Regulation of meiotic telomere dynamics through membrane fluidity promoted by AdipoR2-ELOVL2

Zhang, Jingjing; Ruiz, Mario; Bergh, Per-Olof; Henricsson, Marcus; Stojanović, Nena; Devkota, Ranjan; Henn, Marius; Bohlooly-Y, Mohammad; Hernández-Hernández, Abrahan; Alsheimer, Manfred; Borén, Jan; Pilon, Marc; Shibuya, Hiroki

doi:10.1038/s41467-024-46718-6

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Published: 14 March 2024

Regulation of meiotic telomere dynamics through membrane fluidity promoted by AdipoR2-ELOVL2

Nature Communications volume 15, Article number: 2315 (2024) Cite this article

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Abstract

The cellular membrane in male meiotic germ cells contains a unique class of phospholipids and sphingolipids that is required for male reproduction. Here, we show that a conserved membrane fluidity sensor, AdipoR2, regulates the meiosis-specific lipidome in mouse testes by promoting the synthesis of sphingolipids containing very-long-chain polyunsaturated fatty acids (VLC-PUFAs). AdipoR2 upregulates the expression of a fatty acid elongase, ELOVL2, both transcriptionally and post-transcriptionally, to synthesize VLC-PUFA. The depletion of VLC-PUFAs and subsequent accumulation of palmitic acid in AdipoR2 knockout testes stiffens the cellular membrane and causes the invagination of the nuclear envelope. This condition impairs the nuclear peripheral distribution of meiotic telomeres, leading to errors in homologous synapsis and recombination. Further, the stiffened membrane impairs the formation of intercellular bridges and the germ cell syncytium, which disrupts the orderly arrangement of cell types within the seminiferous tubules. According to our findings we propose a framework in which the highly-fluid membrane microenvironment shaped by AdipoR2-ELOVL2 underpins meiosis-specific chromosome dynamics in testes.

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Introduction

During mitosis, the cytoplasm of the dividing cell forms a temporal bridging structure called the midbody¹. Midbody formation in mitotic cells is regulated not only by a series of midbody-localizing proteins², but also by the local rearrangement of the lipid membrane that enriches classical sphingolipids as well as their glycosylated variants (glycosphingolipids) (as summarized in Supplementary Fig. 1)³. The inhibition of glycosphingolipid biosynthesis leads to defective cytokinesis⁴, suggesting that the rearrangement of the lipid membrane is crucial for faithful cytokinesis.

While the mitotic midbody is a temporal structure, dividing germ cells do not complete cytokinesis and instead form stable intercellular bridge structures resulting in the formation of germline syncytia⁵. In mammalian germ cells, the intercellular bridge is composed of canonical midbody proteins in addition to a germ cell-specific component called Testis Expressed 14 (TEX14). TEX14 stabilizes the temporal midbody structure and forms the stable intercellular bridge by inhibiting the recruitment of proteins essential for cell abscission (Supplementary Fig. 1a)⁶. The deletion of Tex14 and the subsequent disruption of the intercellular bridge results in the death of prophase I spermatocytes and male infertility in mice⁷. In contrast, female mice with a deletion of Tex14 exhibit a disruption of intercellular bridge structures while maintaining fertility, suggesting the sexually dimorphic requirement of the germline syncytium⁸.

Accumulating evidence suggests that the formation of the intercellular bridge in mammalian germ cells requires specialized forms of sphingolipids that contain very-long-chain polyunsaturated fatty acids (VLC-PUFAs) (Supplementary Fig. 1b)^{9,10,11,12,13}. VLC-PUFAs are not present in ordinary dietary sources and thus need to be synthesized in situ. VLC-PUFAs are uniquely synthesized in the brain, retina, and testis and have specialized roles in these tissues such as brain physiology, photoreceptor activities, and male reproduction, respectively¹⁴. In the testis, the synthesis of VLC-PUFA-containing sphingolipids requires the germ cell-specific ceramide synthase CerS3 as well as the fatty acid (FA) elongase ELOVL2^9,10. CerS3 deficiency results in the disorganization of the intercellular bridge and thus the death of prophase I spermatocytes, similar to Tex14 knockout (KO) mice. Notably, TEX14 still localizes to the disorganized intercellular bridge in CerS3 KO spermatocytes, suggesting that lipid rearrangement is essential for intercellular bridge stability in addition to the TEX14-dependent regulation of midbody proteins. However, the detailed molecular regulations underlying the VLC-PUFA synthesis in the testis and, more crucially, which aspects of meiosis require VLC-PUFAs are poorly understood.

