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The Journal of Reproduction and Development logoLink to The Journal of Reproduction and Development
. 2024 Jul 15;70(5):296–302. doi: 10.1262/jrd.2024-023

Perilipin2 depletion causes lipid droplet enlargement in the ovarian corpus luteum in mice

Megumi IBAYASHI 1, Takayuki TATSUMI 2, Satoshi TSUKAMOTO 1
PMCID: PMC11461514  PMID: 39010158

Abstract

Lipid droplets (LDs) are endoplasmic reticulum-derived organelles that store neutral lipids (mostly triglycerides and cholesterol esters) within a phospholipid monolayer and appear in most eukaryotic cells. Perilipins (PLINs, comprising PLIN1–5) are abundant LD-associated proteins with highly variable expression levels among tissues. Although PLINs are expressed in the mammalian ovaries, little is known about their subcellular localization and physiological functions. In this study, we investigated the localization of PLIN1–3 and their relationship with LD synthesis using mCherry-HPos reporter mice, thereby enabling the visualization of LD biogenesis in vivo. PLIN2 and PLIN3 were localized as puncta in granulosa cells with low levels of LD synthesis in developing follicles. This localization pattern was quite different from that of PLIN1, which was mainly localized in the theca and interstitial cells with high levels of LD synthesis. In the corpus luteum, where LD synthesis is highly induced, PLIN2 and PLIN3 are abundant in the particulate structures, whereas PLIN1 is poorly distributed. We also generated global Plin2-deficient mice using the CRSPR/Cas9 system and demonstrated that the lack of PLIN2 did not alter the distribution of PLIN1 and PLIN3 but unexpectedly induced LD enlargement in the corpus luteum. Collectively, our results suggest that the localization of PLIN1–3 is spatiotemporally regulated and that PLIN2 deficiency influences LD mobilization in the corpus luteum within the ovaries.

Keywords: Lipid droplet, Mouse, Ovary, Perilipin


Ovarian follicles develop through the reciprocal stimulation of the oocyte and the surrounding somatic cells, namely, granulosa cells (GCs) and theca cells (TCs) [1, 2]. TCs are located outside the GCs and are separated from each other by the basement membrane. As the follicle develops, proliferating GCs cover the oocyte in multiple layers, and a vascular network is formed around the TCs. During this process, androgens produced by TCs are transported through the basement membrane to the GCs, where they are converted to estradiol through aromatase activity (known as the two-cell/two-gonadotropin theory). After ovulation, the follicle undergoes a morphological change to form the corpus luteum (CL), which functions as a temporary endocrine structure that produces the high levels of progesterone required to support implantation and pregnancy. To maintain the supply of hormones and energy indispensable for follicular growth and CL function, neutral lipids are assumed to be stored as precursors within the cell. Therefore, the ovaries are expected to be lipid-rich.

Lipid droplets (LDs) are organelles in which neutral lipids comprising mainly triglycerides (TGs) and cholesterol esters (CEs) are surrounded by phospholipid monolayers [3, 4]. These organelles are ubiquitous in most eukaryotic cells. The birthplace of LDs is the endoplasmic reticulum (ER) because proteins such as diacylglycerol acyltransferase 1 (DGAT1) and DGAT2, which are involved in neutral lipid synthesis, and Berardinelli-Seip congenital lipodystrophy type 2 (BSCL2, which encodes SEIPIN), which is involved in LD formation, reside in the ER [5,6,7]. As LDs bud from the ER and grow, cytoplasmic proteins, such as perilipins (PLINs), become localized on the surface of LDs, in addition to proteins that migrate from the ER. Recent proteomic studies have identified a wide variety of LD-associated proteins (see the comprehensive open-resource for LD biology dataset, the Lipid Droplet Knowledge Portal [https://lipiddroplet.org/]) [8]. Some of these proteins were unrelated to lipid metabolism, suggesting that LDs have diverse physiological functions beyond their role as lipid depots.

