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Perilipin family members preferentially sequester toeither triacylglycerol-specific or cholesteryl-ester-specific intracellular lipid storage droplets

Kai Hsieh1, Yun Kyung Lee1, Constantine Londos1, Bruce M. Raaka2, Knut Tomas Dalen1,3 andAlan R. Kimmel1,*1Laboratory of Cellular and Developmental Biology, National Institute of Diabetes and Digestive and Kidney Diseases, The National Institutes ofHealth, Bethesda, MD 20892, USA2Clinical Endocrinology Branch, National Institute of Diabetes and Digestive and Kidney Diseases, The National Institutes of Health, Bethesda,MD 20892, USA3Department of Nutrition, Institute of Basic Medical Sciences, Medical Faculty, University of Oslo, PO Box 1046 Blindern, N-0316 Oslo, Norway

*Author for correspondence (ark1@helix.nih.gov)

Accepted 15 May 2012Journal of Cell Science 125, 4067–4076� 2012. Published by The Company of Biologists Ltddoi: 10.1242/jcs.104943

SummaryPerilipin family proteins (Plins) coat the surface of intracellular neutral lipid storage droplets in various cell types. Studies across diversespecies demonstrate that Plins regulate lipid storage metabolism through recruitment of lipases and other regulatory proteins to lipiddroplet surfaces. Mammalian genomes have distinct Plin gene members and additional protein forms derived from specific mRNA

splice variants. However, it is not known if the different Plins have distinct functional properties. Using biochemical, cellular imagingand flow cytometric analyses, we now show that within individual cells of various types, the different Plin proteins preferentiallysequester to separate pools of lipid storage droplets. By examining ectopically expressed GFP fusions and all endogenous Plin protein

forms, we demonstrate that different Plins sequester to different types of lipid droplets that are composed of either triacylcerides orcholesterol esters. Furthermore, Plins with strong association preferences to triacylceride (or cholesterol ester) droplets can re-direct therelative intracellular triacylceride–cholesterol ester balance toward the targeted lipid. Our data suggest diversity of Plin function, alter

previous assumptions about shared collective actions of the Plins, and indicate that each Plin can have separate and unique functions.

Key words: PLIN, ADRP, TIP47, LSDP5, S3-12, Triacylglyceride, Cholesterol, Fatty acids, Lipolysis

IntroductionIntracellular neutral lipid storage droplets (LSDs) are unique

organelles that store metabolic precursors of cellular energy,

membrane biosynthesis, steroid hormone synthesis, and signaling

(Farese and Walther, 2009; Kimmel et al., 2010; Londos et al.,

2005). LSDs contain different lipids [e.g. triacylglycerides

(TAGs) or cholesteryl esters (CEs)] at their core, surrounded

by a phospholipid monolayer. LSD surfaces in organisms as

diverged as mammals, Drosophila and Dictyostelium are targeted

by an evolutionarily related family of proteins (Kimmel et al.,

2010; Lu et al., 2001; Miura et al., 2002), the Perilipins (Plins).

Mammalian genomes have five distinct Plin gene members and

additional protein forms derived from specific mRNA splice

variants (Kimmel et al., 2010).

Plin1 (Perilipin 1) is the major LSD coat protein in adipocytes

and steroidogenic cells (Greenberg et al., 1993; Servetnick et al.,

1995). Other Plins exhibit different expression patterns. Plin2 is

the predominant, but not exclusive, form in liver (Dalen et al.,

2006), whereas Plin5 is primarily expressed in oxidative tissues,

including heart, soleus muscle, and brown fat (Dalen et al., 2007;

Wolins et al., 2006; Yamaguchi et al., 2006). Based on Plin1

function (Martinez-Botas et al., 2000; Sztalryd et al., 2003;

Tansey et al., 2001; Wang et al., 2009), the Plins are viewed as

fundamental regulators of lipolytic activity. Loss of Plin1

(Martinez-Botas et al., 2000; Tansey et al., 2001) or Plin2 (also

known as ADRP) (Chang et al., 2006) in mice significantly

reduces intracellular lipid levels in adipocytes and hepatocytes,

respectively. Furthermore, heterozygous loss-of-function

mutations in human PLIN1 leads to a familial partial

lipodystrophy, supporting a required role for Perilipin in TAG

storage within human adipocyte LSDs (Gandotra et al., 2011).

Regardless, little is known of lipid interaction specificity of the

various Plins. Here, we show that distinct Plins differentially

sequester to either TAG- or CE-specific LSDs and can alter

relative intracellular TAG or CE levels toward the preferentially

targeted lipid. These data demonstrate and emphasize diverse

functions for the different Plins.

ResultsExogenous fatty acids and cholesterol differentially

stabilize accumulation of Plin protein family members

Intracellular LSDs accumulate substantially when cells are

cultured overnight in the presence of high concentrations of

various exogenous lipids (Xu et al., 2005). Since Plins primarily

This is an Open Access article distributed under the terms of the Creative Commons AttributionNon-Commercial Share Alike License (http://creativecommons.org/licenses/by-nc-sa/3.0/),which permits unrestricted non-commercial use, distribution and reproduction in any mediumprovided that the original work is properly cited and all further distributions of the work oradaptation are subject to the same Creative Commons License terms.

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sequester to LSD surfaces (Miura et al., 2002), we determined if

different Plins exhibited differential regulation in response to

either fatty acids or cholesterol, lipids that mobilize separate

pathways.

Y1 mouse adrenocortical cells have robust capacity for steroid

hormone synthesis and accumulate TAG and CE LSDs as energy

and metabolic precursor stores. Further, steroidogenic cells are

able to synthesize all 4 Plin1 mRNA splice variants (Servetnick

et al., 1995; Xu et al., 2005) and express all other Plin genes.

Y1 cells were cultured under standard conditions or in medium

supplemented with oleic acid and/or cholesterol. Endogenous

Plin proteins were quantified in whole cell lysates by specific

immunoblotting (Fig. 1A). In general, none of the Plins exhibited

significant accumulation in unsupplemented medium. However,

significant Plin accumulation differences were observed in the

presence of oleic acid or cholesterol. The two major Plin1

variants of steroidogenic cells, Plin1a and Plin1c, exhibited

reciprocal patterns. Plin1a was enhanced by oleic acid, but not by

cholesterol, whereas the Plin1c response was exactly opposite

(Fig. 1A). The effects were largely activating, since the

expressions of Plin1a and Plin1c were not diminished in cells

cultured simultaneously with oleic acid and cholesterol. Plin1b

and Plin1d proteins are not easily detected in Y1 cells (Servetnick

et al., 1995), although Plin1b appears to be regulated similarly to

Plin1a (Fig. 1A).

Plin2 and Plin3 (also known as TIP47) accumulate similarly

regardless of the exogenous lipid moiety, although Plin2 may be

slightly more responsive to oleic acid. Conversely, Plin4 (also

known as S312) and Plin5 (also known as LSDP5) show extreme

lipid specificity, largely mimicking that of Plin1c and Plin1a,

respectively (Fig. 1A).

Since exogenous lipids may have differential regulatory effects

on the transcription or translation of endogenous Plin mRNAs

and, thus, indirectly impact Plin protein accumulation, we also

examined the effects of oleic acid and cholesterol using GFP–

Plin protein fusions expressed from identical constitutively active

promoter vectors. McARH7777 rat hepatoma cells were

transiently transfected with vectors to separately express each

GFP–Plin protein fusion and cultured under standard conditions

or in medium supplemented with oleic acid and/or cholesterol.