During meiotic prophase I, homologous chromosomes are paired and recombined¹⁵. To achieve the homology search, chromosomes attach to the nuclear envelope (NE) via their telomeres and move along the NE^16,17. The misregulation of telomere-driven chromosome movements leads to failures in pairing and recombination and to human infertility, including azoospermia and primary ovarian insufficiency^18,19,20,21. Even though the responsible molecular regulations of meiotic telomere-binding proteins have been clarified, how the membrane is regulated for this purpose remains unexplained. Telomeres are mechanically fused with the lipid bilayer and subsequently move along the membrane²², thus there must be a need for the rearrangement of lipid species to fluidize the membrane.

Here, we report that the highly-fluid lipid environment ensured by the synthesis of VLC-PUFA in male germ cells is required for the nuclear peripheral distribution of meiotic telomeres and thus for faithful homologous recombination and synapsis. The synthesis of VLC-PUFA in testes depends on AdipoR2 (Adiponectin receptor 2), an evolutionarily conserved regulator of membrane homeostasis^23,24,25,26. AdipoR2 ensures the robust expression of ELOVL2 in the testis. Our findings establish an essential role for AdipoR2-ELOVL2 in shaping the highly-fluid membrane microenvironment, thus allowing the chromosome dynamics needed for proper meiosis progression in male germlines.

Results

AdipoR2 is essential for male reproduction

AdipoR2, as well as its paralog AdipoR1, were initially identified as the receptors for Adiponectin, a secretory protein produced by adipocytes²³; however, recent studies have demonstrated their prime role as evolutionarily conserved regulators of membrane homeostasis^24,25,26. To analyze their tissue-specific roles, we used reverse transcription PCR (RT-PCR) to assess mRNA expressions in murine tissue samples. AdipoR1 and AdipoR2 were widely expressed in somatic tissues, while AdipoR2 alone demonstrated predominant expression in the testis (Fig. 1a).

**Fig. 1: AdipoR2 is indispensable for male reproduction.**

To investigate AdipoR2’s roles in the testis, we analyzed spermatogenesis defects in AdipoR2 KO testes (Fig. 1b). The KO testes were significantly smaller compared to those of their WT littermates (Fig. 1c). Histological staining showed that there were only a small number of post-meiotic round spermatids in the KO seminiferous tubules and that elongated spermatids were completely absent (Supplementary Fig. 2). Of note, the round spermatids in the KO testis were abnormally large and multinucleated, and these cells were undergoing apoptosis (Fig. 1d, e), which is similar to the defects reported in lipid metabolism-defective mutants such as CerS3 KO and Elovl2 KO testes^9,10. Consistent with the histological analysis, transmission electron microscopy (TEM) analysis found that multiple round spermatid nuclei lacking the discernable acrosome structure resided in a large cytoplasmic area enclosed by a plasma membrane (Fig. 1f). Consistent with the testicular defects, the epididymides from AdipoR2 KO males were devoid of sperm (Fig. 1g), which led to the complete loss of male fertility (Fig. 1h).

AdipoR2 deficiency causes homologous synapsis and recombination failures

The TUNEL assay showed that a large number of germ cells underwent apoptosis in mid-pachytene spermatocytes at epithelial cycle stage V–VIII (Fig. 2a). To investigate meiotic defects further, we analyzed the progression of meiotic prophase I by staining with SYCP1 (a homologous synapsis marker) and SYCP3 (an axial element marker) (Supplementary Fig. 3a). Quantification revealed a significant increase in leptotene (unsynapsed) and zygotene (partially synapsed) populations in the AdipoR2 KO testis (Fig. 2b). Closer inspection of AdipoR2 KO pachytene spermatocytes showed the presence of two major synapsis defects—unsynapsis (represented by a few chromosomes remaining unsynapsed along the whole length of their chromosome axes even when the other chromosomes have completely synapsed) or incomplete synapsis (represented by abnormal Y-shaped branching structures) (Fig. 2c).

**Fig. 2: AdipoR2 ensures the synapsis and recombination of homologous chromosomes.**

During meiosis, unsynapsed chromatin activates a response termed meiotic silencing of unsynapsed chromatin (MSUC) and leads to the accumulation of DNA-damage markers such as the phosphorylated serine 139 of histone H2AX (γH2AX)^27,28. In WT pachytene cells, the γH2AX signal was restricted to sex chromosomes and formed one flare signal on the condensed sex chromosomes (Fig. 2d). In contrast, there was a significant increase in the number of γH2AX flares in the AdipoR2 KO pachytene spermatocytes. The chromatin near the branching axis was stained with γH2AX, suggesting that MSUC was activated at the incompletely synapsed regions (Fig. 2d, magnified pictures). The RAD51 recombinase was not detected at the γH2AX flares in the AdipoR2 KO pachytene spermatocytes, suggesting that it is not unrepaired DNA double-strand breaks (DSBs) but MSUC that is responsible for γH2AX flare formation (Supplementary Fig. 3b). Further, thickened and condensed SCYP3-stained chromosome axes seen on the sex chromosomes (the so-called “sex body”) in WT spermatocytes were barely distinguishable in the AdipoR2 KO pachytene spermatocytes (Fig. 2d, rightmost panel), suggesting that the activation of MSUC led to imperfect silencing and to the decondensation of sex chromosomes.