PLINs (PLIN1–PLIN5) are the most abundant LD-associated proteins and are evolutionarily present in metazoans [9]. The characteristics of PLINs have been thoroughly reviewed by Najt et al. [10]. Briefly, PLINs share conserved sequences that mediate LD binding, such as PAT domains (the N-terminus in PLIN1–3 and -5) and 11-mer repeats of different lengths (in all PLINs); however, their expression varies considerably among mammalian tissues. PLIN1 is abundant in white adipose tissue but is also expressed in steroidogenic cells (PLIN1 has several isoforms, but PLIN1a is the major isoform) [11]. In addition, PLIN2 and PLIN3 are widely distributed among tissues [12], PLIN4 is enriched in adipose tissue and differentiating neurons [13], and PLIN5 is predominantly expressed in highly oxidative tissues, such as skeletal muscle, heart, and brown adipose tissues [14, 15]. PLINs participate in lipid homeostasis by regulating basal or stimulated lipolysis in vivo, and various Plin-deficient mice have been reported to exhibit phenotypes associated with metabolic disorders, such as obesity, fatty liver, and insulin resistance [16,17,18,19]. Specifically, Plin2-deficient mice are protected against diet-induced obesity, insulin resistance, and fatty liver disease [17, 20, 21], although these phenotypes differ among mouse models.

Although PLINs are expressed in the mammalian ovaries, including oocytes [22], little is known about their distribution or physiological functions during follicular development. We previously generated mCherry-HPos mice that systemically expressed a fusion protein of red fluorescent protein (mCherry) and HPos, which harbored the minimal sequence required for specific localization to the ER and nascent LDs [23]. Using mCherry-HPos mice, we observed that LD synthesis occurred synchronously with angiogenesis during folliculogenesis [24]. Unexpectedly, we found that PLIN1 was localized to a subset of TCs and interstitial cells (ICs) with active LD synthesis [24]. Our findings, together with the ubiquitous expression of PLIN2 and PLIN3, suggest that various PLINs may localize to the ovaries in a cell type- and stage-dependent manner during follicular development.

In this study, we investigated the localization of PLIN1–3 in the ovaries. Moreover, we generated Plin2-deficient mice using the CRISPR/Cas9 system to investigate whether the loss of PLIN2 alters the localization of PLIN1 and PLIN3 as well as LD synthesis and lipid storage. Our study demonstrated that the localization of PLIN1–3 is spatiotemporally regulated according to LD synthesis and that PLIN2 deficiency induces LD enlargement in the CL within the ovaries.

Materials and Methods

Mice

Generation and genotyping of mCherry-HPos mice have been previously reported [24]. Plin2 knockout mice were generated using CRISPR/Cas9-mediated genomic engineering, as previously described [25], with several modifications. Single-guide RNA (sgRNA) was targeted to exon 3 of Plin2 and was produced via in vitro transcription (MEGAshortscript T7 transcription kit; Thermo Fisher Scientific, Inc., Waltham, MA, USA) using a DNA template with the following sequence: 5′-GCGGCCTCTAATACGACTCACTATAGGGTGGCTCCAGCT TCTGGATGAGTTTTAGAGCTAGAAATAGCAAGTTAAAAT AAGGCTAGTCCGTTATCAACTTGAAAAAGTGGCACCGAG TCGGTGCTTTTTT-3′. This sequence contained a T7 promoter-binding sequence, an sgRNA target sequence (underlined), and a scaffold sequence, as previously reported [26]. Template DNA was synthesized using gBlocks Gene Fragments (Integrated DNA Technologies Inc., Coralville, IA, USA). Synthesized sgRNAs were purified using a MEGAclear Transcription Clean-Up Kit (Thermo Fisher Scientific) and stored at −80°C. The sgRNA (50 ng/μl) and Cas9 protein (30 ng/μl; Nippon Gene Co., Ltd., Tokyo, Japan) were mixed and microinjected into the cytoplasm of one-cell embryos, which were derived from C57BL/6J mice (Japan SLC, Inc., Shizuoka, Japan), to generate Plin2 knockout progeny. F0 founder mice were identified by PCR and sequence analysis, and subsequently mated with wild-type mice to generate F1 offspring. The mutant mice used in this study had a 4 bp deletion in exon 3, resulting in the generation of sequence resistance to the cutting action of the restriction enzyme Van91l (Fig. 2A).

Fig. 2.

Fig. 2.

