evidence of colocalization on the surface of lipid droplets (Fig. 5A), Nile red was not in sufficient FRET proximity to Plin2 to indicate direct contact.

Fig. 5.
Colocalization and FRET imaging between CFP-Plin2 and Nile red. CFP-Plin2-overexpressing cells were incubated with Nile red to determine colocalization (A) and FRET efficiency (E). The extent of colocalization was shown graphically as a pixel fluorogram ...

Live cell targeting and FRET analysis of CFP-Plin2 with NBD-labeled Chol and stearic acid.

Based on binding studies (7, 27, 54, 61), there is the possibility that Plin2 interacts directly with other lipids found in the lipid droplet monolayer such as Chol (6, 58) and stearic acid (7). Therefore, the ability of CFP-Plin2 to interact with NBD-labeled Chol and stearic acid was examined by live cell LSCM. First, the ability of CFP-Plin2 to colocalize with NBD-labeled lipids such as NBD-Chol (Fig. 4C) and NBD-stearic acid (Fig. 4E) was established. CFP-Plin2-overexpressing cells were incubated with NBD-Chol or NBD-stearate separately and then imaged by LSCM. For the CFP-Plin2/NBD-Chol pair, simultaneous acquisition of confocal images showed strong spatial overlap (yellow-to-orange, Fig. 4C) in highly fluorescent areas identified as lipid droplets by morphology. In similar fashion, NBD-stearic acid incubated with CFP-Plin2-overexpressing cells exhibited strong overlap with the fluorescently labeled lipid droplets (Fig. 4E). The extent of the overlap or colocalization was quantified in pixel fluorograms (data not shown). For the CFP-Plin2/NBD-cholesterol pair, 100% of the CFP probe colocalized with the NBD-stain and 78% of the NBD probe colocalized with CFP probe. With the CFP-Plin2/ NBD-stearic acid pair, correlation coefficients indicated that 99% of the CFP-label (red channel) colocalized with the NBD-label (green channel), while 36% of the NBD stain colocalized with the CFP probe. Overall, these experiments revealed that NBD-labeled lipids colocalized with CFP-Plin2-labeled lipid droplets, indicating strong spatial overlap of Plin2 with lipids on the surface of lipid droplets to within 2,200 Å, the optical limit of light microscopy. It should be considered that while NBD-Chol was a useful probe to examine intracellular Chol dynamics, the overall behavior of the fluorescent analogue was not identical to that of cholesterol. Notably, NBD-Chol uptake/efflux was significantly faster than [3H]-Chol, possibly reflecting the higher aqueous solubility of NBD-Chol versus Chol (8). The faster kinetics of absorption/desorption from plasma membrane receptors observed with NBD-Chol led to limited incorporation of the probe in some membranes (such as the plasma membrane), yet NBD-Chol was esterified similarly as cholesterol and was shown to traffic by similar uptake and secretory pathways (6, 7, 27, 65) making NBD-Chol an acceptable probe for examining intracellular Chol targeting and localization.

While results from the colocalization imaging studies suggested that Plin2 interacts directly with lipids on the lipid droplet surface, optical resolution did not allow a definitive interpretation. Therefore, live cell FRET experiments were performed on the CFP-Plin2/NBD-lipid pairs as described in materials and methods (44, 73). For the CFP-Plin2/NBD-Chol FRET pair, emission of CFP-Plin2-labeled lipid droplets was imaged by exciting the CFP label at 408 nm to determine bleed-through parameters. The CFP-Plin2-labeled cells were then incubated with NBD-Chol, and fluorescent molecules in the desired field were imaged after excitation at 488 nm (Fig. 4D, cells in grayscale), with fluorescence bleed-through to other channels minimized as necessary. Cells in the field were photobleached by repeated scanning at 488 nm until no NBD signal was detected (image not shown). The same photobleached area was then excited at 408 nm (CFP excitation), and a post-bleach CFP-labeled Plin2 image was acquired (image not shown). Several fluorescently labeled lipid droplets in the post-bleach CFP-Plin2 image exhibited stronger intensity than in the pre-bleach CFP-Plin2 image, results consistent with a previous FRET interaction. Image pixel intensities of the pre- and post-bleach CFP-label were then used to calculate the means of E and the intermolecular distance R between the CFP-Plin2 and NBD-Chol as described previously. E and R were calculated to equal 45 ± 4% and 50 ± 1 Å, respectively (Table 1). In order to more clearly visualize the areas in the cell where FRET efficiency was greatest, FRET efficiency images were also generated (Fig. 4D). From these images, several areas of high FRET efficiency were clearly observed within lipid droplets, consistent with FRET interaction between CFP-tagged Plin2 and NBD-labeled Chol.