Meiotic homologous recombination is facilitated by homologous synapsis, thus failures in homologous synapsis frequently coincide with defects in homologous recombination^29,30. The counting of RPA2 foci, a single-strand DNA-binding protein that decorates the DSB sites, and the type I crossover marker MutL homolog 1 (MLH1) in AdipoR2 KO spermatocytes, revealed defects in DSB repair and crossover formation (Fig. 2e, f). Taken together, AdipoR2 depletion significantly impairs homologous chromosome synapsis and recombination, which causes aberrant activation of MSUC on autosomes leading to the massive, if not complete, elimination of male germ cells at the mid-pachytene stage.

AdipoR2 is essential for the synthesis of testis-specific VLC–PUFA-containing sphingolipids

To gain insight into the mechanism underlying the meiotic errors, we performed a whole-testis lipidomics analysis to determine whether the FA composition is changed in AdipoR2 KO testis. To minimize the heterogeneity in the cell population between WT and AdipoR2 KO testes, we used juvenile testis samples at postnatal day 28. We determined the composition of two dominant phospholipid classes, phosphatidylcholine (PC) and phosphatidylethanolamine (PE), as well as of the sphingolipid ceramide (Fig. 3). Consistent with previous studies^9,10, the WT testes samples were enriched in PC, PE, and ceramides containing VLC-PUFAs, which is a unique feature of the testis lipidome (Fig. 3a). However, the AdipoR2 KO testes showed significant accumulation of FAs containing shorter carbon chains such as PC 32:0, PC 34:1, PE 34:1, and ceramide 16:0 (Cer 16:0) and a significant reduction of VLC-PUFAs such as PC 36:4, PC 38:5, PC 38:6, PE 36:4, ceramide 30:6, and ceramide 30:5 (Fig. 3a). The most significantly affected lipid species was Cer 16:0, a sphingosine anchored with a palmitic acid, which was 22 mol % more overrepresented in AdipoR2 KO testes compared to WT (Fig. 3b). Similar trends were detected even when the lipids were quantified in terms of nmol or pmol/mg tissue (Supplementary Fig. 4). The heat map visualization showed a clear gradient where the saturated or monounsaturated FAs with shorter carbon chains were overrepresented and, conversely, the VLC-PUFAs were mostly depleted in the AdipoR2 KO samples (Fig. 3c). The analysis of another sphingolipid species, sphingomyelin, showed the same trends (Supplementary Fig. 5). These results suggest that AdipoR2 is essential for the synthesis of testis-specific VLC-PUFA–containing phospholipids and sphingolipids.

**Fig. 3: The depletion of VLC-PUFA in the *AdipoR2*^–/– testis lipidome.**

AdipoR2 regulates ELOVL2 at both the transcriptional and post-transcriptional levels

VLC-PUFA–containing ceramide is synthesized through multiple steps, including FA desaturation by desaturases (such as FADS1, FADS2, and SCD1), FA elongation by elongases (such as ELOVL2, ELOVL4, and ELOVL5), and the transfer of the acyl chain from acyl-CoA to a sphingoid base by ceramide synthase (such as testis-specific CerS3) (Fig. 4a). To address the mechanism underlying VLC-PUFA–containing ceramide synthesis by AdipoR2, we examined the mRNA expression of key enzymes by quantitative PCR (qPCR). Among the candidates tested, the mRNA expression of Elovl2 and CerS3 was significantly decreased in AdipoR2 KO testes compared to WT (Fig. 4b and CerS1,2,4,5,6 data in Supplementary Fig. 6a). Consistent with this, Western blotting analysis showed a moderate reduction of CerS3 protein levels and an almost complete abrogation of ELOVL2 protein expression in AdipoR2 KO testes (Fig. 4c, d). These results suggest that AdipoR2 regulates ELOVL2 expression both at the transcriptional and post-transcriptional levels. The lipidomics analysis showed that the total amount of ceramide, normalized to the total amount of PC, was not significantly changed in the AdipoR2 KO lipidome compared to WT (Supplementary Fig. 6b). Further, we found comparable localization of CerS3 protein in endoplasmic reticulum membranes in mid-pachytene spermatocytes as reported in previous studies (Supplementary Fig. 6c)⁹. These results suggest that the moderate reduction of CerS3 protein has only a minor effect on the bulk ceramide level in the AdipoR2 KO lipidome. Instead, the almost complete loss of ELOVL2 protein expression in AdipoR2 KO testes readily explains the depletion of VLC-PUFA in the AdipoR2 KO lipidome.

