NLRP2 controls age-associated maternal fertility

Nucleotide-binding domain and leucine-rich repeat (NLR) proteins are well-established hub proteins regulating a diversity of inflammatory and host defense responses in the immune system. Class II major histocompatibility complex transactivator (CIITA), nucleotide-binding oligomerization domain receptor 1 (NOD1), and NOD2 are founding family members that, respectively, control MHC-II transcription and peptidoglycan-induced NF-κB and MAP kinase activation, whereas NLRC5 regulates MHC-I gene expression (Kanneganti et al., 2007; Ting et al., 2008; Geddes et al., 2009; Lupfer and Kanneganti, 2013). Another subset of NLRs plays central roles in the assembly of inflammasomes, multiprotein scaffolds that couple direct and indirect detection of pathogen- and danger-associated molecular patterns to recruitment and activation of the prototype inflammatory caspase-1 for production of bioactive IL-1β and IL-18. We distinguish the Nlrp1a and Nlrp1b, Nlrp3, and Nlrc4 inflammasomes, among several other emerging inflammasomes (Lamkanfi and Dixit, 2014). The NLRP1a inflammasome currently lacks a defined activating agent, but bacterial pathogens such as Salmonella enterica serovar Typhimurium, Pseudomonas aeruginosa, Legionella pneumophila, and Shigella flexneri all engage the NAIP/Nlrc4 inflammasome. The Nlrp1b inflammasome is triggered by anthrax lethal toxin (LeTx). Finally, the Nlrp3 inflammasome reacts to a broad range of bacterial, viral, and fungal pathogens, as well as danger-associated molecular patterns linked to autoimmune diseases (Lamkanfi and Dixit, 2012).

Unlike the aforementioned NLRs, the roles of NLRP2 are ill-defined. Ectopically expressed NLRP2 was shown to regulate inflammasome signaling and to inhibit NF-κB activation (Agostini et al., 2004; Bruey et al., 2004; Kinoshita et al., 2005; Fontalba et al., 2007). Knockdown studies have also suggested Nlrp2 to be essential for early mouse embryogenesis (Peng et al., 2012). Despite these advances, the physiological roles of Nlrp2 are unknown. Here, we describe Nlrp2-deficient mice and report that Nlrp2 deletion altered neither innate nor adaptive immune responses in ex vivo–stimulated leukocytes and in in vivo–challenged mice. However, we identified a role for oocyte-expressed Nlrp2 in early embryogenesis and show that Nlrp2 contributed importantly to in vivo fertility with progressing maternal age. Thus, this work highlights Nlrp2 as a critical and selective regulator of oocyte quality, suggesting that NLRP2 variants with reduced activity may contribute to maternal age-associated fertility loss in humans.

To study the in vivo roles of Nlrp2, we generated Nlrp2-deficient mice by flanking exon 5—which encodes most of the central NACHT ATPase domain—with LoxP sites to allow Cre recombinase-assisted conditional deletion of the targeted exon (Fig. S1). Surprisingly, full-body Nlrp2^−/− mice were born from heterozygous parents with expected Mendelian ratios, and no phenotypic abnormalities were noted (Table S1). To probe for putative roles of Nlrp2 in inflammasome regulation, we assessed IL-1β secretion from LPS-primed Nlrp2^−/− and WT BM-derived macrophages (BMDMs) after exposure to the canonical Nlrp3 inflammasome stimulus nigericin. The levels of IL-1β detected in culture supernatants of Nlrp2^−/− macrophages were indistinguishable from those of WT BMDMs (Fig. 1 a). In agreement, pyroptosis (Fig. 1 b) and nigericin-induced caspase-1 maturation (Fig. 1 c) were unaffected. Moreover, activation of neither the noncanonical Nlrp3 inflammasome (Fig. 1, d–f) nor that of other canonical inflammasomes was altered in Nlrp2^−/− macrophages in terms of IL-1β secretion, caspase-1 maturation, and pyroptosis induction (Fig. 1, g–o). Together, these results indicate that Nlrp2 is dispensable for known inflammasome pathways.