Generation of Plin2-null mice using CRISPR/Cas9-mediated genome editing. (A) Schematic diagram of the Plin2 gene structure and sgRNA targeting site for the CRISPR/Cas9-mediated generation of Plin2-mutant alleles. Targeting sites of sgRNA are indicated in red. The PAM sequence (NGG) is labeled in green. The four deleted nucleotides (Δ4) are indicated with a red hyphen. Underline shows the recognition sequence for the restriction enzyme (Van91l). ATG, translation initiation site. Bottom: the Plin2 sequence for wild-type and mutant is indicated. (B) Representative genotyping of PCR and subsequent Van91l digestion in Plin2-deficient mice. Bands corresponding to Van91l-digested (369 bp and 132 bp for wild-type [Plin2+/+]) or non-digested (497 bp for mutant [Plin2+/− and Plin2−/−]) Plin2 gene are indicated. (C) Representative immunoblots against PLIN2 and PLIN3 in whole ovarian lysates from hormone (PMSG + hCG)-treated Plin2+/+, Plin2+/−, and Plin2−/− mice. Tubulin was used as a loading control. (D) Representative immunoblots against PLIN2 in whole ovarian lysates from Plin2+/+, Plin2+/−, and Plin2−/− mice treated with or without PMSG + hCG. Actin was used as a loading control. (E) Representative confocal images of frozen ovarian sections from Plin2+/+ and Plin2−/− mice immunolabeled with anti-PLIN2 antibody. Cell nuclei were counterstained with DRAQ5. Ovarian follicles are outlined by dashed lines. Bottom panels show enlarged images of the boxed area. Numbers indicate the follicle stage. Arrows, PLIN2-puncta in Plin2+/+ GC. Scale bars, 50 μm and 10 μm (inset).

For genotyping, PCR was performed using Ex Premier DNA Polymerase (Takara Bio Inc., Kusatsu, Japan) with P1 (5′-GGACAGGAATTGGTCCTCAA-3′) and P2 (5′-TGGGTAGAGGAGGGATTGTG-3′) primers, which amplified a wild-type band of 501 bp and a mutant band of 497 bp (Fig. 2B; see PCR products). These amplicons were further digested with Van91l, which generated either wild-type-specific bands of 369 bp and 132 bp or a 497 bp mutant-specific band (Fig. 2B; see Van91l cleaved).

To stimulate follicular development, female mice were intraperitoneally injected with 5 IU pregnant mare serum gonadotropin (PMSG; Aska Animal Health, Tokyo, Japan). After 46–48 h, the ovaries were harvested, and frozen sections were prepared. To induce CL formation, female mice injected intraperitoneally with 5 IU of PMSG 48 h beforehand were intraperitoneally injected with 5 IU of human chorionic gonadotropin (hCG; Aska Pharmaceutical, Tokyo, Japan), and their ovaries were collected 46–48 h later.

All mice used in this study were 8–12 weeks old, except for the 3–4-week-old mice shown in Fig. 1 (preantral and antral follicles) and Fig. 2E, which were maintained on a C57BL/6J background. The mice were housed under a stable (12-h) light/dark cycle with free access to water and rodent chow. All the mouse experiments were approved and registered by the Animal Care and Use Committee of the National Institutes for Quantum Science and Technology (approval number: 16-1012-5).

Fig. 1.

Fig. 1.

Distribution of PLIN1, 2, and 3 in mouse ovaries. Representative confocal images of frozen ovarian sections from PMSG- or hCG-treated mCherHPos mice immunolabeled with antibodies against PLIN1 (A), PLIN2 (B), or PLIN3 (C). (D) Schematic illustration of the characteristics of mCherry-HPos, which accumulates in nascent to growing LDs synthesized in the ER and allows visualization of LD synthesis. Cell nuclei were counterstained with DRAQ5. The image on the right of each panel is a higher-magnification image of the boxed region in the left image. GC, granulosa cells; TC, theca cells; IC, interstitial cells; CL, corpus luteum. The ovarian follicles and CL are outlined with dashed lines. Numbers indicate the follicle stage (see Materials and Methods for details). Arrows, PLIN2- or PLIN3-puncta in the GC; arrowheads, PLIN2- or PLIN3-granules in the CL. Scale bars, 50 μm and 10 μm (inset).