**Fig. 4: AdipoR2 regulates ELOVL2 both transcriptionally and post-transcriptionally.**

Membrane rigidification impairs the nuclear peripheral distribution of meiotic telomeres

To investigate the consequences of the depletion of VLC-PUFA in the AdipoR2 KO testis lipidome, we quantitatively measured the fluidity of the cellular membranes. Isolated live male germ cells were cultured in vitro and stained with Laurdan dye to measure membrane packing (often a proxy measurement for fluidity). Laurdan dye binds to membranes and emits light at 440 nm in gel-phase membranes (packed) and at 490 nm in liquid-phase membranes (loose) (Fig. 5a). The AdipoR2 KO cells showed a significantly higher generalized polarization (GP) index compared to WT cells in both spermatocytes and round spermatids (Fig. 5b, c), suggesting that the membrane became more packed and thus less fluid in the mutant cells. Moreover, bar-shaped solid-like membranous structures were frequently observed in the cytoplasm of AdipoR2 KO cells (Fig. 5b, asterisks), which likely correspond to the rigidified endoplasmic reticulum membranes reported in AdipoR1 and AdipoR2 double KO mouse embryonic fibroblasts²⁶.

**Fig. 5: A rigidified membrane impairs the telomere distribution in meiosis.**

A unique event in meiosis is the attachment of telomeres to the NE. Telomere attachment to the NE is mediated by a meiosis-specific protein complex, TERB1-TERB2-MAJIN, which recruits the SUN1-KASH5 transmembrane complex for the movement of telomeres (Fig. 5d)^18,19,31,32. Telomere movement along the NE facilitates homologous pairing, synapsis, and recombination and thus is crucial for faithful meiotic progression. Interestingly, the staining of the telomere marker TRF1 in AdipoR2 KO pachytene spermatocytes showed a significant increase in internalized telomeres dislocated from the nuclear periphery (Fig. 5e, asterisks). Notably, these internalized telomeres coincided with the internalization of Lamin B, which is a marker for the nuclear lamina normally attached to the inner NE (Fig. 5e). The same phenotypes were observed by the staining of TERB1 and TERB2 (Supplementary Fig. 7a, b). Further, the integral inner NE proteins, MAJIN, SUN1, and the outer NE protein, KASH5, which accumulate at the telomere attachment sites, were also internalized (Fig. 5e), suggesting that telomeres are internalized together with the inner and outer NE components.

The TERB1-TERB2-MAJIN complex is responsible for the formation of the telomere-NE fusion structure, known as the telomere attachment plate, as seen by TEM¹⁹. The TEM investigation of AdipoR2 KO spermatocytes showed that the formation of the telomere attachment plate was intact, which was in contrast to the Terb2 KO spermatocytes where the formation of the telomere attachment plate was completely abrogated (Fig. 5f). However, the TEM analysis still showed characteristic NE defects in AdipoR2 KO spermatocytes involving the invagination of the NE toward the inside of nucleoplasm (Fig. 5g, left). The NE invagination appeared like a membranous bubble inside the nucleus when seen in cross-section (Fig. 5g, right). Telomeres were frequently found attached to the membranous bubble (Fig. 5h and Supplementary Fig. 7c). The membranous bubble enclosed the mitochondria, supporting the notion that it is a cytoplasmic compartment invaginated into the nucleus (Fig. 5h). Taken together these results suggest that the stiffening of the nuclear membrane in AdipoR2 KO spermatocytes leads to the NE invagination, which disturb the nuclear peripheral distribution of telomeres in meiosis. It is likely that the dynein-dependent forces applied to telomeres, which normally induce telomere movement within the fluid membrane in WT, instead lead to NE curvature and subsequent invagination in cases where telomeres are tightly embedded within the rigidified membrane.

Destabilization of the intercellular bridge disrupts the germ cell syncytium

Another distinctive characteristic of the cellular membrane in germ cells is the formation of intercellular bridges between synchronously developing germ cells⁵. The intercellular bridge is visualized by the staining of the integral component TEX14⁷. The intercellular bridge matures during germ cell development, and the diameter of the TEX14 ring became larger from spermatocytes to round spermatids (Fig. 6a). In AdipoR2 KO testis the staining of TEX14 showed clear abnormalities. First, the diameter of the TEX14 ring was abnormally larger in round spermatids compared to WT spermatocytes (Fig. 6a). Further, the TEX14 signals sometimes appeared as elongated linear structures surrounding the multinucleated giant round spermatids, and the individual spermatid nuclei were no longer connected by the TEX14 ring (Fig. 6b and Supplementary Movies 1–3). These results suggest that the integrity of the intercellular bridge was significantly impaired in AdipoR2 KO cells leading to the defects in the formation of germ cell syncytia. Similar defects in TEX14 integrity were reported in germ cell-specific conditional knockouts of CerS3 and Gcs (a gene encoding a glucosylceramide synthase responsible for the addition of glycan moieties to sphingolipids)⁹, suggesting that the defects in the intercellular bridge integrity are hallmark issues caused by abnormal sphingolipid and glycosphingolipid composition in male germ cells.