Although ectopically expressed Nlrp2 inhibited activation of the proinflammatory transcription factor NF-κB (Bruey et al., 2004; Kinoshita et al., 2005; Fontalba et al., 2007), we found that production levels of the NF-κB–dependent cytokine TNF in Salmonella Typhimurium–infected macrophages were not modulated by Nlrp2-deficiency (Fig. 1 p). Nor did we detect differential levels of TNF and IL-6 in Nlrp2-sufficient and -deficient DCs that have been stimulated for 6 or 24 h with the TLR2 ligands FSL-1 and Pam3CSK4 or the TLR4 ligand LPS, respectively (Fig. 1, q–t).

Given that Nlrp2 appeared to be negligible for regulation of the examined inflammasome and TLR-induced NF-κB signaling pathways, we sought to more broadly address the role of Nlrp2 in the immune system. To this end, we selectively ablated Nlrp2 expression in the hematopoietic compartment by breeding Nlrp2^flox/flox mice to mice expressing the Cre recombinase under control of the mouse HS21/45-vav control regions. CD4⁺ T lymphocytes were collected and stimulated ex vivo with anti-CD3 and anti-CD28 monoclonal antibodies to mimic stimulation by antigen-presenting cells. This treatment regimen triggered Nlrp2^−/− CD4⁺ T cells to produce the hallmark cytokines IL-2 and IFN-γ, as well as TNF and IL-6, with similar potency to WT CD4⁺ T cells (Fig. 2, a–d). Nlrp2 deficiency also failed to modulate T cell receptor–independent cytokine production and lymphocyte proliferation in CD4⁺ T lymphocytes that have been treated with PMA and ionomycin (Fig. 2, e–h). In vivo immunophenotyping further revealed normal distribution profiles of lymphocyte and myeloid cell fractions in thymus, spleen, lymph nodes, and BM of unchallenged full-body Nlrp2^−/− mice (Fig. 2, i–m). Moreover, Nlrp2^−/− mice had normal in vivo T cell responses in the NP-KLH in alum immunization model, and expected lethality rates in the LPS-endotoxemia model (unpublished data). Together, these findings suggest that unlike many other NLR family members, Nlrp2 is dispensable for innate and adaptive immunity.

In seeking to define the physiological role of Nlrp2, we characterized its mRNA expression profile in different organs. In agreement with reported findings (Peng et al., 2012), a query of public microarray data revealed abundant expression of Nlrp2 mRNA in oocytes, whereas a variety of other murine tissues and cell types returned negative for Nlrp2 expression (Fig. 3 a). A similar oocyte-restricted expression profile emerged from probing human expression data (Fig. 3 b). Quantitative RT–PCR analysis confirmed the selective expression of Nlrp2 in ovaries (containing oocytes), and its absence in the male reproductive system and somatic tissues of the mouse (Fig. 3, c and d). Several other NLR family members with oocyte expression have been reported (Tong et al., 2000; Peng et al., 2012, 2015), but unlike Nlrp2, transcripts of Nlrp5, Nlrp9b, and Nlrp14 were also abundantly expressed in the male reproductive system (testis, epididymis, and sperm; Fig. 3 e). Together, these results suggest that Nlrp2 expression largely is confined to oocytes.