Imaging of frozen ovarian sections

Immunofluorescence analysis of frozen ovarian sections was conducted as previously described [25, 27]. PLINs were visualized with anti-PLIN1 antibody (1:50; #9349, Cell Signaling Technology, Beverly, MA, USA), anti-PLIN2 antibody (rabbit anti-PLIN2 [1:50; NB110-40877, Novus Biologicals, Centennial, CO, USA], or chicken anti-PLIN2 [1:50; ab37516, Abcam, Cambridge, UK], as shown in Supplementary Fig. 1) or anti-PLIN3 antibody (1:50; NB110-40764; Novus Biologicals), followed by Alexa Fluor 488-conjugated goat anti-rabbit IgG or chicken IgY (1:500; A-11008 or A-11039, Thermo Fisher Scientific). The sections ware counterstained with DRAQ5 (10 µg/ml; DR50050, BioStatus Ltd., Shepshed, UK) for 20–30 min. Finally, the sections were mounted on glass slides using ProLong Glass Antifade Mountant (P36980, Thermo Fisher Scientific) on glass slides. When necessary, frozen ovarian sections were stained with LipidTox Deep Red (100× dilution in PBS; H34477, Thermo Fisher Scientific) for 20–30 min before mounting.

Confocal images were captured under a 60× oil immersion objective mounted on an IX83 inverted microscope (Olympus, Tokyo, Japan) equipped with a spinning disk confocal system, as previously described [28]. The signal intensities were optimized for each channel with a representative section and the images were captured in a single plane. Confocal images were thresholded, and particles were detected using the Analyze Particles tool using Fiji [29]. The threshold was adjusted for representative images and identical settings were used for all images from the same experiment. For quantification, 5–10 different images per section were averaged from the same ovary (n > 3 mice in each group). Follicular development was classified according to the scheme of Pedersen & Peters [30] by determining granulosa cell numbers: type 3a, less than 20 granulosa cells; type 3b, 21–60 granulosa cells; type 4, 61–100 granulosa cells; type 5a, 101–200 granulosa cells; type 5b, 201–400 granulosa cells; type 6, 401–600 granulosa cells; type 7, > 600 granulosa cells; and type 8 (preovulatory follicle), large follicles with a single cavity with follicular fluid. Quantification of mCherry-HPos foci in frozen ovarian sections was performed as described previously [24].

Oil Red O staining

Oil Red O staining was performed as previously described [27]. After staining, the sections were visualized using a 40× objective lens (UPIanSApo ×40/0.95 NA, Olympus) mounted on a microscope (IX83, Olympus) equipped with a digital camera (DP74, Olympus).

Immunoblotting

Immunoblotting was performed as previously described [25]. Briefly, ovaries collected from female mice treated with or without hormones (PMSG + hCG) were homogenized using glass beads in lysis buffer (×10 dilution in PBS; #9803, Cell Signaling Technology) containing a complete protease inhibitor cocktail (#5871; Cell Signaling Technology). Following centrifugation at 10,000 g, the supernatants were mixed with 4× Laemmli buffer, separated on SuperSep™ Ace 10%–20% gels (Fujifilm Wako Pure Chemical Corporation, Osaka, Japan), and transferred to PVDF membranes using a Turbo Transfer System (Bio-Rad Laboratories, Hercules, CA, USA). Membranes were blocked with Blocking One (03953-95, Nacalai Tesque, Kyoto, Japan) for 1 h, incubated overnight at 4°C with anti-PLIN2 antibody (1:1,000; NB110-40877; Novus Biologicals) or anti-PLIN3 antibody (1:1,000; NB110-40764, Novus Biologicals), and then incubated with horseradish peroxidase (HRP)-linked anti-rabbit IgG (1:10,000; #7074, Cell Signaling Technology). To detect tubulin and actin, HRP-conjugated anti-β-tubulin (1:1,000; #5346, Cell Signaling Technology) or anti-β-actin (1:1,000; A00730, GenScript, NJ, USA) antibodies were used, respectively. The membranes were visualized using SuperSignal West Femto Maximum Sensitivity Substrate (34094, Thermo Fisher Scientific) and analyzed using a ChemiDoc-It imaging system equipped with a BioChemi camera (UVP, Upland, CA, USA).

Statistical analysis

Statistical analyses were performed using a two-tailed Student’s t-test using GraphPad Prism 6 (GraphPad Software, Boston, MA, USA). Data represent means ± SEM.