**Fig. 6: Defects in the formation of intercellular bridges and germ cell syncytium.**

Male germ cells are orderly arranged within the seminiferous tubules (Fig. 6c). To test if the defects in intercellular bridge formation affect this tubular structuring, we investigated the distribution of different germ cell types in AdipoR2 KO seminiferous tubules. In WT cases, spermatocytes localized to the periphery of the seminiferous tubule, while spermatids localized to the internal lumina, and thus spermatocytes and spermatids never coexisted in the same niche within a seminiferous tubule. However, in AdipoR2 KO seminiferous tubules spermatocytes were frequently localized in the internal lumen and mixed with round spermatids in the same niche (Fig. 6d). The immunostaining of SYCP3 confirmed that SYCP3-positive spermatocytes were indeed localized in the luminal region of seminiferous tubules and were juxtaposed with round spermatids in AdipoR2 KO seminiferous tubules (Fig. 6e).

Sertoli cells create the testicular cell niche by forming the blood-testis barrier (BTB), which separates the seminiferous tubule into a basal compartment and an endoluminal compartment³³. The proper composition of lipid species in the testis is essential for the integrity of the BTB, as demonstrated by its disruption in ether lipid-deficient mice³⁴. To exclude the possibility that the abnormal distribution of germ cells within the AdipoR2 seminiferous tubule is a consequence of similar Sertoli cell defects, we tested the integrity of the BTB using a biotin tracer assay. In WT mice, the biotin tracer was observed in the basal compartment but was excluded from the endoluminal compartment (Fig. 6f, left). Cadmium chloride disrupted the BTB, leading to the leakage of the biotin tracer into the endoluminal compartment (Fig. 6f, right). In the AdipoR2 KO seminiferous tubule, however, the biotin tracer was restricted to the basal compartment and did not spread to the endoluminal compartment, which was similar to WT (Fig. 6f, middle). These data suggest that the integrity of the BTB is intact in AdipoR2 KO seminiferous tubules. Thus, we conclude that the orderly arrangement of cell types within the seminiferous tubules was impaired following the destabilization of intercellular bridges but not the BTB in AdipoR2 KO seminiferous tubules (Fig. 6f).

Discussion

AdipoR1/2 were identified as the receptors for the adipocyte-secreted protein adiponectin encoded by the Adipoq gene²³, but later studies demonstrated that they are evolutionarily conserved regulators of membrane fluidity^24,35,36. In this study, we have shown that AdipoR2 is necessary for fluidizing the cell membrane in male germ cells and thus ensuring proper meiosis progression and male fertility (Fig. 7). AdipoR2 deficiency impaired the nuclear peripheral distribution of meiotic telomeres, homologous synapsis, and recombination in spermatocytes, providing evidence linking membrane fluidity defects with specific chromosomal anomalies in meiosis. Of note, Adipoq KO mice were reported to be fertile³⁷, suggesting that AdipoR2’s role in the testis is likely to be independent of adiponectin binding, consistent with the reported adiponectin-independent roles of AdipoRs in C. elegans (where no adiponectin homolog is conserved) and in cultured mammalian cells²⁵.

AdipoR2 exhibits ceramidase activity, hydrolyzing ceramides to produce sphingosine and free FAs³⁸. The resulting sphingosine can be phosphorylated to produce the signaling molecule sphingosine-1-phosphate (S1P). In C. elegans and cultured mammalian cells, S1P is a critical downstream effector of AdipoR2 that activates specific transcription factors and promotes the transcription of FA desaturases, particularly the stearoyl-COA desaturase (SCD)²⁶. Our study found that AdipoR2 KO did not affect the mRNA expression of SCD1 (the murine homolog of human SCD) in germ cells. Instead, we found that the upregulation of CerS3 and ELOVL2 at both the transcriptional and post-transcriptional levels is the key downstream event regulated by testicular AdipoR2. In particular, the expression of the ELOVL2 protein was almost completely abolished in AdipoR2 KO testis. AdipoR2 and its C. elegans homolog PAQR-2 directly interact with an FA elongation complex³⁹, suggesting that this physical interaction may contribute to the stabilization of ELOVL2 in the testis. Overall, our findings imply that AdipoR2 ensures membrane fluidity homeostasis by controlling a comprehensive network involved in lipid metabolism, including gene transcription and protein stability, beyond the previously identified SCD-dependent pathway.