In similar fashion, FRET efficiencies were determined in CFP-Plin2-overexpressing cells labeled with NBD-stearic acid from the following images: 1) donor (CFP) emission image excited at 408 nm of cells colabeled with CFP-Plin2 and NBD-stearic acid before acceptor (NBD) photobleaching; 2) acceptor (NBD) emission image excited at 488 nm of cells colabeled with CFP-Plin2 and NBD-stearic acid before acceptor photobleaching (Fig. 4F, cells in grayscale); 3) acceptor (NBD) emission image excited at 488 nm of cells colabeled with CFP-Plin2 and NBD-stearic acid after acceptor photobleaching; and 4) donor (CFP) emission image excited at 408 nm of cells colabeled with CFP-Plin2 and NBD-stearic acid after acceptor (NBD) photobleaching. As with the CFP-Plin2/NBD-Chol pair, the FRET overlay with CFP-Plin2 with NBD-stearic acid revealed several areas of increased FRET efficiency (Fig. 4F). Mean FRET efficiencies were found to be 47 ± 4%, with the mean interaction distance R equal to 48 ± 1 Å (Table 1). In summary, FRET with CFP-labeled Plin2 paired to NBD-cholesterol or NBD-stearic acid was measured as increased donor (CFP-label) emission upon bleaching the acceptor (NBD-label) in fluorescently labeled lipid droplets. Calculations of E and R for the two FRET pairs revealed the close mean intermolecular distances between the CFP-labeled Plin2 and NBD-labeled lipids to within 48–50 Å, indicating a strong molecular association between CFP-Plin2 and NBD-lipids on the surface of lipid droplets.

Effect of CFP-Plin2 overexpression on lipid composition.

Results from the FRET studies raised questions on the structural and functional role of Plin2 in lipid droplet biology. Therefore, it was important to establish whether increased expression of Plin2 could influence lipid composition in transfected cells. Lipids from CFP-Plin2 overexpressing and control cell homogenates including CE, DG, TG, Chol, and total phospholipids were extracted and analyzed as described in materials and methods. Consistent with other work that shows increased expression of Plin2 promotes cellular lipid accumulation (29, 48, 50, 51), levels of TG and CE, interior core neutral lipids, were increased 1.9- and 1.7-fold, respectively in CFP-Plin2-expressing cells (Fig. 6A, P < 0.02), In addition, levels of total phospholipids, found in the surface layer of lipid droplets, were each increased 1.5-fold in CFP-Plin2-expressing cells (Fig. 6A, P < 0.05). Since total phospholipids were increased, individual levels of sphingomyelin (SM), phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylinositol (PI), and phosphatidylserine (PS) were next examined in control and CFP-Plin2-overexpressing cell homogenates. Levels of SM, PS, and PC were increased 1.4-, 1.6-, and 1.8-fold in CFP-Plin2-overexpressing cells (Fig. 3B). Cellular levels of PI and PE were not affected by CFP-Plin2 overexpression. Taken together, these results indicate that Plin2 overexpression affected cellular levels of lipids found in both the LD interior core (CE, TG) and surface monolayer (SM, PS, PC).

Fig. 6.
Distribution of individual lipids in CFP-Plin2-overexpressing cells. The mass (nmol/mg protein) of individual lipids (A) and phospholipids (B) in control (open bar) and CFP-Plin2 overexpressing (closed bar) cells was determined as described in materials ...

Effect of CFP-Plin2 overexpression on proteins associated with TG and PL metabolism: HSL, MGAT, DGAT, ATGL, GPAT, CCT, SMS, CETP.