The observation that Nlrp2 was abundant in oocytes prompted us to investigate its significance for early embryogenesis. Oocytes from superovulated Nlrp2^−/− mice and WT controls were collected and analyzed for their developmental potential upon parthenogenetic activation with strontium chloride (Ma et al., 2005). Oocytes of 6–10-wk-old Nlrp2^−/− mice—approximating young adult age in humans—progressed through the two-cell, four-cell, and morula embryonic stages with normal efficiency (Fig. 4, a and b) and kinetics (Fig. 4 c), with ∼30% of both Nlrp2-sufficient and -deficient oocytes having adopted the morphological appearance of blastocysts 5 d after parthenogenetic activation (Fig. 4, a–c). However, Nlrp2^−/− oocytes from mice aged 22–25 wk—which approximates mature adult age in humans—progressed asynchronously and with significantly reduced rates through the four-cell and morula stages (Fig. 5, a–c). This contrasted markedly with oocytes from similarly aged WT mice, close to 80% of which successfully developed into blastocysts (Fig. 5 c). Instead, most Nlrp2^−/− embryos were lost or appeared fragmented by day 5, and <10% reached the blastocyst stage (Fig. 5). Despite the striking age-associated defect in early embryogenesis, metaphase II–arrested oocytes collected from both 6–10- and 22–25-wk-old Nlrp2^−/− mice had spindles of seemingly normal size and shape (Fig. S2, a–d). Also spindle-mediated chromosome segregation appeared to be grossly normal when analyzed 1 h after parthenogenetic activation (Fig. S2, e and f). Many genes that contribute importantly to early embryogenesis in mammals are subject to imprinting, an epigenetic process of gene silencing in reproductive cells that allows selective expression of alleles in a parent-of-origin–dependent manner (Van Gorp et al., 2014). Given that the nucleotide sequences of Nlrp2 transcripts of C57BL/6J, BALB/c, and 129S1 mice are fully conserved in their open reading frames (unpublished data), naturally encoded Nlrp2 mRNA variants could not be exploited to distinguish maternal- and paternal-derived Nlrp2 transcripts in oocytes. We therefore took advantage of the differential Nlrp2 alleles of WT and Nlrp2^−/− mice to establish whether Nlrp2 was imprinted (Fig. S3 a). After breeding Nlrp2^−/− females to Nlrp2^+/+ males, RT-PCR analysis of oocytes of the resulting heterozygous offspring showed the presence of transcripts corresponding to both the WT and knockout Nlrp2 alleles (Fig. S3 b), indicating that Nlrp2 is not imprinted because both the parental and maternal Nlrp2 alleles were actively transcribed. Together, these results suggest that Nlrp2 regulates early embryogenesis after oocyte activation.

Although more work is needed to pinpoint the molecular mechanisms involved, the aforementioned findings led us to hypothesize that Nlrp2 may exert a role in vivo in early embryogenesis that gains significance with progressing maternal age. To empirically address this possibility, WT and Nlrp2^−/− females were bred to randomly selected WT and Nlrp2^−/− males when aged 7 wk, and their reproductive performance was monitored over time until they became 45 wk old. Notably, litters were comparably sized when WT and Nlrp2^−/− females were 9–15 wk old, but Nlrp2^−/− females delivered significantly less pups/litter as they grew older (Fig. 6 a). Nlrp2 likely controls maternal fertility independently of inflammasomes because females lacking the central inflammasome proteases caspases 1 and 11 did not have reduced litter sizes with progressing age (Fig. 6 b). We next addressed whether the in vivo role of Nlrp2 in regulation of age-associated fertility extended to males. To this end, Nlrp2^−/− and WT males that were aged >16 wk were mated with WT females of similar age to compare the reproductive performance of these breeding couples. Nlrp2^−/− males had litter sizes that were comparable to those of WT males (Fig. 6 c), whereas a control group of Nlrp2^−/− males that was mated to Nlrp2^−/− females showed a significant reduction in the number of pups delivered per litter (Fig. 6 c). We also performed a reciprocal experiment in which WT males were bred to either WT or Nlrp2^−/− females. In agreement with our other findings, Nlrp2^−/− females had markedly less pups per litter relative to their WT counterparts (Fig. 6 d). These results confirm that Nlrp2 selectively controls maternal fertility in mature adult mice. To further validate and expand on these findings in a controlled setting, four Nlrp2^−/− males of proven fertility were each co-housed with one randomly selected WT and one Nlrp2^−/− female of >14 wk old. Litter sizes in this experiment were determined based on genotype of the pups with all heterozygous pups in the breeding cage being assigned to the WT female, and Nlrp2^−/− pups being born from the knockout female. In agreement with our other observations, we found that litters of mature adult Nlrp2^−/− females were significantly smaller than those of WT females (Fig. 6 e), confirming that Nlrp2 is required for optimal fertility with progressing maternal age.