Results

Distribution of PLIN1–3 in ovaries

Our previous study using mCherry-HPos mice demonstrated high LD synthesis in TCs and ICs, and low LD synthesis in GCs during follicular development [24]. To investigate the possible link between LD synthesis and PLIN (PLIN1–3) localization during follicular development, frozen ovarian sections from PMSG-primed mCherry-HPos mice were immunostained with antibodies against endogenous PLIN1, PLIN2, and PLIN3. In agreement with our previous observations, PLIN1 was preferentially distributed to TCs and ICs, where mCherry-HPos foci were abundant (indicating high levels of LD synthesis), during follicular development (Fig. 1A; see also the schematic representation of mCherry-HPos in Fig. 1D). In contrast to PLIN1, PLIN2 and PLIN3 were localized to defined regions and appeared as puncta in GCs within growing follicles where mCherry-HPos foci were undetectable (indicating low levels of LD synthesis; arrows in Figs. 1B and C, preantral follicles). However, they were rarely visible in the TCs and ICs, where mCherry-HPos foci were abundant (Figs. 1B and C). This distribution was also observed in the late stages of follicular development (Figs. 1B and C, antral follicles). In the CL of hCG-primed mCherry-HPos mice (during the luteal phase), PLIN1 was less abundant within the CL but was mainly distributed in the TC layer (Fig. 1A, corpus luteum), whereas PLIN2 and PLIN3 appeared to have granular structures, some of which were close to the mCherry-HPos foci (arrowheads in Fig. 1B and C, corpus luteum).

Taken together, these results indicate that the localization of PLIN2 and PLIN3 is closely linked but quite different from that of PLIN1 during follicular development. This result also suggests that PLIN1–3 localization changes in response to LD synthesis in the ovaries.

Generation of Plin2-deficient mice

To determine whether the loss of PLIN2 influences the distribution of PLIN1 and PLIN3, as well as ovarian function, we generated Plin2-deficient mice via CRISPR/Cas9-mediated genome engineering. To this end, we designed sgRNAs targeting exon 3 of Plin2, which generated mice with a 4 bp indel, resulting in the introduction of a premature stop codon into exon 4 (Fig. 2A). The deletion generated sequence resistance to the cutting action of the restriction enzyme Van91l, facilitating the genotyping of the enzyme (Fig. 2B, see also Materials & Methods). Intercrossing of Plin2 heterozygous mice (Plin2+/−) provided healthy homozygous mice (Plin2−/−) at the expected Mendelian ratio (13:30:11 for Plin2+/+:Plin2+/−:Plin2−/−). Mating of sexually mature Plin2−/− females with wild-type males produced pups at the same ratio as Plin2+/− or wild-type females (data not shown), suggesting that global Plin2 deficiency does not have a significant impact on female fertility. This is consistent with previous studies showing that Plin2−/− mice are grossly normal when fed a regular diet [16, 17].

Western blot analysis revealed the complete absence of PLIN2 from the ovaries of these Plin2−/− mice, in contrast to Plin2+/+ or Plin2+/−mice (Fig. 2C). Previous studies have shown that PLIN3 expression is upregulated in the embryonic fibroblasts and adrenal glands of Plin2-null mice [31, 32] and Plin2-deficient livers [33]. However, no marked upregulation of PLIN3 was detected in Plin2−/− ovaries (Fig. 2C). Additionally, we confirmed that hormone treatment did not alter the expression level of PLIN2 in the Plin2+/+ or Plin2+/− ovaries (Fig. 2D).

To further verify the specificity of the immunostaining shown in Fig. 1B, frozen ovarian sections from Plin2−/− mice were stained with an anti-PLIN2 antibody. We found that PLIN2, which was visualized as puncta in GCs in Plin2+/+ ovaries, was completely absent from Plin2−/− ovaries (Fig. 2E). Similar results were obtained when the ovarian sections from Plin2−/− mice were labeled with another anti-PLIN2 antibody (Supplementary Fig. 1). These results indicate that our immunostaining method selectively labeled endogenous PLIN2.