The GFP–Plin proteins in McARH7777 cells showed identical

responses to oleic acid and cholesterol as their endogenous

counterparts in Y1 cells (Fig. 1A,B). GFP–Plin1a and GFP–Plin5

were specifically responsive to the positive effects of oleic acid,

whereas GFP–Plin1c and GFP–Plin4 were only detected in the

presence of cholesterol (Fig. 1B). GFP–Plin2 and GFP–Plin3 did

not show a significant preference to either lipid. The responses of

Plin1b and Plin1d were more clear in this heterologous

expression system. Plin1b is structurally most similar to Plin1a

(Lu et al., 2001) and GFP–Plin1b behaves identically to both

Plin1a and GFP–Plin1a. Plin1d is the smallest variant (Lu et al.,

2001) and GFP–Plin1d shows limited lipid preference, acting

more like Plin2 and Plin3.

Differential sub-cellular localizations of tagged fatty acidand cholesteryl markers

Data (Fig. 1) suggest that the sequestration and stabilization of

individual Plins to different classes of LSDs may influence the

specific accumulation of particular Plin proteins. To examine this

further, we established conditions to preferentially tag and purify

TAG- and CE-specific LSDs.

Y1, McARH7777 and AML12 cells were cultured overnight in

the presence of both oleic acid and cholesterol, plus BODIPY

558/568 C12 [as a fluorescent fatty acid (FA) dye marker] and

cholesteryl BODIPY 500/510 FL C12 [as a fluorescent cholesteryl

(Chl) dye marker] and imaged (Fig. 2A). All three types of cells

showed definitive separation of FA (red) and Chl (green) markers

into distinct LSDs clusters. Few sectors show any colocalization,

a pattern similarly observed in 4T1 mouse mammary tumor cells,

primary mouse liver cells, C2C12 mouse myoblasts, 3T3-L1

mouse fibroblasts, J774A.1 mouse monocyte-macrophages, and

CHOK1 Chinese hamster ovary cells (supplementary material

Fig. S1A). In general, the different markers, though distinctly

separate, were largely intermingled. However, the McARH7777

cells (Fig. 2A) were most distinctive. Droplets with tagged FAs

(hereafter referred to as FA-tagged droplets) segregated to

entirely separate sub-cellular regions to those with tagged Chl

(hereafter referred to as Chl-tagged droplets). FA-tagged droplets

were polarized to the cell periphery, while Chl-tagged droplets

were centrally localized. Less distinctive labeling and polarized

separation of the droplets is seen in the McARH7777 cells

cultured for shorter incubation periods (supplementary material

Fig. S1B).

The polarized localizations of FA- and Chl-tagged droplets

suggested that it might be possible to distinguish separate

associations of these distinct droplets. McARH7777 cells were

cultured with oleic acid, cholesterol, and FA- or Chl-tagged

fluorescent markers, and co-stained using antibodies against

several organelle-specific proteins (Fig. 2B) or organelle-specific

dye trackers (Fig. 2C). In McARH7777 cells, the FA-tagged

droplets localized entirely separate from lysosomes,

Fig. 1. Differential accumulation of Plins in cells cultured in the absence or

presence of fatty acid and/or cholesterol. (A) Y1 adrenal cells were cultured

overnight in the absence or presence of oleic acid and/or cholesterol. Whole cell

lysates were prepared and endogenous Plins assayed by immunoblotting. Data

are representative of three experiments. (B) McARH7777 rat liver cells were

transiently transfected with the indicated GFP–Plin-expressing constructs and

cultured overnight in the absence or presence of oleic acid and/or cholesterol.

Whole cell lysates were prepared and GFP–Plin assayed by immunoblotting.

Data are representative of three experiments.

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mitochondria, and early endosomes (Fig. 2B,C). The Chl-tagged

droplets and other organelles are more centrally localized and

perinuclear. Nonetheless, there was nominal overlap of Chl-

tagged droplets with these other structures (Fig. 2B,C).

Lysosomes and mitochondria remained largely separate, and

there was minimal intermingling with early endosomes. The data

indicate that the FA or Chl fluorescent tags do not broadly label

these other organelles.

FACS purification of LSDs enriched in either TAG or CE

To determine if FA- and Chl-tagged LSDs had distinct lipid

compositions, we first developed conditions for their separate

purifications. Labeled Y1, McARH7777 and AML12 cells were

lysed, LSDs floated by gradient centrifugation, and the FA- and

Chl-fluorescently tagged LSDs separated by FACS (Fig. 3A;

supplementary material Fig. S2A–C). The FA- and Chl-specific

labelings were relatively similar and reproducible. Generally, .70%of the particles were dye tagged in dual label experiments. FACSseparations were distinct with ,5% of unsorted particles carryingboth FA and Chl dye markers (Fig. 3A,B). The particles also

showed reasonable size homogeneity; ,80% of all particles haddiameters of 2–6 mm. Experiments with a mixture of labeled andunlabeled lysates and particles showed ,2% dye marker transfer

and/or non-specificity (supplementary material Fig. S2A–C).

Lipids were extracted from sorted FA- and Chl-tagged LSDs

and analyzed and quantified by thin layer chromatography(Fig. 3C; Table 1; supplementary material Fig. S3A). Total LSDcontent was similar within and between cell types. The Chl-tagged

droplets were primarily comprised of cholesteryl ester and asmaller amount of cholesterol; we were unable to detecttriacylglycerols, fatty acids and related metabolites in the sortedChl-tagged droplets. Conversely, the isolated FA-tagged droplets

were comprised primarily of triacylglycerol and minor quantitiesof metabolites (di- and mono-acylglycerides and fatty acids); smallamounts of cholesterol were present, but cholesteryl ester was not

detected. Thus, we have established conditions to fluorescentlytag, image and purify LSDs specific to either CE or TAG.

We also traced the cell fate of the BODIPY precursor markers(supplementary material Fig. S3B). The native BODIPY–Chlmigrates during TLC at a unique position that differs from both

cholesterol and cholesterol ester and retains its identical motilityeven after integration within the CE droplets. Thus, BODIPY–Chl appears to be incorporated directly into CE droplets and isnot significantly metabolized into other droplet-specific lipids

during the time frame studied. BODIPY–FA migrates duringTLC less far than does untagged FA, but changes mobility whenintegrated into TAG droplets. We suggest that this represents the

incorporation of the BODIPY–FA into TAG, which migrates lessfar than untagged TAG and distinctly from phospholipids.Regardless, the FA and Chl markers represent specific tracer

tags that allow us to separately purify the distinct populations ofLSDs that are biochemically enriched in either TAG or CE,hereafter referred to as TAG-LSDs and CE-LSDs, respectively.

Plin associations with TAG- or CE-LSDs

We next examined Plin protein associations with isolated TAG-or CE-LSDs. Y1 cells were cultured with oleic acid andcholesterol, plus FA- and Chl-fluorescent markers. LSDs were

floated, TAG- or CE-LSDs purified by FACS, and associations ofY1 endogenous Plins within unsorted LSDs or to TAG- or CE-specific populations then assayed by specific immunoblotting.

All of the Plin proteins were detected in the unsorted LSDpopulation (Fig. 3D), consistent with previous data (Fig. 1A,B).

The relative increase in endogenous Plin1b and Plin1d signalsprobably reflects the enrichment of these forms in floatingdroplets compared to whole cell lysates (see Fig. 1A, Fig. 3D). Inthis assay, the individual Plins show distinct sequestration to

TAG- or CE-specific LSDs. Plin1a, Plin1b and Plin5 arerestricted to TAG populations, whereas Plin1c and Plin4localize with the CE-LSDs (Fig. 3D). Plin2, Plin3 and Plin1d

show limited association preference (Fig. 3D).