In conclusion, we showed here that Nlrp2 selectively controls maternal age-associated fertility. Together with the reported sterility of female mice with a hypomorphic Nlrp5 allele (Tong et al., 2000), our results define a growing subset of NLRs with demonstrated in vivo roles in maternal fertility. Defective Nlrp5 causes female sterility as a result of an oocyte-intrinsic inability to develop beyond the two-cell embryonic stage, whereas the results presented here implicate Nlrp2 in an oocyte-intrinsic developmental checkpoint that gains significance with progressing maternal age. These findings set the experimental context for future studies aimed at elucidating the molecular mechanisms through with NLRP2 controls early embryogenesis. Among other possibilities, Nlrp2 may be a maternal effect gene that directly or indirectly supports proper imprinted gene expression patterns and pluripotency of developing zygotes under ovarian age-associated stress conditions (Barton et al., 1984; Van Gorp et al., 2014). Defective NLRP2 signaling may thus affect reproductive function in patients. Indeed, anecdotal evidence links mutations in human NLRP2 with increased susceptibility to reproductive disorders, such as sporadic Beckwith-Wiedemann syndrome (Meyer et al., 2009) and Silver–Russell syndrome (Court et al., 2013). Moreover, humans encode a second (primate-specific) Nlrp2 orthologous gene, NLRP7, that is frequently found mutated in patients with familial cases of biparental molar pregnancies (Nguyen and Slim, 2014; Van Gorp et al., 2014). As increased maternal age is a prime factor driving the fast-growing societal demand for in vitro fertilization and other assisted reproductive technologies, our findings additionally support examination of whether NLRP2 and NLRP7 polymorphisms impact on embryo quality and success rates in these patient groups. Thus, this work sheds light on the role of Nlrp2 as a critical checkpoint of early preimplantation embryogenesis, and warrants investigation of defective NLRP2 and NLRP7 function in women with early menopause manifestation and other conditions associated with declining maternal fertility.

Nlrp2^floxNeo+ mice were generated at Ingenious Targeting Laboratory Inc. by conditional targeting of exon 5 in C57BL/6 embryonic stem (ES) cells. Resulting chimeras were bred to Flp-transgenic mice to delete the FLP recombinase target (FRT) sequences-flanked neomycin selection cassette. Nlrp2-deficient mice were obtained after breeding to appropriate Cre recombinase-expressing mice. Casp1^−/−Casp11^−/− mice have been reported (Kuida et al., 1995). All mice were housed under specific pathogen–free conditions, and animal studies were conducted under protocols and guidelines approved by the Ghent University animal care and use committee (ECD14/74; ECD15/02).

The following primers were used for genotyping Nlrp2^−/− mice: R1, 5′-CCTGGTGAATCCAATGTCTAGG-3′; F1, 5′-GGAGCCCTGATGATTCTTTGG-3′; F2, 5′-CTGAAGTCTCTCATTCTCTATAGG-3′. Nlrp2⁺ (primers R1, F2), 406 bp; Nlrp2^flox (primers R1, F2), 583 bp; and Nlrp2⁻ (primers R1, F1), 691 bp.

Females of the indicated age and genotypes were superovulated by i.p. injection of pregnant mare serum gonadotropin (PMSG; 5 IU; Folligon; Intervet), 48 h later followed by i.p. injection of human chorionic gonadotrophin (hCG; 5 IU; Chorulon; Intervet). Mice were euthanized, cumulus-enclosed oocytes were recovered and treated with hyaluronidase (0.3 mg/ml; H4272; Sigma-Aldrich) in M2 media (M7167-50ML; Sigma-Aldrich) to remove cumulus cells. Oocytes were then washed and maintained in KSOM (MR-121-D; EMD Millipore) in an incubator (at 37°C and 5% CO₂) for 2 h. Subsequently, oocytes were incubated in Ca²⁺-free HTF media containing SrCl₂ (10 mM) and cytochalasin D (2 µg/ml; C8273; Sigma-Aldrich). 4 h later, embryos were washed and cultured further in KSOM medium in a 5% CO₂ incubator at 37°C. Morulas were transferred to blastocyst media (K-SIBM-50; Cook Medical). Developing embryos were analyzed under 20× objective of IX71 inverted Microscope by CCD Camera XM-10 (Olympus) with resolution 0.6442 µm/pixel in Imaging software CellB (Olympus).