To determine the effect of Plin2 overexpression on enzymes involved in neutral lipid (DG, TG) and phospholipid (PC, SM) synthesis, quantitative analysis of multiple Western blots was performed using antibodies against the following proteins: 1) enzymes involved primarily in neutral lipid metabolism including HSL, MGAT, DGAT, and ATGL (Fig. 7A); 2) enzymes involved in phospholipid (PC, SM) metabolism such as GPAT, CCT, and SMS (Fig. 7B); and 3) enzymes involved in neutral lipid (CE, TG) transport including CETP and HSL (Fig. 7, A and B). Results describing the effect of Plin2 expression on cellular protein and lipid content are summarized in Fig. 8.

Fig. 7.
Effect of CFP-Plin2 overexpression on key proteins involved with TG, Chol, and phospholipid metabolism. Quantitative Western blot analysis was performed on cell homogenates from CFP-Plin2-overexpressing (closed bar) and control (open bar) cells using ...
Fig. 8.
Schematic diagram of neutral lipid and phospholipid biosynthetic pathways. The effect of Plin2 overexpression on levels of key enzymes and lipids involved in TG, CE, and phospholipid synthesis is depicted. In brief, a primary de novo route towards neutral ...

Levels of DGAT1, directly involved in TG synthesis, were increased 2.6-fold but no change was observed with DGAT2. These results were in keeping with the observed increase in TG levels in the CFP-Plin2-overexpressing cells reported here and in other Plin2-overexpressing systems (29, 48, 50). In addition, there was no significant change in expression levels of ATGL, the primary lipase involved in TG hydrolysis (60), resulting in an overall shift towards increased TG. Levels of the enzyme MGAT1, directly involved in DG synthesis (62), were increased but DG levels were balanced by mixed changes observed in enzymes involved in DG hydrolysis/metabolism. For example, levels of HSL significantly decreased 1.2-fold while DGAT1 (involved in forming TG from DG) (62) and CCT (converts DG to PC) (46) increased 2-fold. For proteins involved in PC and SM synthesis (24, 46), levels of CCT and SMS1 (but not SMS2) were increased 2- and 1.3-fold, respectively with no observed changes in levels of GPAT3, the rate-limiting enzyme in the Kennedy pathway to TG synthesis (23). Taken together, these results were consistent with the increased PC and SM content observed in CFP-Plin2-overexpressing cells.

Effect of CFP-Plin2 overexpression on proteins associated with Chol synthesis, uptake, and efflux.

To determine the effect of Plin2 overexpression on enzymes involved in Chol synthesis, uptake, and efflux, quantitative analysis of multiple Western blots was performed using antibodies against the following proteins: 1) HMGR, the enzyme involved in Chol synthesis (19); 2) SR-B1 and LDLR, plasma membrane receptors that govern uptake and efflux of Chol/CE into the cell; and 3) ACAT-1, the enzyme responsible for catalyzing the formation of CE from Chol (22) (Fig. 7C). Levels of HMGR were decreased 1.4-fold in CFP-Plin2-overexpressing cells but no change was observed with SR-B1 or LDLR. In contrast, levels of ACAT-1 increased threefold, in keeping with the significant increase observed in CE levels (Fig. 6A). Interestingly, levels of CETP, an enzyme involved in transporting CE and TG from the ER to lipid droplets (41), were slightly decreased. Despite this, intracellular levels of CE and TG were increased (Fig. 6) in cells overexpressing Plin2, suggesting that Plin2 may act to retain neutral lipids despite decreased transport (CETP), synthesis (HMGR), or lack of change (LDLR, SR-B1) in enzymes involved in Chol metabolism.