Sub-cellular localization of GFP-Plins to TAG- or CE-LSDsThe whole cell accumulations of specific Plin proteins in

response to either oleic acid or cholesterol correlate with theassociations of the same Plins, to either TAG- or CE-LSDs(Fig. 1, Fig. 3D). Still, we wished to investigate specific Plin/

Fig. 2. Spatially distinct intracellular accumulations of fatty acid and

cholesteryl dye markers. (A) Y1 adrenal, McARH7777 rat liver and AML12

mouse liver cells were cultured overnight in the presence of oleic acid and

cholesterol, plus fatty acid BODIPY 558/568 C12 (FA) and cholesteryl

BODIPY 500/510 FL C12 (Chl). Representative confocal images are shown

with red and green indicating the localization of FA and Chl, respectively.

(B) McARH7777 cells were cultured overnight in the presence of oleic acid

and cholesterol, plus fatty acid BODIPY 558/568 C12 and cholesteryl

BODIPY 500/510 FL C12 and probed for organelle localization using

immunofluorescence detection of LAMP1 (Lysosomal-associated membrane

protein 1), TOM2 (Translocase of outer mitochondrial membranes 20 kDa)

and EEA1 (Early endosome antigen 1). (C) McARH7777 cells were cultured

overnight in the presence of oleic acid and cholesterol, plus fatty acid

BODIPY 500/510 C12 or cholesteryl BODIPY 500/510 FL C12 and probed for

organelle localization by staining with organelle-specific markers.

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droplet associations by alternative methods. Since McARH7777

cells accumulate TAG- and CE-LSDs that are completely

separated (Fig. 2A), we could visually observe if GFP–Plins

exhibited parallel localization differences. In addition we could

quantify the relative distributions of GFP–Plins among CE- or

TAG-LSDs by FACS.

Fig. 3. Differential localization of Plins to TAG or CE droplets. Y1 adrenal, McARH7777 rat liver and AML12 mouse liver cells were cultured overnight in

the presence of oleic acid and cholesterol, plus fatty acid BODIPY 558/568 C12 (FA) and cholesteryl BODIPY 500/510 FL C12 (Chl). (A) Representative FACS

profiles of purified FA- and Chl-tagged lipid droplets are shown, with relative distributions of total lipid droplet numbers indicated in the different quadrants

(FA2/Chl2, FA+/Chl2, FA2/Chl+ and FA+/Chl+); blue represents dual signals of the FA and Chl probes. Y1 cells: 25,000 total droplets; McARH7777 cells:

20,000 total droplets; AML12 cells: 25,000 total droplets. (B) Confocal images of unsorted and FACS sorted FA- and Chl-tagged lipid droplets from Y1,

McARH7777 and AML12 cells. (C) Lipids isolated from sorted FA- and Chl-droplets were separated by TLC in parallel with lipid markers and detected by

staining with iodine vapor. 16 indicates 26105 LSD particles; 56 indicates 16106 LSD particles. Marker lanes have 50 mg CE (*) and 10 mg each of TAG, FA,

diacylglycerides (DAG), cholesterol (Chol), MAG and phospholipid (PL). PLs do not migrate from the origin in this system. (D) Proteins were prepared from

unsorted and FACS-sorted TAG- or CE-LSD isolated from Y1 cells. Endogenous Plin proteins were assayed by immunoblotting. Identical numbers of lipid

droplet particles were loaded in each lane. Data are representative of three experiments.

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McARH7777 cells were transiently transfected individually

with vectors that specifically encode different eGFP–Plin fusions

and cultured with oleic acid and cholesterol plus either BODIPY

558/568 C12 (TAG marker) or cholesteryl BODIPY 576/589 C11

(CE marker). Both dye markers have excitation/emission spectra

that differ from eGFP (488/509). GFP/TAG and GFP/CE

experiments were always performed in pairs and data analyzed

in parallel to assess reciprocal responses (Figs 4, 5, 6;

supplementary material Fig. S4).

GFP–Plin1a localizes to the polarized periphery of

McARH7777 cells and primarily with TAG-LSDs (Fig. 4A,

Fig. 6). In contrast, while CE-LSDs are sequestered to the cell

interior, none of the GFP–Plin1a signal colocalizes with the CE

marker (Fig. 4A). FACS profiles of cell populations support these

Table 1. Lipid specificities in FACS-purified lipid droplets

Lipid

Chl-tagged droplets FA-tagged droplets

Y1 McA AML12 Y1 McA AML12

CE 90 mg 65 mg 70 mg ND (,3 mg) ND (,3 mg) ND (,3 mg)TAG ND (,0.5 mg) ND (,0.5 mg) ND (,0.5 mg) 70 mg 100 mg 100 mgFA ND (,0.5 mg) ND (,0.5 mg) ND (,0.5 mg) 10 mg 15 mg 20 mgDAG ND (,0.5 mg) ND (,0.5 mg) ND (,0.5 mg) 20 mg 10 mg 10 mgChol 50 mg 15 mg 15 mg 3–5 mg 3–5 mg 3–5 mgMAG ND (,2 mg) ND (,2 mg) ND (,2 mg) 15 mg 10 mg 10 mg

Chl- and FA-tagged lipid droplets from Y1, McARH7777 and AML12 cells were separated by FACS, and lipid profiles analyzed by TLC (see Fig. 3C). Values(from 26106 droplets) were extrapolated (630%) from relative staining intensity data in comparison to standards (see supplementary material Fig. S3B) for CE,cholesteryl ester; TAG, triacylglycerol; FA, fatty acid; DAG, diacylglycerol; Chol, cholesterol; and MAG, monoacylglycerol. Phospholipid spot staining wassimilar for each, but did not resolve from the origin. ND, not detected.

Fig. 4. Differential localization of Plin1 proteins to TAG or CE lipid droplets. McARH7777 cells were transiently transfected with the indicated GFP–Plin1-

expressing constructs and cultured overnight in the presence of oleic acid and cholesterol, plus either BODIPY 558/568 C12 (TAG) or cholesteryl BODIPY 576/

589 C11 (CE). Representative confocal images are shown for each with red indicating the localization of either TAG or CE droplets, green indicating localization

of GFP–Plin1 proteins, and yellow (or rings) indicating colocalization. Representative FACS profiles of TAG- and GFP-labeled lipid droplets or of CE- and

GFP-labeled lipid droplets are shown, with relative distributions of total particle numbers indicated in the different quadrants, and the relative Plin1 variant

localizations indicated as a percentage of the total GFP+ signal that co-sorts (blue) with either TAG+ or CE+ tags. Each experiment was performed at least three

times (see Fig. 6). (A) Plin1a (25,000 total droplets for each population); (B) Plin1b (20,000 total droplets for each population); (C) Plin1c (25,000 total droplets

for each population); (D) Plin1d (10,000 total droplets for each population).

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conclusions; ,95% of expressed GFP–Plin1a co-sorts with the

TAG marker, while only ,5% GFP–Plin1a co-sorts with CE-

LSDs (Fig. 4A, Fig. 6).

GFP–Plin1b also exhibits a localization preference for TAG-

LSDs compared to CE-LSDs, although the FACS profiles show

slightly less specificity (Fig. 4B, Fig. 6).

GFP–Plin1c shows the expected reciprocal pattern to GFP–

Plin1a (see Fig. 1, Fig. 3D). GFP–Plin1c localizes to the cell

interior, surrounding CE-LSDs, and distinctly separate from the

polarized TAG-LSDs (Fig. 4C). The FACS data are consistent,

where .90% of GFP–Plin1c sorts with CE and separate from

TAG (Fig. 4C, Fig. 6).