The synthesis of sphingolipids and the VLC-PUFA chain in testes depends on CerS3 and Elovl2, respectively, and the deletion of these genes causes cell death in spermatocytes^9,10. In AdipoR2 KO testis, the levels of ceramides were unchanged, but VLC-PUFA chains were largely absent from ceramides due to a significant reduction in ELOVL2 protein expression. Previous studies didn’t delve into the specific meiotic defects caused by abnormalities in the lipidome in testes. The present work identifies multiple meiotic chromosomal defects arising from the loss of VLC-PUFA. Some chromosomal defects may be direct consequences of membrane rigidification. For example, the defects in telomere attachment to the NE correlated with altered NE morphology, such as pronounced invaginations. In AdipoR2 KO spermatocytes with a highly-rigid NE lacking VLC-PUFA, the cytoplasmic dynein-dependent movement forces on telomeres might induce NE curvature, contrasting with the movements of telomeres within the fluid membrane observed in the wild-type scenario. Chromosome movements driven by the NE-attached telomeres are required for faithful homologous pairing and recombination. The defective NE properties and the telomere distribution can thus explain the observed homologous synapsis and recombination errors in AdipoR2 KO spermatocytes, similar to what has been reported in a number of different KO mice defective in telomere-driven chromosome movements^{18,19,31,40,41}. The necessity of unsaturated FAs for NE architecture appears to be evolutionarily conserved, as observed in recent findings highlighting their importance for the NE structure and function in yeast cells⁴². Alternatively, the defects in the formation of the intercellular bridges could be one of the indirect causes of the observed meiotic errors in AdipoR2 KO spermatocytes. The Tex14 KO mice showed meiotic arrest in prophase I⁷, suggesting that defects in the intercellular bridges alone without lipidome defects are sufficient to cause prophase I arrest. In summary, the meiotic defects observed in AdipoR2 KO testes are likely to result from a complex interplay, with direct consequences arising from membrane rigidification and indirect effects stemming from the disruption of intercellular bridges.

In addition to testes, VLC-PUFAs are also abundant in the retina and play crucial roles in retinal biology and photoreceptor renewal^43,44, and both Elovl2 and AdipoR1 are reported to be essential for normal retinal functions^44,45. In the retina, ELOVL2-dependent VLC-PUFA synthesis is progressively attenuated with aging due to an age-related increase in Elvol2 gene methylation^44,46,47. Similarly, AdipoR1 deficiency showed specific defects in the retina, where PCs containing VLC-PUFAs are significantly decreased leading to progressive photoreceptor degeneration⁴⁸. A recent study reported a molecular link showing that AdipoR1 upregulates Elovl2 at the transcriptional level in the retina⁴⁹, paralleling our discovery of AdipoR2’s roles in the testis. Thus, the AdipoR1/2-ELOVL2 functional axis as the master regulator of VLC-PUFA synthesis is likely to be conserved beyond the specific tissue contexts studied here.

Methods

Mouse

Mice were congenic with the C57BL/6J background. Knockout mice for AdipoR2 and Terb2 were reported earlier^19,50. All animal experiments were approved by the Regional Ethics Committee of Gothenburg, governed by the Swedish Board of Agriculture (#1316/18).

Reverse transcription PCR

Total RNA was isolated from tissues using the RNeasy Mini kit (Qiagen). cDNAs were generated using the iScript reverse transcription super mix (Bio-Rad), and PCR amplification was performed using standard DNA polymerase. The primers designed for this study are shown in Supplementary Table1.

Histological analysis

Testes and epididymides were fixed in Bouin’s fixative for 24 h at room temperature and embedded into paraffin blocks. Slices of 8 μm thickness were stained with hematoxylin and eosin. TUNEL analysis was carried out with an ApopTag Plus In Situ Apoptosis Fluorescein Detection Kit (S 7111; Millipore).

Antibodies

The following antibodies were used: rabbit antibodies against AdipoR2 (1:800 for WB)⁵⁰, TEX14 (1:1000 for WB, 1:200 for IF, Abcam; Ab41733, GR109649-1), SYCP1 (1:5000 for IF, Abcam; Ab15090, GR3184119-1), γH2AX (1:3000 for IF, Abcam; Ab11174, GR294890-8), RAD51 (1:500 for IF, Thermo Fisher Scientific; PA5-27195, UI2840658J), ELOVL2 (1:1000 for WB, Abcam; Ab176327, GR139744-4), CerS3(1:1000 for WB, 1:100 for IF)⁵¹, SUN1 (1:1000 for WB, 1:100 for IF, Abcam; Ab103021, 1014380-1), KASH5 (1:1000 for IF, Hiroki Shibuya lab), MAJIN (1:1000 for IF)¹⁹, TERB1 (1:1000 for IF)¹⁸, and TERB2 (1:1000 for IF)¹⁹; rat antibody against RPA2 (1:200 for IF, Cell Signaling Technology; 2208 S, 3); mouse antibodies against TRF1(1:1000 for IF)¹⁸, MLH1 (1:50 for IF, BD Biosciences; 51-1327GR, 4136717), and β-Actin (1:2000 for WB, Sigma; A2228-200UL, 067M4856V); goat antibody against Lamin B (1:100 for IF, Santa Cruz Biotechnology; sc-6216, F1715); and chicken antibody against SYCP3 (1:5000 for IF)⁵².