DISCUSSION

Recent advances have increased our understanding of the many roles lipid droplets play in maintaining lipid homeostasis (reviewed in Refs. 28, 31, 37, 55), yet molecular details governing lipid exchange across the phospholipid monolayer remain scarce. It was recently shown that Plin1 and other proteins associated with lipolysis including HSL and ATGL target to highly organized areas within the phospholipid monolayer to facilitate lipolytic function (54), suggesting that lipid exchange may be directed by, and be dependent on, the protein and/or lipid composition of the lipid droplet hemi-membrane. Sequestering of proteins with shared function is common in cholesterol-rich areas of the plasma membrane (1, 3, 64) to facilitate cholesterol/fatty acid uptake and efflux (18, 25, 49, 57), but in lipid droplets it represents a novel mechanism to regulate lipid flux across the phospholipid monolayer. Given the surface proximity and ubiquitous nature of Plin2 in all cell types, along with its high binding affinity for several lipid droplet lipids (6, 7, 27, 54, 61), a role for Plin2 in binding lipids and regulating lipid exchange across the surface is strongly indicated. Therefore, the present work was undertaken to examine potential roles of Plin2 in regulating intracellular lipid metabolism through direct lipid interactions on the lipid droplet surface.

Results show for the first time that, in living cells, Plin2 is directly associated with fluorescent lipids on the surface of lipid droplets with protein-lipid interactions in the range of 44–57 Å. This represents an important advance over previous work where colocalization studies indicated that Plin2 interacted with lipids to within the limits of optical resolution (2,200 Å) but did not permit conclusive evidence of a direct physical molecular association (6, 7, 27, 54). Therefore, live cell FRET imaging was performed to determine the extent of interaction between CFP-Plin2 and NBD-labeled lipids on the lipid droplet surface. Since FRET occurs as a result of a nonradiative transmission of energy from an excited donor molecule (CFP) to a nearby acceptor molecule (NBD) and is dependent on the dipole-dipole interactions of fluorescent donor to acceptor molecules, it represents a way to detect molecule-molecule interactions at distances in the range of 10–100 Å. In the case of CFP-Plin2 and NBD-labeled lipids, the measured intermolecular distances R between Plin2 and lipids (PC, SM, Chol, stearic acid) ranged from 44 to 57 Å (Table 1), indicating that Plin2 was directly associated with lipids on the lipid droplet surface. Evidence that both the donor and acceptor molecules reside on the surface monolayer and not within the interior neutral lipid core was based primarily on the fact that higher fluidities of neutral lipids within the inner core greatly increase randomization of the dipole orientation of NBD-lipid acceptor molecules to result in a nonfavorable orientation factor between the FRET donor and acceptor pairs. Moreover, since the diameter of fibroblast lipid droplets is in the range of 2,000–20,000 Å, there is a much greater volume for dilution within the neutral lipid rich core, further decreasing the probability of labeled lipids remaining in close FRET proximity to the protein (where energy transfer occurs as R−6) within the inner core. The volume of the lipid droplet size contrasts with the monolayer thickness which is on the order of 20–30 Å, allowing Plin2 close proximity to lipids and polar probes. In a simple calculation based on a near spherical shape, the diameter of the 47-kDa Plin2 can be estimated to be ~50 Å. Since FRET interaction distances R were in the range of 44–57 Å for the NBD-labeled lipids used herein, it can be predicted that the fluorescent lipids were within the monolayer or bound to the protein at the monolayer interface. In summary, the observed mean interaction distances suggest that, on average, the NBD-labeled lipids were separated from the fluorescent donor CFP molecule to no more than the estimated diameter of the Plin2 protein (50 Å).

The strong molecular interaction between Plin2 and lipids on the lipid droplet surface was also evident from measured E values. E, representing the efficiency of energy transfer between donor and acceptor, is dependent on the distance R and the orientation of the donor and acceptor molecules. E ranged from 30 ± 5% for NBD-PC to 60 ± 3% for NBD-SM (Table 1). In addition, E was quantitated on a pixel-by-pixel basis to generate FRET image maps that identified specific areas within the cell where the percentage of E was highest. Areas of the cell identified by morphology and lipid stain as lipid droplets consistently exhibited E between 60 and 100% as designated by the red to yellow colored areas on the FRET image maps (Figs. 2E, ,4B,4B, ,4D,4D, and and4F).4F). These images clearly identified the FRET-active sites within the cell where Plin2 was in direct contact with the labeled lipids on the surface of the lipid droplet. It should be noted that Nile red, a lipid probe that targeted to lipid droplets and colocalized with Plin2 (Fig. 5A), did not show observable FRET with CFP-labeled Plin2. The measured distance R between CFP-Plin2 and Nile red exceeded 120 Å, and E was <0.5%, indicating that Nile red was not closely associated with Plin2. Thus, despite the potential for FRET between CFP-Plin2 and Nile red as evidenced by the strong spectral overlap between CFP emission and Nile red excitation spectra (Fig. 2B), distances in living cells were beyond FRET proximity (Fig. 5E). In all, these studies provided evidence that Plin2 selectively associates with several lipids on the surface of lipid droplets.