The GFP fusions of Plin1d, Plin2, and Plin3 show minimal

preference for TAG- or CE-LSDs and segregate with both

(Fig. 4D, Fig. 5A,B). All 3 protein fusions are found in both

interior and peripheral cell regions, colocalize with both TAG-

and CE-LSDs within the cell, and co-segregate with both by

FACS, but to varying degrees (Fig. 4D, Fig. 5A,B, Fig. 6).

GFP–Plin4 and GFP–Plin5 exhibit reciprocal LSD preferences

and are, thus, respectively, similar to Plin1c and Plin1a (see Figs 1,

3). While GFP–Plin4 sequesters with CE-LSDs in the cell interior

(Fig. 5C), GFP–Plin5 polarizes with TAG-LSDs to the cell

periphery (Fig. 5D). FACS data substantiate these preferences;

,85% of GFP–Plin4 co-sorts with a CE marker, while ,85% of

GFP–Plin5 segregates with TAG (Fig. 5C,D, Fig. 6).

Plins with TAG- or CE-binding preferences can alter

cellular TAG/CE distributions

Loss of Plin1 and Plin2 in mice alters the targeted accumulation

of lipid levels in defined cell types (Martinez-Botas et al., 2000;

Tansey et al., 2001; Chang et al., 2006). We were, thus, interested

to determine if there were a preferential relationship among

individual Plin proteins, their lipid targeting preference, and the

cellular accumulation of either TAG or CE. We first attempted

siRNA approaches to deplete Plins targeted to either TAG- or

CE-LSDs. Data from other systems clearly show that depletion of

Fig. 5. Differential localization of Plin2, -3, -4 and -5 proteins to TAG or CE lipid droplets. McARH7777 cells were transiently transfected with the indicated

GFP–Plin-expressing constructs and cultured overnight in the presence of oleic acid and cholesterol, plus either BODIPY 558/568 C12 (TAG) or cholesteryl

BODIPY 576/589 C11 (CE). Representative confocal images are shown for each, with red indicating the localization of either TAG or CE droplets, green

indicating localization of GFP–Plin proteins, and yellow (or rings) indicating colocalization. Representative FACS profiles of TAG- and GFP-labeled lipid

droplets or of CE- and GFP-labeled lipid droplets are shown, with relative distributions of total particle numbers indicated in the different quadrants, and the

relative Plin2–5 variant localizations indicated as a percentage of total GFP+ signal that co-sorts (blue) with either TAG+ or CE+ tags. Each experiment was

performed at least three times (see Fig. 6). (A) Plin2 (15,000 total droplets for each population); (B) Plin3 (20,000 total droplets for each population); (C) Plin4

(25,000 total droplets for each population); (D) Plin5 (10,000 total droplets for each population).

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one Plin protein type results in compensation by other Plins

(Martinez-Botas et al., 2000; Tansey et al., 2001; Chang et al.,

2006; Sztalryd et al., 2006). Further, Plin2 and Plin3 are

expressed in most cells and exhibit no TAG or CE targeting

preference. Thus, experiments directed toward defining Plin

effects on TAG or CE levels required us to simultaneously target

multiple Plins in any individual cell. Several cultured cells lines

were selected but we were unsuccessful in depleting any of the

TAG- or CE-LSD specific Plins in combination with Plin2 and

Plin3. As we were unable to deplete all Plins associated with

either TAG- or CE-LSDs, we sought an alternative approach.

When we compared FACS analyses of GFP–Plin1a- and GFP–

Plin1c-expressing cells (Fig. 4), we noticed a conspicuous

difference in the relative distributions of TAG- and CE-LSDs.

More than 60% of total LSD particles in Plin1a-expressing cells

were tagged by the TAG dye marker (Fig. 4A), whereas the

Plin1c-expressing cells were predominantly (.60%) populated

with CE-containing LSDs (Fig. 4C). These data suggested that

the ectopic expression of Plins with specific lipid targeting

preferences might quantitatively alter the balance of TAG/CE

levels in individual cells. We thus analyzed the relative TAG/CE

distributions in cells expressing various Plin proteins.

AML12 cells were transiently transfected individually with

vectors that specifically encode different Plin proteins and

cultured with oleic acid and cholesterol. Cells cultured without

exogenous lipids accumulated only limited levels of TAG or

CE regardless of Plin expression (data not shown), whereas

untransfected control cells accumulate large quantities of both

TAG and CE. LSDs were isolated from the various Plin-

expressing and control cells and their TAG/CE levels quantified

by TLC. Cells expressing Plin1a or Plin1c were analyzed in pairs,and normalized in parallel experiments of lipid loaded and

unloaded control cells (Fig. 7). Similar paired and normalized

experiments were used to analyze cells expressing Plin4 or Plin5.

Plin1a and Plin1c exhibited largely reciprocal effects on TAG/CE

levels, with an ,60% TAG lipid bias observed in Plin1a-expressing

cells and an ,60% CE bias observed for Plin1c-expressing cells

(Fig. 7). Plin5/Plin4 differences in relative TAG/CE accumulationswere similarly polarized toward either TAG or CE, respectively.

Thus, ectopic expression of Plins that exhibit lipid targeting

preferences can polarize relative cellular lipid-type distributions.

These effects are seen for both the TAG-specific (i.e. Plin1a and

Plin5) and the CE-specific (i.e. Plin1c and Plin4) Plin proteins, butnot for the non-preferential Plins 2, 3 and 1d (data not shown).

DiscussionWe have demonstrated that distinct Plins differentially sequester

to either TAG- or CE-specific LSDs, emphasizing diversity of

function for the different Plins. These significant functional

differences towards TAG- or CE-LSDs impact previous

assumptions about commonality of Plin action. The largeunilocular TAG-LSDs that are targeted by Plin1a in adipocytes

are proposed to derive from nascent droplets marked by Plin2,

Plin3 and Plin4 (Wolins et al., 2005). While Plin2 and Plin3

Fig. 6. Relative distributions of Plin family proteins to TAG- or CE-

specific intracellular lipid storage droplets. McARH7777 cells were

transiently transfected with the indicated GFP–Plin-expressing constructs and

cultured overnight in the presence of oleic acid and cholesterol, plus either

BODIPY 558/568 C12 (FA) or cholesteryl BODIPY 576/589 C11 (Chl) dye

markers. TAG or CE distributions are the percentage of the total GFP+ signal

that co-sorts, respectively, with either FA+ or Chl+ tags (see Figs 4, 5). Data

are from at least three independent, paired experiments, where the TAG+CE

sum for each pair was 98–102%. Values are the means 6 standard deviation.

Plin1a, Plin1b and Plin5 show strong preference for localization to TAG-

LSDs, with Plin1a consistently exhibiting a stronger TAG signal than Plin1b

or Plin5. Plin1c and Plin4 show strong preference for localization to CE-

LSDs, with Plin1c consistently exhibiting a stronger CE signal than did Plin4.

Plin1d, Plin2 and Plin3 exhibit less specific localization. Values are 6

standard deviation.