Immunostaining of spermatocytes

Testis cell suspensions were prepared and washed in PBS, centrifuged, and resuspended in hypotonic buffer (30 mM Tris (pH 7.5), 17 mM trisodium citrate, 5 mM EDTA, and 50 mM sucrose) followed by centrifugation and resuspension in 100 mM sucrose. The cell suspensions were placed on slides in the same volume of fixation buffer (1% paraformaldehyde and 0.1% Triton X-100), fixed for 3 h at room temperature, and air dried. For immunostaining, the slides were incubated with primary antibodies in PBS containing 5% BSA for 2 h and then with Alexa Fluor 488-, 594-, or 647-conjugated secondary antibodies (1:1000 dilution, Invitrogen) for 1 h at room temperature. The slides were washed with PBS and mounted with VECTASHIELD medium with DAPI (Vector Laboratories). Signal intensities were measured using the SoftWorx Data Inspector tool. The background signals were measured in the same way and subtracted.

Lipidomics

Mouse testes were removed and immediately frozen on dry ice, and tissues were stored at –80 °C until analysis. For lipid extraction, tissues were extracted using the BUME method according to previous work⁵³. Internal standards were added during lipid extraction. Phospholipids and sphingomyelins were measured using direct infusion mass spectrometry. For phospholipids, a part of the lipid extracts was evaporated and reconstituted in chloroform:methanol [1:2] with 5 mM ammonium acetate. This solution was infused directly (shotgun approach) into a QTRAP 5500 mass spectrometer (Sciex) equipped with a TriVersa NanoMate (Advion Bioscience) as described previously⁵⁴. A similar procedure was used for sphingomyelin. However, prior to analysis the lipid extracts were evaporated and exposed to alkaline hydrolysis (0.1 M KOH in methanol for 60 minutes at room temperature) in order to remove the interfering phospholipids. For phosphatidylcholines and sphingomyelins, mass spectra were obtained in precursor ion scanning mode using the phosphocholine headgroup (m/z 184.1) as the fragment ion. For phosphatidylethanolamines, neutral loss scanning of m/z 141.1 was used^55,56. Ceramides were measured using ultra-performance liquid chromatography coupled to tandem mass spectrometry as previously described⁵⁷. In brief, the separation of ceramides species was performed using a Waters BEH C8 column (2.1 × 100 mm with 1.7 µm particles) with water, acetonitrile, and isopropanol as mobile phases. The detection was made in positive MRM (targeted) mode using m/z 264 as a fragment ion. The lipidomics data were evaluated using the LipidView and MultiQuant software (Sciex), and Qlucore Omics Explorer software was used for the multivariant analysis. The complete lipid composition data are provided in the Source data.

Measurement of membrane packing

Live cells were stained with Laurdan dye (6-dodecanoyl-2-dimethylaminonaphthalene) (Invitrogen) at 15 μM for 1 h. Images were acquired with an LSM880 confocal microscope (Zeiss) equipped with a live cell chamber (set at 33 °C and 5% CO2) and ZEN software with a 40× water-immersion objective. Briefly, cells were excited with a 405 nm laser and the emission was recorded between 410 nm and 461 nm (ordered phase) and between 470 nm and 530 nm (disordered phase). Images were acquired with 16-bit image depth and 1024 × 1024-pixel resolution using a pixel dwell of ~1.02 μs. The GP index was calculated using ImageJ version 1.47 software following published guidelines⁵⁸. SYTO59 (5 mM in DMSO) (Invitrogen) was added together with the Laurdan dye to stain the nuclei and allow distinguishing between cell types. SYTO59 was excited with a 561 nm laser, and the emission was recorded between 570 nm and 758 nm.

Biotin tracer assay

WT and AdipoR2 KO mice were anesthetized, and 20 μl EZ-Link Sulfo-NHS-LC-Biotin (10 mg/ml in PBS containing 1 mM CaCl2) (Thermo Fisher Scientific) was injected into the testis interstitium. Mice were euthanatized after 40 minutes. The testes were removed and fixed in 4% formaldehyde overnight, and 5 µm sections were obtained and stained with FITC-conjugated streptavidin (Invitrogen) and DAPI. For quantification, the distance traveled by biotin (D_Biotin) from the basement membrane in a seminiferous tubule versus the radius of the tubule (D_Radius) was calculated. For oval tubules, the average of the longest distance and the shortest distance from the basement membrane was used. As the positive control, WT mice were treated with daily doses of CdCl2 (5 mg/kg, i.p.) (Sigma) for 3 consecutive days before the BTB assay.

qPCR

Total RNA was extracted from both WT and AdipoR2 KO testis biopsies using the RNeasy Mini Kit according to the manufacturer’s protocol (Qiagen) and quantified by a NanoDrop spectrophotometer (ND-1000; Thermo Fisher). cDNA was synthesized using the SuperScript III First-Strand Synthesis System (Invitrogen), and the qPCR experiment was performed with a CFX Connect Real-Time PCR Detection System (BioRad) using the iTaq Universal SYBR Green Supermix qPCR system (BioRad) and standard primers. The relative expression of each gene was measured by the delta-Ct method. The primers designed for this study are shown in Supplementary Table1.