Oocytes were collected from oviducts of superovulated mice, transferred to extraction/fixation solution (200 µl 50 mM Hepes, 4% paraformaldehyde, 0.2% Triton X-100, 5 mM MgCl₂, 5 mM EGTA, pH 7.4, and 0.1 M PIPES, pH 7.4, in water) in 9-well Pyrex plates (13-748B; Thermo Fisher Scientific) and incubated at room temperature for 1 h before incubation in 200 µl blocking solution (PBS; BSA [4%] and Tween [0.25%]) for an additional 1 h. Oocytes were stained with an α-tubulin antibody (1:200; T9026; Sigma-Aldrich) in blocking solution, washed, and incubated with a donkey anti–mouse IgG conjugated to Alexa Fluor 488 (A-21202; Thermo Fisher Scientific). Mounted samples were visualized under a TCS SP5 (Leica) and confocal microscopes. Spindle length was measured after 3D reconstruction of 2-µm z-stacks using Imaris software.

RNA was extracted from mouse tissues using RNeasy mini kit (74104; QIAGEN) according to the manufacturer’s protocol. RNA concentration was measured using a NanoDrop ND-8000, and cDNA was synthesized using QuantiTect Reverse Transcription kit (205311; QIAGEN) according to the manufacturer’s instructions.

The following primers were used for RT-PCR analysis: Nlrp2F: 5′-CCCTGAAAGTAGAAAAGGCGG-3′; Nlrp2R: 5′-CCACTTTCTGGAGACAACAGG-3′; Nlrp9bF: 5′-TCAGATGTATTATCAGTGACTGC-3′; Nlrp9bR: 5′-GGAGACATCGAATTACAGAGG-3′; Nlrp5F: 5′-GCTGCTGATTAACCAGAAGATGG-3′; Nlrp5R: 5′-GGCCCGTTGACTCAGGATGC-3′; Nlrp14F: 5′-GCTCAGCCATACCTTGAAGC-3′; Nlrp14R: 5′-CTCCAGCTCCTGAAGATTGC-3′. Nlrp2 cDNAs were amplified for imprinting analysis with the following primers: Nlrp2R2: 5′-AAAAATGAAATGAGGAAGAGGAGG-3′; Nlrp2F2: 5′-AGTGAAAGCTGGTGTGGTGG-3′. Phusion Hot Start Flex 2X Master Mix (M0536L; NEB) was used for PCR amplification.

qRT-PCR was performed on a mouse normal tissue qPCR array (MNRT301; Origene) with the LightCycler 480 SYBR Green I Master (4707516001; Roche) and the following primers: NLRP2F: 5′-CCCTGAAAGTAGAAAAGGCGG-3′; Nlrp2R: 5′-CCACTTTCTGGAGACAACAGG-3′; GapdhF: 5′-GGTGAAGGTCGGTGTGAACG-3; GapdhR: 5′-CTCGCTCCTGGAAGATGGTG-3′. Nlrp2 expression levels were normalized to Gapdh.