Fig. 7. Relative change in TAG- or CE-specific intracellular lipid storage

upon expression of various Plin proteins. AML12 cells were transiently

transfected with the indicated GFP–Plin-expressing constructs and cultured

overnight in the presence of oleic acid and cholesterol. Transfection

efficiencies were confirmed by visualizing GFP fluorescence. Untransfected

cells, cultured with/without exogenous oleic acid and cholesterol, were grown

in parallel. LSDs were isolated by centrifugation and lipids were extracted,

separated by TLC in parallel with lipid markers, and TAG and CE detected

after staining with iodine vapor. Relative TAG/CE ratios were quantified in

cells transfected with each specific Plin-expressing construct and analyzed in

parallel with identically grown untransfected cells for normalization and TLC

background correction. The TAG/CE ratio for untransfected lipid-loaded cells

(controls) was set to 0.5. The relative TAG/CE-ratio of Plin1a- and Plin1c-

expressing cells were always analyzed in parallel and normalized to those of

control cells, and then secondarily compared to results determined for its Plin-

expressing counterpart. Values .0.5 indicate a proportional increase in TAG

lipid bias, whereas values ,0.5 indicate a proportional increase in CE lipid

bias. Relative distributions of TAG/CE levels are shown as the means 6

standard deviation for each paired comparison. Data for each pair are based

on three independent experiments. Plin4- and Plin5-expressing cells were

similarly analyzed as pairs and internally compared as described for the

Plin1a/Plin1c pair. Plin1a and Plin5 show a relative bias for cellular TAG

accumulation and a strong preference for binding specificity to TAG-LSDs

(see Fig. 6). Conversely, Plin1c and Plin4 show a relative increase in cellular

CE levels and a strong preference for binding to CE-LSDs.

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interact with TAG-LSDs, this association, unlike that of Plin1a, isnot exclusive, as Plin2 and Plin3 also co-segregate with CE-

LSDs. Since Plin4 primarily targets CE-LSDs, conclusionsregarding linear development of Plin-specific LSD populations,based upon static global imaging of cells cultured withexogenous fatty acids (Wolins et al., 2005), must be tempered.

Plin structural domains that direct LSD targeting are stillpoorly defined. All Plins share an N-terminal, ,100 amino acidPAT domain and a distal 11-mer amphipathic helical repeat

(Bussell and Eliezer, 2003; Lu et al., 2001; Miura et al., 2002). C-terminal to both, Plins are increasingly diverged. Several groupshave probed for LSD-interacting motifs through domain-specific

expressions (Garcia et al., 2003; Hickenbottom et al., 2004;McManaman et al., 2003; Nakamura and Fujimoto, 2003; Ohsakiet al., 2006; Subramanian et al., 2004a; Subramanian et al.,2004b; Yamaguchi et al., 2006), but these studies have focused

on TAG-enriched cells and ignored effects of CE-LSDs. Inaddition, some constructs have exposed amphipathic helices thatare usually masked in endogenous Plins and, thus, may target

inappropriately (Hickenbottom et al., 2004).

No simple and consistent structural model for preferential Plinassociations with TG- or CE-LSDs can be easily deduced from

sequence or structural motif scanning and interrogation. Whilethe PAT and associated 11-mer domains may be involved in LSDinteraction, these regions cannot be sufficient determinants forspecific targeting. Although all Plin1 variants have identical N-

terminal 198 residues, which include the PAT and 11-mer regions(Kimmel et al., 2010; Lu et al., 2001; Miura et al., 2002), theyhave dissimilar LSD targeting. Further, the PAT and 11-mer

domains differ greatly among the other Plin proteins in bothlength and sequence. Lipid binding discrimination can also not besimply deduced by analyses of C-termini, which differ highly

among Plin4, Plin5, and each of the unique Plin1 forms.Signaling or targeting motifs may instead reside in non-common segments, as hydrophobic segments are suggested to

facilitate Plin1 targeting to LSDs (Garcia et al., 2003). WhilePlin-specific interacting proteins possibly help direct TAG- orCE-LSD recognition, one must consider that the extremelydiverged single Plin species in Dictyostelium specifically targets

LSDs when expressed in mammalian CHO cells (Lu et al., 2001;Miura et al., 2002). Furthermore, although Plin interactions thatare unique to TAG- or CE-specific LSDs may involve the

surrounding phospholipid monolayer, these would also requirecommonality in both diverse tissues (e.g. adrenal and liver cells)and species.

Pathogenesis associated with abnormal lipid storage hasserious health consequences. Thus, understanding themechanisms that direct lipid storage and lipolytic breakdown isparamount. The Plins regulate access of lipases to lipids stored

within the LSD core (Granneman et al., 2009; Granneman et al.,2011; Martinez-Botas et al., 2000; Miyoshi et al., 2007; Sztalrydet al., 2003; Tansey et al., 2001; Wang et al., 2011; Wang et al.,

2009; Yamaguchi et al., 2004), and the cellular content of Plinsand accumulated LSDs seem intimately coordinated. In addition,various Plins may have significantly different regulatory effects

on cellular lipolytic activity depending upon tissue context(Dalen et al., 2007; Tansey et al., 2003). Accordingly, recentattention has been directed towards possible causal linkages of

aberrant Plin function with human disease. In particular,heterozygous loss-of-function mutations of PLIN1 causes afamilial partial lipodystrophy in humans (Gandotra et al., 2011)

and polymorphisms in human genes for Plin1 (Qi et al., 2004)

and Plin4 (Richardson et al., 2011) have been associated with

obesity.

Our data add novel conceptual parameters for LSD and

Perilipin function and analyses. The various Plins target different

classes of LSDs even within a single cell population and Plins

with LSD lipid specificities can preferentially affect the

accumulation of the targeted LSD class. Intriguingly, tissues

such as adipose, heart and oxidative muscle, that predominantly

accumulate TAG, have highest expression of Plin 1a, 1b or 5,

which specifically target TAG-LSDs. Conversely, steroidogenic

cells, which accumulate CE, express these Plins more poorly in

comparison to Plin1c. We suggest that Plin expression may not

impact all cellular lipids and LSDs equivalently, but that the

actions of individual Plin types may be differentially targeted to

distinct LSD classes within a given cell. It will, thus, be of

interest to evaluate the full spectrum of Plin variants in broad

tissue panels that exhibit differential TAG/CE biases. Our new

findings underscore the view that each Plin is likely to have very

separate and perhaps unique functions associated with their LSD-

specific targeting.

Materials and MethodsMaterials

Fatty-acid-free bovine serum albumin (BSA) was purchased from Fisher Scientific(Pittsburgh, PA). Phosphate-buffered saline (PBS), glutamine, fetal bovine serum(FBS), horse serum, cholesteryl BODIPYH 500/510 FL C12 [cholesteryl 4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene-3-dodecanoate; cholesteryl dye],BODIPYH 558/568 C12 [4,4-difluoro-5-(2-thienyl)-4-bora-3a,4a-diaza-s-indacene-3-dodecanoic acid; fatty acid dye], cholesteryl BODIPYH 576/589 C11 [cholesteryl 4,4-difluoro-5-(2-pyrrolyl)-4-bora-3a,4a-diaza-s-indacene-3-undecanoate; cholesteryl dye],and BODIPYH 500/510 FL C12 [4,4-difluoro-5-methyl-4-bora-3a,4a-diaza-s-indacene-3-dodecanoate; fatty acid dye] were from Invitrogen (Carlsbad, CA). MitoTracker(579/599) and LysoTracker (577/590) were also from Invitrogen (Carlsbad, CA).Antibodies to LAMP1 and EEa1 were from Abcam (Cambridge, MA) and to TOM2was from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). The 250 mm silica gel HTLC plates were from Analtech (Newark, DE). Complete protease inhibitor cocktailtablets were from Roche Diagnostics (Indianapolis, IN). Paraformaldehyde was fromElectron Microscopy Sciences (Hatfield, PA). Cell media and all other chemicals werefrom Sigma-Aldrich (St Louis, MO).