TEM

For TEM, testis tissue samples were prepared and fixed as previously described⁵⁹. Small testis tissue pieces were fixed at 4 °C in cacodylate-buffered glutaraldehyde solution (2.5% glutaraldehyde, 50 mM KCl, 2.5 mM MgCl, 50 mM cacodylate; pH 7.2) for 1 h and subsequently washed five times for 3 min each in fresh cacodylate buffer (50 mM cacodylate, pH 7.2). The samples were then incubated in 2% osmium tetroxide in 50 mM cacodylate for 2 h at 4 °C washed three times in distilled H₂O and contrasted overnight at 4 °C with 0.5% uranyl acetate (in distilled H₂O). The tissue pieces were dehydrated in an increasing ethanol series and incubated three times in propylene oxide for 30 min each time at room temperature. Finally, the samples were embedded in epon. Ultrathin sections (65 nm to 100 nm) were transferred to Formvar-coated copper grids and treated with fresh 2% uranyl acetate solution in distilled H₂O for 20 min at room temperature. After washing in distilled H₂O, testis sections were counterstained in Reynold’s lead citrate for 10 min, washed in distilled H₂O, and dried at room temperature. Samples were analyzed with a JEOL JEM-1400 Flash Scanning Transmission Electron Microscope (Jeol, Eching, Germany) operated at 120 kV. For the analysis of Terb2 KO samples, some minor changes in the fixation/embedding protocol were performed as previously described¹⁹.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

All data are available in the main text or the supplementary materials. All other data supporting the findings of this study are available from the corresponding author upon request. Source data are provided in this paper.

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Acknowledgements

We thank Roger Sandhoff (German Cancer Research Center) for providing the CerS3 antibody. The Jeol JEM-1400 STEM was funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation)—426173797 (INST 93/1003-1 FUGG). This work was supported by the German Research Foundation Al 1090/4-2 (M.A), the European Research Council StG-801659 (H.S.), the Swedish Research Council 2018-03426 (H.S.), and the Knut och Alice Wallensbergs Stiftelse KAW2019.0180 (H.S.).

Funding

Open access funding provided by University of Gothenburg.

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Abrahan Hernández-Hernández
Present address: National Genomics Infrastructure, Science for Life Laboratory, Department of Cell and Molecular Biology, Karolinska Institute, Stockholm, Sweden

Authors and Affiliations

Department of Chemistry and Molecular Biology, University of Gothenburg, 41390, Gothenburg, Sweden
Jingjing Zhang, Mario Ruiz, Nena Stojanović, Ranjan Devkota, Marc Pilon & Hiroki Shibuya
Department of Molecular and Clinical Medicine/Wallenberg Laboratory, Institute of Medicine, University of Gothenburg, 41467, Gothenburg, Sweden
Per-Olof Bergh, Marcus Henricsson & Jan Borén
Department of Cell and Developmental Biology, Biocenter, University of Würzburg, 97074, Würzburg, Germany
Marius Henn & Manfred Alsheimer
Discovery Sciences, BioPharmaceuticals R&D, AstraZeneca, Gothenburg, Sweden
Mohammad Bohlooly-Y
Department of Cell and Molecular Biology, Karolinska Institutet, Stockholm, Sweden
Abrahan Hernández-Hernández
Laboratory for Gametogenesis, RIKEN Center for Biosystems Dynamics Research (BDR), Kobe, Japan
Hiroki Shibuya

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Jingjing Zhang
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Contributions

J.Z., M.R., N.S., R.D. and M.B. performed the mouse experiments and analyzed the data; P.O.B. and M. Henricsson performed the lipidomics analysis with supervision from J.B.; M.H., M.A., and A.H. performed the TEM analysis; M.P. and H.S. supervised the project; and H.S. wrote the manuscript.

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Zhang, J., Ruiz, M., Bergh, PO. et al. Regulation of meiotic telomere dynamics through membrane fluidity promoted by AdipoR2-ELOVL2. Nat Commun 15, 2315 (2024). https://doi.org/10.1038/s41467-024-46718-6

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Received: 01 September 2023
Accepted: 29 February 2024
Published: 14 March 2024
DOI: https://doi.org/10.1038/s41467-024-46718-6

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