The physical association between Plin2 and NBD-labeled lipids was also corroborated from fluorescence binding studies where binding affinities in the nanomolar range (15–257 nM) were exhibited by Plin2 for NBD-PC (54), NBD-SM (54), NBD-stearic acid (61), and NBD-Chol (7, 27). From these studies the extent of interaction between lipid and Plin2 could be approximated by examining the NBD-lipid emission maxima shifts in the presence of Plin2. The emission maxima for NBD-labeled stearic acid, PC, and SM red-shifted 5–9 nm in the presence of Plin2, indicating that the NBD group in the ligand binding site sensed a more polar environment as compared to a lipid micelle in solution (54, 61). In contrast, NBD-cholesterol was blue-shifted 15 nm upon binding (27), indicating a more hydrophobic environment. One possible explanation for the more polar or hydrophobic environment sensed by Plin2 bound NBD-lipids may be the orientation of the respective ligands in the Plin2 lipid binding site. The X-ray crystal structure of Plin2 has yet to be determined, but the COOH-terminal domain of TIP47 has been resolved and this region is highly conserved between TIP47 and Plin2 (35). In the COOH-terminal domain of TIP47 (and presumably in other members of the PAT family with homology in this region including Plin2) a deep hydrophobic cleft between a α/β domain and a four-helix bundle domain was found that was rimmed by basic residues. Cleft size, shape, and hydrophobicity suggested this site as a possible area of interaction for hydrophobic molecules such as fatty acids and sterols (35). Based on this, binding in this region would allow the cholesterol and fatty acid groups of the NBD-lipids to bury deep in the hydrophobic cleft with differences in hydrophobicity of the cholesterol versus fatty acyl and polar head groups dictating the orientation of the attached NBD group. The NBD-group attached to the acyl chains in NBD-labeled stearic acid, PC, or SM could be expected to be relatively closer to the opening (more polar) of the Plin2 binding pocket than the NBD group in NBD-cholesterol (attached to carbon 22 in the alkyl tail). Consistent with the position of the fluorescent label within the lipid, NBD-labeled stearic acid, PC, and SM binding to Plin2 resulted in relatively small red-shifts of the NBD emission spectrum (up to 9 nm), indicating that the NBD group was bound near the surface opening of the Plin2 binding pocket in closer proximity to the basic residues on the rim of the hydrophobic cleft while the blue-shift experienced by the NBD-cholesterol ligand upon binding indicated the NBD group was located more in the hydrophobic cleft. In all, given the surface proximity of Plin2, the high lipid binding affinity resulting in fluorescence emission shifts, and the strong FRET interactions with lipids in the lipid droplet monolayer, a role for Plin2 in binding to, and directing lipid exchange across, the lipid droplet surface is strongly indicated.