Cell culture

Y1 mouse adrenal-cortical, AML12 mouse liver, and McARH7777 rat hepatomacells were obtained from the American Tissue Culture Collection (Manassas, VA).Cells were grown to subconfluence in medium supplemented with 100 mg/mlpenicillin and 100 mg/ml streptomycin and incubated in humidified air containing5% CO2 at 37 C. Y1 adrenal cells were grown in Dulbecco’s modified Eagle’smedium (DMEM)/Ham’s nutrient mixture F-12 supplemented with 15% horseserum, 2.5% FBS and 2 mM glutamine. McARH7777 cells were grown in DMEMsupplemented with 10% FBS. AML12 cells were grown in DMEM/Ham’s nutrientmixture F-12 supplemented with 5 mg/ml insulin, 5 mg/ml transferrin, 5 ng/mlselenium, 10% FBS and 2 mM glutamine.

Plin–GFP fusion constructs and transfection

The perilipin expression vectors were generated using Multi-Site Gate-Way(Invitrogen, Carlsbad, California). Mouse Plins 1a, 2, 3, and 5 cDNAs wereamplified from previously described pSG5 vectors (Dalen et al., 2007). Plins 1b,1c, 1d and 4 cDNAs were amplified from Y-1 cells or adipose tissue mRNA usingPfuTurboH DNA Polymerase (Stratagene). Primers used contained overhangs forinsertion into the pDONR-221 P4r-P3r vector, Kozak sequence (ACCTAG) andstop codon (CTA), and were designed using Vector NTI 10.0 (Invitrogen) with Tmset to 65 C.

PCR products were recombined into the pDONR-221 P4r-P3r vector using BPClonase II (Invitrogen) to produce pENTR vectors. GFP was amplified frompEGFP-C1 (Clontech) and cloned into the pDONR-221 P1-P4. The V5-6x-His-Glyepitope was cloned into the pDONR-221 P3-P2 vector.

The destination vector was generated by replacing the multi-cloning site ofpcDNA3 (Invitrogen) with the attR1-ccdB-chloramphenicol-attR2 cassette (R1-R2) from pLenti6/v5-DEST (Invitrogen). The R1-R2 cassette was amplified withPfuTurbo, and digested with HindIII/ApaI prior to ligation into HindIII/ApaI-digested pcDNA3 vector. The ligation mixture was transformed into ccdB Survival

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TR cells (Invitrogen) and clones selected on Ampicilin (100 mg/ml) andChloramphenicol (25 mg/ml) plates. The novel pcDNA3-R1-R2 vector(pcDNA3-KTD2-DEST) was subsequently recombined with the above pENTRvectors using LR clonase II (Invitrogen) to generate the pKTD2-G-Perilipin-VHexpression vectors. Due to the stop codon inserted into the pENTR-perilipinvectors, the 3-end V5-6xHis-G tag will not be translated. Correctly amplifiedsequences were confirmed by sequencing (Macrogen, Korea).

Transient transfection was carried out following the manufacturer’s instructions(Invitrogen) using Lipofectamine LTX reagent. McARH7777 cells (cell density6.256104/cm2) were incubated in standard growth conditions in medium lackingantibiotic for one day and then incubated with fresh media containing plasmidDNA (250 ng/cm2), Lipofectamine LTX reagent (1 ml/cm2) and Opti-MEM(50 ml/cm2) for 1 day.

Immunoblot analyses

Proteins were separated by electrophoresis in 10% NuPAGE gels (Invitrogen,Carlsbad, CA) using MOPS running buffer and then subjected to immunoblotanalyses as described (Kim et al., 2002). For immunoblot analyses, we used rabbitpolyclonal antisera (1:3000) to mouse Plin1 (Servetnick et al., 1995), Plin2 (Xuet al., 2005), or Plin3 (Sztalryd et al., 2006), and guinea pig polyclonal antisera(1:3000) to human Plin4 (American Research Product, 03-GP31) and human Plin5(Progen Biotechnik, GmbH GP34). Rabbit antibody (1:5000) to b-actin and GFPwere, respectively, from Abcam (Cambridge, MA) and Invitrogen (Carlsbad, CA).Secondary antibodies to rabbit or guinea pig IgG were from JacksonImmunoResearch (West Grove, PA) and used at 1:5000 dilution.

Lipid loading

Cells were grown overnight to ,50% confluence in media supplemented with100–200 mM oleic acid bound to fatty acid free BSA (2.5:1 mol oleic acid: molBSA) (Dalen et al., 2006) and/or 50 mM cholesterol complexed with b-methylcyclodextrin (8:1 mol b-methyl cyclodextrin: mol cholesterol) (Christian et al.,1997). For fatty acid and cholesteryl dye labeling, cells were grown overnight to,50% confluence in media supplemented with 100 mM oleic acid and 50 mMcholesterol, plus 1.0 mM BODIPYH 558/568 C12 (fatty acid dye) and 0.5 mMcholesteryl BODIPYH 500/510 FL C12 (cholesteryl dye). GFP–Plin expressingcells were grown overnight to ,50% confluence in media supplemented with100 mM oleic acid and 50 mM cholesterol, plus either 1.0 mM BODIPYH 558/568C12 (fatty acid dye) or 0.5 mM cholesteryl BODIPYH 576/589 C11 (cholesteryldye).

Lipid droplet preparation

Cells were grown overnight, washed with PBS, scraped into PBS, and pelleted bycentrifugation (300 g for 5 min). The cell pellet was resuspended in 4 mlhypotonic lysis solution (50 mM HEPES pH 7.3, 0.1 M KCl, 2 mM MgCl2) at4 C, containing protease inhibitors [20 mg/ml leupeptin, 1 mM benzamidine and100 mM 4-(2-aminoethyl)-benzenesulfonylfluoride] and lysed by incubation on icefor 30 min. 1 ml of 50% (w/v) sucrose in lysis solution was added to the cell lysatefor a total volume of 5 ml. This 10% w/v sucrose solution was layered at thebottom of an ultracentrifuge tube and a step gradient of 5%, 2.5% and 0% sucrosewas carefully layered above; centrifugation (Beckman Coulter Optima XL100KUltracentrifuge) was at 154,000 g for 1 h at 4 C. The floating lipid layer wascollected using a Beckman tube slicer (Brea, CA) for immunoblotting or flowcytometry (Brasaemle et al., 2004; Hsieh and Huang, 2005).

FACS separation of fluorescently labeled lipid droplets

Lipid droplets isolated from cells labeled with fluorescent fatty acid dye,fluorescent cholesteryl dye, and/or expressing GFP were sorted using a FACSAriaII cytometer (BD Biosciences). All fluorochromes were excited using a 488 nmlaser. Fluorescent emissions from lipid droplets labeled with cholesteryl BODIPYH500/510 FL C12 or with GFP-tagged perilipin proteins were collected through a502 nm longpass filter followed by a 530/30 nm bandpass filter. Emissions fromlipid droplets labeled with BODIPYH 558/568 C12 or with cholesteryl BODIPYH576/589 C11 were collected through a 556 nm longpass filter followed by a 575/26bandpass filter. Compensation settings for experiments with multiple fluorescentdyes were established using lipid droplets isolated from cells labeled with a singlefluorescent dye (see supplementary material Figs. S1, S3). The forward scatterthreshold was set at the lowest possible linear signal height of 200 (within anallowed range of 200 to 262,143), permitting detection of lipid droplets withdiameters .1 mm. For sorting, ,5000 events/second were processed and sortprecision was set on the default purity mode.