LD synthesis is unchanged in Plin2-deficient ovaries

To determine whether the lack of Plin2 influences LD synthesis in the ovaries, frozen ovarian sections obtained from Plin2+/− or Plin2−/− mice expressing transgenic mCherry-HPos (referred to as Plin2+/−;mCherry−HPos or Plin2−/−;mCherry−HPos, respectively) were observed under fluorescence microscopy. Despite the absence of PLIN2, numerous mCherry-HPos foci were detectable in the TCs and ICs of Plin2−/−;mCherry−HPos ovaries, similar to Plin2+/−;mCherry−HPos ovaries (arrows in Fig. 3, preantral and antral follicles). No difference was detected in the CL between the two groups (arrows in Fig. 3, corpus luteum). These results indicated that LDs are normally synthesized in ovaries lacking PLIN2.

Fig. 3.

Fig. 3.

LD synthesis in Plin2-deficient ovaries. Representative images of frozen ovarian sections from PMSG (for preantral and antral follicles) or hCG (for CLs) treated Plin2+/− or Plin2−/−mice systemically expressing mCherry-HPos (indicated by Plin2+/−;mCherry−HPos or Plin2−/−;mCherry−HPos, respectively). The insets show enlarged images of the boxed areas. Numbers indicate follicle stage. Arrows indicate mCherry-HPos foci representing nascent LDs. Scale bars: 50 μm and 5 μm (inset).

LD enlargement in the CL of Plin2-deficient ovaries

PLIN2 acts as an LD coat protein to protect LDs from neutral lipases (i.e., adipose triglyceride lipase [ATGL]) or autophagic-mediated pathway [34,35,36,37]. Hence, a lack of PLIN2 may reduce the levels of LDs and neutral lipids stored within cells. To test this possibility, we stained Plin2+/− or Plin2−/− ovarian sections with LipidTox Deep Red, which specifically labels neutral lipids. We noticed that a small number of LipidTox Deep Red-positive foci were detected in GCs from the Plin2+/− or Plin2−/− ovaries (Fig. 4A, preantral to antral follicles); however, unexpectedly, in the CL of Plin2−/− ovaries, the larger-sized LDs labeled with LipidTox Deep Red were increased compared with those of Plin2+/− ovaries (arrows in Fig. 4A, corpus luteum). Supporting this finding, quantitative analysis revealed that Plin2-deficiency induced a 1.4-fold increase in LD size, whereas LD number was unchanged in the CL (Fig. 4B). Consistent with this increased in LD size in the CL, Oil Red O staining showed that a lack of PLIN2 caused marked LD accumulation in the CL of Plin2−/− ovaries compared with Plin2+/+ ovaries (Fig. 4C). Taken together, these results suggest that the lack of Plin2 causes LD enlargement in the CL.

Fig. 4.

Fig. 4.

LD enlargement in Plin2-deficient ovaries. (A) Representative confocal images of frozen ovarian sections from PMSG- or hCG-treated Plin2+/− and Plin2−/− mice that were immunolabeled with anti-PLIN2 antibody and stained with LipidTox Deep Red to visualize neutral lipids. Lower panels show magnified images of the boxed regions in the upper panel. Dashed lines indicate GC–TC border or CL boundary. Numbers indicate the follicle stage. Arrows (in CL) indicate enlarged LDs labeled with LipidTox Deep Red. Scale bars, 50 μm and 10 μm (inset). (B) Quantification of the average LD size and LD number based on the confocal images shown in A. Data are presented as means ± SEM (n = 10 images from 3–5 mice analyzed per genotype). ** P < 0.01; n.s., not significant. (C) Representative CL images of frozen ovarian sections from hCG-treated Plin2+/+ and Plin2−/− mice stained with Oil Red O to visualize neutral lipids. Cell nuclei were counterstained with Mayer’s hematoxylin (blue). Scale bars, 20 μm.

Discussion

Although the expression of PLINs varies among mouse tissues, this is the first study to demonstrate the spatiotemporal regulation of PLIN1–3 localization in ovaries. Notably, PLIN2 and PLIN3 were found in puncta in restricted regions of GCs in growing follicles. Because GCs have low levels of LD synthesis and hence store low amounts of neutral lipids (i.e., few LDs), PLIN2- and PLIN3-positive puncta are structures that may be unrelated to LD metabolism. Our previous studies demonstrated that LD synthesis is rapidly induced when GCs are cultured in vitro, and under these conditions, PLIN2 and PLIN3 are distributed granularly throughout the cytoplasm ([24] and our unpublished data). Therefore, in GCs with low LD synthesis, PLIN2 and PLIN3 may be sequestered to certain cellular compartments. Because PLIN2, when not bound to LDs in the cytoplasm, is unstable and rapidly degraded via the ubiquitin-proteasome or autophagy-mediated pathway [37,38,39,40], such compartments may also serve to protect PLIN2 itself against such degradation.