It was clear from fluorescence binding studies (7, 27, 54, 61) that Plin2 was capable of high-affinity binding with lipids found in the phospholipid monolayer including PC, SM, Chol, and stearic acid. The FRET studies in the present work clarified these interactions to reveal a close physical association, indicating that the lipids were either bound in the ligand binding pocket and/or immediately surrounding the protein, much as in a lipid shell (1, 42). In the lipid shell model, proteins surrounded by lipid molecules in regions of <10 nm have a dynamic composition that depends upon protein-lipid, lipid-lipid, and nearby protein-protein interactions (26). In the lipid droplet monolayer, these interactions would allow direct access to nearby proteins and could serve to anchor the proteins into the lipid monolayer in select domains that would facilitate protein-lipid interactions along the hemi-membrane. Consistent with this theory, several proteins involved in lipolysis of TG from the lipid droplet neutral lipid core were recently found localized in highly organized, cholesterol-enriched areas of the lipid droplet monolayer, presumably to help facilitate the lipolytic process (66). While Plin2 was not part of that pool (consistent with the fact that Plin2 plays no known part in lipolysis), these results lend support to a domain-driven hypothesis where lipid exchange across the monolayer is mediated based on changes in the protein (and lipid) composition of the phospholipid hemi-membrane. This hypothesis is supported by recent studies where increased expression of Plin2 was shown to reduce the association of other proteins for the lipid droplet surface through differential binding affinity and displacing ability based on changes in surface hydrophobicity in the phospholipid packing (50). In the present work, lipid binding characteristics of Plin2 correlated very well with lipid specificity as evidenced by the FRET data. Results suggest that lipid exchange from the interior core to the outer surface may involve a process whereby high-affinity binding of lipids to Plin2 on the lipid droplet surface and/or Plin2 in direct proximity with a shell of lipids subsequent to binding to the protein helped facilitate the transfer of lipids through the monolayer into/from the neutral lipid-rich inner core where changes in the lipid composition at the surface could serve to regulate the exchange. Moreover, sequestering of lipids to specific areas of the hemi-membrane through protein-lipid interactions may be critical to enzyme function and help define a lipid zip code for lipid exchange across the lipid droplet surface.

Since the physiological significance of the above results may be based on the context of Plin2's influence on key enzymes involved in the synthesis of lipids necessary for lipid droplet structure and function, the effect of Plin2 overexpression on cellular metabolism was also examined. This was demonstrated by directly correlating results from lipid analysis with expression changes in proteins related to TG, Chol, and phospholipid metabolism. The lipid profile reflected the potential role of Plin2 in regulating intracellular lipid storage since levels of the primary lipids found in the interior neutral lipid core (CE, TG) and the phospholipid monolayer (PC, PS, SM) were significantly increased in the CFP-Plin2-overexpressing cells. In keeping with the lipid changes, expression levels of proteins associated with TG, CE, and phospholipid synthesis were upregulated in CFP-Plin2-expressing cells. Specifically, expression levels of proteins involved primarily in neutral lipid (DG, TG) metabolism including HSL, MGAT, DGAT, and ATGL were selectively regulated in CFP-Plin2-overexpressing cells to promote increased TG levels (i.e., increased MGAT, DGAT, decreased HSL, no change in ATGL or perilipin). Levels of DG, standing at the crossroads to TG and phospholipid synthesis, were not affected by Plin2 expression changes, balanced by mixed changes in expression of proteins involved in DG metabolism (i.e., HSL was significantly decreased and levels of MGAT, DGAT and CCT increased). These results were in keeping with the role of DG as a key lipid second messenger (reviewed in Ref. 69), suggesting that careful regulation of this lipid was important to cellular function. Like TG, CE levels were increased in CFP-Plin2-overexpressing cells, reflecting the threefold increase in ACAT-1, an enzyme responsible for esterifying Chol. With respect to phospholipid synthesis, levels of CCT and SMS were upregulated in CFP-Plin2-overexpressing cells, leading to higher levels of total phospholipids. It should be noted that while previous studies document that Plin2 overexpression is associated with increased TG (29, 48, 50) and CE (51) levels, the effect of Plin2 overexpression on phospholipid content is first reported here where levels of PC, PS, and SM were increased up to 1.8-fold in CFP-Plin2-overexpressing cells (Fig. 3B). As with most membranes, the major phospholipid found in the lipid droplet monolayer was PC with lower levels of PE, PS, PI, and SM also present. A lack of SM, PS, and PA in lipid droplets from Chinese hamster ovary (CHO) K2 cells was reported in another study (9) but subsequent work with other cell types and tissue including HepG2 cells (68), Niemann-Pick cells (45), alveolar macrophages (40), mammary tissue (38), and adipose tissue (39, 54) record the presence of these lipids in isolated lipid droplet fractions. Since phospholipids form the boundary between the aqueous environment of the cytoplasm and the interior neutral lipid core, their importance in maintaining lipid droplet structure is clear. Recently, phospholipid remodeling of PC (residing in the ER) to PS and later PE was shown to incorporate into the lipid droplet monolayer (39). Moreover, the enzyme PEMT was able to convert the remodeled PE into PC, suggesting that this enzyme plays an important role in lipid droplet biogenesis. In the present work, overexpression of Plin2 through cellular modulation of key enzymes in the phospholipid synthetic pathway led to significant increases in several phospholipids found in the lipid droplet monolayer, providing evidence that Plin2 plays a valuable role not only in TG metabolism but also in phospholipid biosynthesis.