Confocal laser scanning microscopy

Confocal laser scanning microscopy (CLSM) was carried out using a Zeiss LSM510 (Jena, Germany) inverted confocal microscope with a 1006 (Plan-Apochromat, NA1.40) oil objective lens. GFP and cholesteryl BODIPYH FL500/510 FL C12 were imaged using argon 488-nm laser and a 505–530-nm BPemission filter. BODIPYH 558/568 C12 and cholesteryl BODIPYH 576/589 C11

were imaged using a He/Ne 543-nm laser excitation and a 580-nm LP emissionfilter. The software for confocal microscopic image generation was LSM510

software 3.2. The cells were seeded and manipulated in 35 mm glass bottomculture dishes (MatTek, Ashland, MA), fixed with 4% paraformaldehyde in PBSfor 30 minutes, and washed twice for 5 minutes before observation. To visualize

isolated lipid droplets, the unsorted or sorted lipid droplet populations were firstmixed with an equal volume of glycerol before observation.

Lipid extraction and analyses

Lipid droplets were extracted twice with two volumes ofchloroform:heptane:methanol (4:3:2; v/v/v) (Hsieh and Huang, 2007). The lipids

were applied to TLC plates and separated in hexane:diethyl ether:acetic acid(70:30:1; v/v/v); the plates were stained overnight in an iodine chamber to visualize

the lipids. Extracted lipids were separated in parallel to a dilution series of lipidstandards applied to the same plate. The standards were cholesteryl oleate(cholesteryl ester; CE), glyceryl trioleate (triacylglycerol; TAG), oleic acid (fatty

acid; FA), cholesterol (Chol), dioleoylglycerol (diacyclglycerol; DAG), DL-a-monoolein (monoacyclglycerol; MAG), and L-a-phosphatidylcholine (phospholipid;PL).

TIFF images of the iodine stained TLC plates were analyzed by ImageQuant TLwith module 1D gel analyses (GE Health Life Sciences, Piscataway, NJ). Stored 8-bit grayscale images were created manually, since single lane images had various

shapes and spot distances. Spot intensities were corrected for background and dataexported to generate curve standards for the various lipid marker controls. Massesof the unknown lipids were extrapolated in comparison to standard curves (see

supplementary material Fig. S3A).

Statistical analyses

Confocal data (Figs 4, 5, 6) are from at least three independent, paired experiments(see Fig. 3), where the TAG/CE sum for each pair was 98–102% (see Fig. 3).McARH7777 cells were transiently transfected with the indicated GFP-Plin-

expressing constructs and cultured overnight in the presence of 100 mM oleic acidand 50 mM cholesterol, plus either 1 mM BODIPY 558/568 C12 (FA) or 0.5 mMcholesteryl BODIPY 576/589 C11 (Chl) dye markers. TAG or CE distributions

represent the percentage of the total GFP+ signal that co-sorts, respectively, witheither FA+ or Chl+ tags.

FundingThis work was supported by the Intramural Research Program of theNational Institutes of Health; the National Institute of Diabetes andDigestive and Kidney Diseases; and a travel grant from the Henningand Johan Throne-Holst’s Foundation to K.T.D. Deposited in PMCfor release after 12 months.

Supplementary material available online at

http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.104943/-/DC1

ReferencesBrasaemle, D. L., Dolios, G., Shapiro, L. and Wang, R. (2004). Proteomic analysis of

proteins associated with lipid droplets of basal and lipolytically stimulated 3T3-L1

adipocytes. J. Biol. Chem. 279, 46835-46842.

Bussell, R., Jr and Eliezer, D. (2003). A structural and functional role for 11-mer

repeats in alpha-synuclein and other exchangeable lipid binding proteins. J. Mol. Biol.

329, 763-778.

Chang, B. H., Li, L., Paul, A., Taniguchi, S., Nannegari, V., Heird, W. C. and Chan,

L. (2006). Protection against fatty liver but normal adipogenesis in mice lacking

adipose differentiation-related protein. Mol. Cell. Biol. 26, 1063-1076.

Christian, A. E., Haynes, M. P., Phillips, M. C. and Rothblat, G. H. (1997). Use of

cyclodextrins for manipulating cellular cholesterol content. J. Lipid Res. 38, 2264-

2272.

Dalen, K. T., Ulven, S. M., Arntsen, B. M., Solaas, K. and Nebb, H. I. (2006).

PPARalpha activators and fasting induce the expression of adipose differentiation-

related protein in liver. J. Lipid Res. 47, 931-943.

Dalen, K. T., Dahl, T., Holter, E., Arntsen, B., Londos, C., Sztalryd, C. and Nebb, H.

I. (2007). LSDP5 is a PAT protein specifically expressed in fatty acid oxidizing

tissues. Biochim. Biophys. Acta 1771, 210-227.

Farese, R. V., Jr and Walther, T. C. (2009). Lipid droplets finally get a little

R-E-S-P-E-C-T. Cell 139, 855-860.

Gandotra, S., Le Dour, C., Bottomley, W., Cervera, P., Giral, P., Reznik, Y.,

Charpentier, G., Auclair, M., Delepine, M., Barroso, I. et al. (2011). Perilipin

deficiency and autosomal dominant partial lipodystrophy. N. Engl. J. Med. 364, 740-

748.

Garcia, A., Sekowski, A., Subramanian, V. and Brasaemle, D. L. (2003). The central

domain is required to target and anchor perilipin A to lipid droplets. J. Biol. Chem.

278, 625-635.

Perilipin lipid specificities 4075

Journ

alof

Cell

Scie

nce

Granneman, J. G., Moore, H. P., Krishnamoorthy, R. and Rathod, M. (2009).Perilipin controls lipolysis by regulating the interactions of AB-hydrolase containing5 (Abhd5) and adipose triglyceride lipase (Atgl). J. Biol. Chem. 284, 34538-34544.

Granneman, J. G., Moore, H. P., Mottillo, E. P., Zhu, Z. and Zhou, L. (2011).Interactions of perilipin-5 (Plin5) with adipose triglyceride lipase. J. Biol. Chem. 286,5126-5135.

Greenberg, A. S., Egan, J. J., Wek, S. A., Moos, M. C., Jr, Londos, C. and Kimmel,

A. R. (1993). Isolation of cDNAs for perilipins A and B: sequence and expression oflipid droplet-associated proteins of adipocytes. Proc. Natl. Acad. Sci. USA 90, 12035-12039.

Hickenbottom, S. J., Kimmel, A. R., Londos, C. and Hurley, J. H. (2004). Structureof a lipid droplet protein; the PAT family member TIP47. Structure 12, 1199-1207.

Hsieh, K. and Huang, A. H. (2005). Lipid-rich tapetosomes in Brassica tapetum arecomposed of oleosin-coated oil droplets and vesicles, both assembled in and thendetached from the endoplasmic reticulum. Plant J. 43, 889-899.

Hsieh, K. and Huang, A. H. (2007). Tapetosomes in Brassica tapetum accumulateendoplasmic reticulum-derived flavonoids and alkanes for delivery to the pollensurface. Plant Cell 19, 582-596.

Kim, H. U., Hsieh, K., Ratnayake, C. and Huang, A. H. (2002). A novel group ofoleosins is present inside the pollen of Arabidopsis. J. Biol. Chem. 277, 22677-22684.

Kimmel, A. R., Brasaemle, D. L., McAndrews-Hill, M., Sztalryd, C. and Londos, C.

(2010). Adoption of PERILIPIN as a unifying nomenclature for the mammalian PAT-family of intracellular lipid storage droplet proteins. J. Lipid Res. 51, 468-471.

Londos, C., Sztalryd, C., Tansey, J. T. and Kimmel, A. R. (2005). Role of PATproteins in lipid metabolism. Biochimie 87, 45-49.