In contrast to the GCs, PLIN2 and PLIN3 were detected as granular structures in the CL, which showed high levels of LD synthesis and abundant LDs. However, the distribution of PLIN1 was quite different from that of PLIN2 and PLIN3, with PLIN1 mainly localized to some TCs and ICs. Further studies are needed to determine the possible significance of these differential distributions in the ovaries, even though all proteins examined belong to the same PLIN family.

Because PLIN2 protects LDs from neutral lipases such as ATGL, as well as autophagy, PLIN2 deletion should reduce LD content, as reported in the Plin2-deficinet liver [41]; however, unexpectedly, a lack of PLIN2 did not alter LD synthesis, but rather caused LD enlargement in the CL. Why do Plin2-deficient ovaries exhibit enlarged LDs in the CL? If PLIN3 compensates for the PLIN2 function (i.e., PLIN3 coats LDs instead of PLIN2), PLIN3 may be more resistant to the cytoplasmic degradation pathway (e.g., lipolysis and/or autophagy) than PLIN2, resulting in LD enlargement. Further studies are required to clarify whether this increase in LD size is due to fusion between LDs or an increase in the amount of neutral lipids that they possess.

Li et al. reported that Plin2 deficiency induces adrenal gland enlargement and causes CE-rich LD accumulation in the adrenal cortex [32]. They also revealed that LD accumulation accelerates with age [32]. It is worth investigating whether aging contributes to LD enlargement, leading to reduced fertility in Plin2-deficient mice with increasing maternal age.

Lack of Plin2 causes LD enlargement in the CL, but no marked changes in ovarian functions, such as follicular development, ovulation, and pregnancy (although fertility declines with increasing maternal age; unpublished data). Why was no apparent phenotype observed in the ovaries of Plin2-deficient mice? Given the closely related distribution patterns of PLIN2 and PLIN3 during follicular development, PLIN3 may compensate for the PLIN2 deficiency. However, the expression level of PLIN3 was unchanged in Plin2-deficient ovaries, whereas PLIN3 was upregulated in Plin2-deficient mouse embryonic fibroblasts, adrenal glands, and the liver [31,32,33], indicating that the regulatory mechanisms involved in PLIN2 and PLIN3 compensation may depend on the cell type and tissue. Additionally, PLIN2 may not be essential for ovarian function (particularly in younger age). To determine whether PLIN2 is indeed required for ovarian function, it is necessary to generate mice that are simultaneously deficient not only in Plin2 but also in other Plins, and to investigate their ovarian function. Furthermore, it may be useful to generate conditional knockout mice specifically deficient in PLIN2 in granulosa and luteal cells, in which PLIN2 is abundant in the ovary.

Given that the localization of PLIN1–3 depends on the ovarian cell type, further studies focusing on the remaining PLINs (PLIN4 and PLIN5) are critical to obtain a complete overview of the physiological functions of PLINs within the ovary. Notably, PLIN5 is highly expressed in oxidative tissues such as heart muscle and brown adipose tissues and plays a role in tethering LDs to mitochondria to promote fatty acid metabolism (i.e., β-oxidation) [14, 15, 42, 43]. Our previous electron microscopy analysis revealed that mitochondria are distributed in a bead-like pattern in the proximity of LDs, which are abundant in the CL; however, LDs are rarely found around mitochondria distributed in GCs in growing follicles [24]. Further research should investigate whether PLIN5 is involved in the distribution of LDs according to the mitochondrial content.

Given that PLINs, which are evolutionarily conserved LD surface proteins, have been recognized as physiological regulators of lipid accumulation in many tissues [10], addressing the roles of PLINs is important for understanding the molecular basis underlying ovarian LD biology and ovarian dysfunction with metabolic disorders.

Conflict of interests

The authors declare no competing or financial interests.

Supplementary

Supplement Figure
jrd-70-296-s001.pdf (1.4MB, pdf)

Acknowledgments

We thank Mami Hayashi for the technical assistance. This study was supported by a Japan Society for the Promotion of Science KAKENHI grant (22H03230 to S.T.).

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