In summary, the present work describes two important findings not previously reported: 1) Plin2 forms a direct physical association with lipids involved in lipid droplet structure (PC, SM) and function (Chol, FA); and 2) Plin2 overexpression affects TG, CE, and phospholipid synthesis by modulating levels of key enzymes involved in lipolysis and lipogenesis. In all, this work presents a novel view of the role of Plin2 in lipid droplet biology and supports the hypothesis that Plin2 can regulate lipid exchange from lipid droplets by facilitating direct protein-lipid interactions on the lipid droplet surface.

GRANTS

This work was supported by National Institutes of Health Grant DK-70965.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

A.L.M. and B.P.A. conception and design of the research; A.L.M., S.S., K.C.M., S.G., J.S.L., C.C.M., S.M.S., and B.P.A. performed the experiments; A.L.M., S.S., K.C.M., S.G., J.S.L., C.C.M., S.M.S., and B.P.A. analyzed the data; A.L.M., S.S., K.C.M., S.G., J.S.L., C.C.M., S.M.S., and B.P.A. interpreted the results of the experiments; A.L.M. and B.P.A. prepared the figures; A.L.M. and B.P.A. drafted the manuscript; A.L.M., S.S., K.C.M., and B.P.A. edited and revised the manuscript; A.L.M., S.S., K.C.M., S.G., J.S.L., C.C.M., S.M.S., and B.P.A. approved the final version of the manuscript.

ACKNOWLEDGMENTS

The technical assistance of Meredith Dixon and Ashley R. Stone is much appreciated.

Glossary

ACAT-1
acyl-coenzyme A:cholesterol acyltransferase-1 (also known as sterol-acyltransferase 1)
ADRP
adipose differentiation-related protein
ATGL
adipose triglyceride lipase
CCT
phosphocholine cytidylyltransferase
CE
cholesteryl ester
CETP
cholesteryl ester transport protein
CFP
cyan fluorescent protein
Chol
cholesterol
CGI-58
comparative gene identification-58
Chol
cholesterol
DG
diacylglycerol
DGAT
diacylglycerol acyltransferase
ECFP
enhanced cyan fluorescent protein
ER
endoplasmic reticulum
FA
fatty acid
G3P
glycerol-3-phosphate
GFP
green fluorescent protein
GPAT
glycerol-3-phosphate acyltransferase
HMGR
3-hydroxy-3-methylglutaryl-CoA reductase
HSL
hormone sensitive lipase
LD
lipid droplet
LDLR
low-density lipoprotein receptor
LPA
lysophosphatidic acid
LSCM
laser scanning confocal microscopy
MAGL
monoacylglycerol lipase
MG
monoacylglycerol
MGAT
monoacylglycerol acyltransferase
NBD-cholesterol
[22-(N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino)-23,24-bisnor-5-cholen-3b-ol]
NBD-stearate
12-N-methyl-(7-nitrobenz-2-oxa-1,3-diazo)aminostearic acid
NBD-PC
1-acyl-2-(N-4-nitrobenzo-2-oxa-1,3-diazole)-aminocaproyl phosphatidylcholine
Nile red
7-diethylamino-3,4-benzophenoxazine-2-one
PA
phosphatidic acid
PAP
phosphatidic acid phosphatase
PC
phosphatidylcholine
PE
phosphatidylethanolamine
PEMT
phosphatidylethanolamine N-methyltransferase
PI
phosphatidylinositol
PL
phospholipid
Plin2
perilipin 2
PM
plasma membrane
PS
phosphatidylserine
SM
sphingomyelin
SMS
sphingomyelin synthase
SR-B1
scavenger receptor-B1
TG
triacylglycerol

Footnotes

1This article is the topic of an Editorial Focus by Jeffrey S. Elmendorf (25a).

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