Lu, X., Gruia-Gray, J., Copeland, N. G., Gilbert, D. J., Jenkins, N. A., Londos, C.

and Kimmel, A. R. (2001). The murine perilipin gene: the lipid droplet-associatedperilipins derive from tissue-specific, mRNA splice variants and define a gene familyof ancient origin. Mamm. Genome 12, 741-749.

Martinez-Botas, J., Anderson, J. B., Tessier, D., Lapillonne, A., Chang, B. H.,Quast, M. J., Gorenstein, D., Chen, K. H. and Chan, L. (2000). Absence ofperilipin results in leanness and reverses obesity in Lepr(db/db) mice. Nat. Genet. 26,474-479.

McManaman, J. L., Zabaronick, W., Schaack, J. and Orlicky, D. J. (2003). Lipiddroplet targeting domains of adipophilin. J. Lipid Res. 44, 668-673.

Miura, S., Gan, J. W., Brzostowski, J., Parisi, M. J., Schultz, C. J., Londos, C.,

Oliver, B. and Kimmel, A. R. (2002). Functional conservation for lipid storagedroplet association among Perilipin, ADRP, and TIP47 (PAT)-related proteins inmammals, Drosophila, and Dictyostelium. J. Biol. Chem. 277, 32253-32257.

Miyoshi, H., Perfield, J. W., 2nd, Souza, S. C., Shen, W. J., Zhang, H. H., Stancheva,Z. S., Kraemer, F. B., Obin, M. S. and Greenberg, A. S. (2007). Control of adiposetriglyceride lipase action by serine 517 of perilipin A globally regulates protein kinaseA-stimulated lipolysis in adipocytes. J. Biol. Chem. 282, 996-1002.

Nakamura, N. and Fujimoto, T. (2003). Adipose differentiation-related protein has twoindependent domains for targeting to lipid droplets. Biochem. Biophys. Res. Commun.

306, 333-338.Ohsaki, Y., Maeda, T., Maeda, M., Tauchi-Sato, K. and Fujimoto, T. (2006).

Recruitment of TIP47 to lipid droplets is controlled by the putative hydrophobic cleft.Biochem. Biophys. Res. Commun. 347, 279-287.

Qi, L., Shen, H., Larson, I., Schaefer, E. J., Greenberg, A. S., Tregouet, D. A.,

Corella, D. and Ordovas, J. M. (2004). Gender-specific association of a perilipingene haplotype with obesity risk in a white population. Obes. Res. 12, 1758-1765.

Richardson, K., Louie-Gao, Q., Arnett, D. K., Parnell, L. D., Lai, C. Q., Davalos, A.,

Fox, C. S., Demissie, S., Cupples, L. A., Fernandez-Hernando, C. et al. (2011).

The PLIN4 variant rs8887 modulates obesity related phenotypes in humans through

creation of a novel miR-522 seed site. PLoS ONE 6, e17944.

Servetnick, D. A., Brasaemle, D. L., Gruia-Gray, J., Kimmel, A. R., Wolff, J. and

Londos, C. (1995). Perilipins are associated with cholesteryl ester droplets in

steroidogenic adrenal cortical and Leydig cells. J. Biol. Chem. 270, 16970-16973.

Subramanian, V., Garcia, A., Sekowski, A. and Brasaemle, D. L. (2004a).

Hydrophobic sequences target and anchor perilipin A to lipid droplets. J. Lipid

Res. 45, 1983-1991.

Subramanian, V., Rothenberg, A., Gomez, C., Cohen, A. W., Garcia, A.,

Bhattacharyya, S., Shapiro, L., Dolios, G., Wang, R., Lisanti, M. P. et al.

(2004b). Perilipin A mediates the reversible binding of CGI-58 to lipid droplets in

3T3-L1 adipocytes. J. Biol. Chem. 279, 42062-42071.

Sztalryd, C., Xu, G., Dorward, H., Tansey, J. T., Contreras, J. A., Kimmel, A. R.

and Londos, C. (2003). Perilipin A is essential for the translocation of hormone-

sensitive lipase during lipolytic activation. J. Cell Biol. 161, 1093-1103.

Sztalryd, C., Bell, M., Lu, X., Mertz, P., Hickenbottom, S., Chang, B. H., Chan, L.,

Kimmel, A. R. and Londos, C. (2006). Functional compensation for adipose

differentiation-related protein (ADFP) by Tip47 in an ADFP null embryonic cell line.

J. Biol. Chem. 281, 34341-34348.

Tansey, J. T., Sztalryd, C., Gruia-Gray, J., Roush, D. L., Zee, J. V., Gavrilova, O.,

Reitman, M. L., Deng, C. X., Li, C., Kimmel, A. R. et al. (2001). Perilipin ablation

results in a lean mouse with aberrant adipocyte lipolysis, enhanced leptin production,

and resistance to diet-induced obesity. Proc. Natl. Acad. Sci. USA 98, 6494-6499.

Tansey, J. T., Huml, A. M., Vogt, R., Davis, K. E., Jones, J. M., Fraser, K. A.,

Brasaemle, D. L., Kimmel, A. R. and Londos, C. (2003). Functional studies on

native and mutated forms of perilipins. A role in protein kinase A-mediated lipolysis

of triacylglycerols. J. Biol. Chem. 278, 8401-8406.

Wang, H., Hu, L., Dalen, K., Dorward, H., Marcinkiewicz, A., Russell, D., Gong, D.,

Londos, C., Yamaguchi, T., Holm, C. et al. (2009). Activation of hormone-sensitive

lipase requires two steps, protein phosphorylation and binding to the PAT-1 domain

of lipid droplet coat proteins. J. Biol. Chem. 284, 32116-32125.

Wang, H., Bell, M., Sreenevasan, U., Hu, H., Liu, J., Dalen, K., Londos, C.,

Yamaguchi, T., Rizzo, M. A., Coleman, R. et al. (2011). Unique regulation of

adipose triglyceride lipase (ATGL) by perilipin 5, a lipid droplet-associated protein. J.

Biol. Chem. 286, 15707-15715.

Wolins, N. E., Quaynor, B. K., Skinner, J. R., Schoenfish, M. J., Tzekov, A. and

Bickel, P. E. (2005). S3-12, Adipophilin, and TIP47 package lipid in adipocytes. J.

Biol. Chem. 280, 19146-19155.

Wolins, N. E., Quaynor, B. K., Skinner, J. R., Tzekov, A., Croce, M. A., Gropler,

M. C., Varma, V., Yao-Borengasser, A., Rasouli, N., Kern, P. A. et al. (2006).

OXPAT/PAT-1 is a PPAR-induced lipid droplet protein that promotes fatty acid

utilization. Diabetes 55, 3418-3428.

Xu, G., Sztalryd, C., Lu, X., Tansey, J. T., Gan, J., Dorward, H., Kimmel, A. R. and

Londos, C. (2005). Post-translational regulation of adipose differentiation-related

protein by the ubiquitin/proteasome pathway. J. Biol. Chem. 280, 42841-42847.

Yamaguchi, T., Omatsu, N., Matsushita, S. and Osumi, T. (2004). CGI-58 interacts

with perilipin and is localized to lipid droplets. Possible involvement of CGI-58

mislocalization in Chanarin-Dorfman syndrome. J. Biol. Chem. 279, 30490-30497.

Yamaguchi, T., Matsushita, S., Motojima, K., Hirose, F. and Osumi, T. (2006).

MLDP, a novel PAT family protein localized to lipid droplets and enriched in the

heart, is regulated by peroxisome proliferator-activated receptor alpha. J. Biol. Chem.

281, 14232-14240.

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