BMDMs and DCs were prepared as previously described (Vande Walle et al., 2014). The following reagents were used as indicated: FSL-1 (100 ng/ml; tlrl-fsl; InvivoGen), Pam3CSK4 (100 ng/ml; tlrl-pms; InvivoGen), S. minnesota LPS (100 ng/ml or 500 ng/ml; tlrl-smlps; InvivoGen), nigericin (20 µM, N7143-10MG; Sigma-Aldrich), C. difficile toxin B (1 µg/ml; #155L; List Biological Laboratories); Lipofectamine-2000 (5 µl; 11668–019; Invitrogen), Fugene (E2311; Promega). CD4⁺ T cells were purified by negative selection with CD4⁺ T cells isolation kit II (130–095-248; Miltenyi Biotec) according to manufacturer’s instructions. CD4⁺ T cell were activated with plate-bound anti-CD3 (4 µg/ml; 14–0031-85; eBioscience) and anti-CD28 antibodies (1 µg/ml; 14–0281-82; eBioscience), or with PMA (100 ng/ml; P1585-1MG; Sigma-Aldrich) together with ionomycin (1 µg/ml; I0634; Sigma-Aldrich).

Cytokines were analyzed with Luminex (Bio-Rad Laboratories) and ELISA (R&D Systems) assays according to guidelines of the manufacturers. Cell death was determined by lactate dehydrogenase (LDH) assay (G1780; Promega).

Cell lysates and culture supernatants were incubated with 1X Caspase Lysis Buffer (0.1% NP-40, 20 mM NaCl, 2 mM Tris-HCl, pH 7.4) on ice and denatured in Laemmli buffer. Samples were boiled and separated by SDS-PAGE. Separated proteins were transferred to PVDF membranes. Blocking, incubation with antibody, and washing of the membrane were done in PBS supplemented with 0.05% Tween-20 (vol/vol) and 3% (wt/vol) nonfat dry milk. Immunoblots were incubated overnight with primary antibodies against caspase-1 (AG-20B-0042-C100; 1:1,000; Adipogen), followed by HRP-conjugated secondary anti–mouse antibodies (1:5,000; 115–035-146; Jackson ImmunoResearch Laboratories) to detect proteins by enhanced chemiluminescence (Thermo Fisher Scientific).

Cells from primary and secondary immune organs were prepared by mechanical dissociation through 70-µm cell strainers (08–771-2; Corning). 1–5 million cells were blocked with anti-mouse CD16/CD32 antibody (14–0161-85, eBioscience), and stained with anti-TCRβ-biotin (13–5961-81, eBioscience) followed by incubation with SA-APC (17–4317-82; eBioscience,) and anti-CD19-PE (12–0193-81, eBioscience) for spleen and LN cells; anti-B220-PE (12–0452-82; eBioscience) for BM cells; anti-CD4-FITC (11–0041-82; eBioscience) and anti-CD8-APC (17–0081-81; eBioscience) for thymocytes; and anti-CD11b-APC (17–0112-81; eBioscience), anti-GR1-FITC (11–5931-82; eBioscience) and anti-CD11c-PE (12–0114-81; eBioscience) for myeloid cells in the presence of propidium iodide for analysis of cell death. Cells were analyzed on a FACSCalibur (BD) machine.

Statistical analyses were performed using the Student’s t test in GraphPad Prism5 software. P < 0.05 was considered statistically significant.

Fig. S1 describes production and genotyping of Nlrp2^−/− mice. Fig. S2 shows normal spindle morphology in Nlrp2^−/− oocytes. Fig. S3 shows that Nlrp2 is not imprinted. Table S1 shows that Nlrp2^−/− mice are born with expected Mendelian ratios from heterozygous parents.

The authors thank the VIB Bio Imaging core for assistance with imaging, and Dr. Marnik Vuylsteke for statistical analysis support.

This work was supported by European Research Council grant 281600, the Fund for Scientific Research-Flanders grant G011315N, and the Baillet Latour Medical Research grant to M. Lamkanfi.

The authors declare no competing financial interests.

Author contributions: A.A. Kuchmiy and M. Lamkanfi designed the study; A.A. Kuchmiy, J. D’Hont, and T. Hochepied performed experiments; A.A. Kuchmiy, J. D’Hont, T. Hochepied, and M. Lamkanfi analyzed data. A.A. Kuchmiy and M. Lamkanfi wrote the manuscript with input from the other authors, and M. Lamkanfi oversaw the project.