Skip to main content
NCBI home page
Genetics logoLink to Genetics
. 2017 Nov 22;208(2):549–563. doi: 10.1534/genetics.117.1123

The Germline-Specific Factor OEF-1 Facilitates Coordinated Progression Through Germ Cell Development in Caenorhabditis elegans

Catherine E McManus 1, Valerie Reinke 1,1
PMCID: PMC5788521  PMID: 29167199

Abstract

The purpose of germ cells is to ensure the faithful transmission of genetic material to the next generation. To develop into mature gametes, germ cells must pass through cell cycle checkpoints while maintaining totipotency and genomic integrity. How germ cells coordinate developmental events while simultaneously protecting their unique fate is not well understood. Here, we characterize a novel nuclear protein, Oocyte-Excluded Factor-1 (OEF-1), with highly specific germline expression in Caenorhabditis elegans. OEF-1 is initially detected early in embryogenesis and is expressed in the nuclei of all germ cells during larval stages. In adults, OEF-1 expression abruptly decreases just prior to oocyte differentiation. In oef-1 mutants, the developmental progression of germ cells is accelerated, resulting in subtle defects at multiple stages of germ cell development. Lastly, OEF-1 is primarily associated with the bodies of germline-expressed genes, and as such is excluded from the X chromosome. We hypothesize that OEF-1 may regulate the rate of progression through germ cell development, providing insight into how these critical maturation events are coordinated.

Keywords: germline, mitosis, meiosis, gametogenesis, checkpoints, C. elegans

AS the only cells passed from generation to generation, the germline of an organism is critical to the survival of the entire species. Germ cells undergo specialized processes such as meiosis and gametogenesis to produce a totipotent zygote upon fertilization (Lesch and Page 2012). Thus, germ cells proceed through multiple stages of maturation while maintaining genomic integrity and preventing somatic differentiation (Robert et al. 2015). Deciphering the mechanisms that protect germ cells while permitting their development is critical to our overall understanding of how cell fates are specified.

In the nematode Caenorhabditis elegans, the germ lineage, known as the P lineage, is specified in the embryo via the asymmetric divisions of embryonic blastomeres (Wang and Seydoux 2013). The germline precursor cell P4 is formed at the 24-cell stage, and undergoes a single symmetric division to generate the two primordial germ cells (PGCs), Z2 and Z3, at the 88-cell stage (Wang and Seydoux 2013). In adult worms, germ cells progress from proliferative progenitor cells to fully differentiated gametes within each of two gonad arms, with precise, multi-level molecular control at each cell fate juncture (Kimble and Crittenden 2007). In part, transcriptional repression of somatic transcripts through chromatin regulation maintains the unique germline fate throughout development (Robert et al. 2015). The entire X chromosome, for example, which houses few germline-expressed genes (Reinke et al. 2004), is held transcriptionally silent through most of germ cell development via the maintenance of repressive histone modifications (Kelly et al. 2002). Extensive post-transcriptional mechanisms function as another level of germ cell fate control. The inhibition or stabilization of specific transcripts by RNA-binding proteins enables rapid switching to different germ cell programs, such as the mitosis-to-meiosis transition or the sperm-to-oocyte switch (Kimble and Crittenden 2007). Finally, during these cell fate transitions, homologous chromosomes must pair, synapse, and recombine so that chromosomes can segregate properly (Hillers et al. 2017). If synapsis of any chromosome pair is delayed or fails, the synapsis checkpoint triggers apoptosis in these germ cells to prevent the formation of aneuploid gametes (Bhalla and Dernburg 2005). While some molecular mechanisms have been implicated in these transcriptional and checkpoint pathways, how critical events in germ cells are coordinated and interconnected remains poorly understood.

Here, we characterize the expression, regulation, and function of a novel, highly germline-specific nuclear factor that we have named OEF-1 (Oocyte-Excluded Factor-1). We define spatial and temporal relationships between oef-1 transcript and protein expression in the C. elegans germline throughout development, including exceedingly early protein expression in the P2 blastomere. OEF-1 is expressed throughout germline development, but appears to be actively excluded from germ cells undergoing oogenesis. oef-1 mutants exhibit faster progression of germ cells through multiple stages of development and differentiation, along with increased apoptosis due to activation of the synapsis checkpoint. Genome-wide binding site analysis demonstrates that OEF-1 preferentially associates with the bodies of germline-expressed genes on autosomes, and is largely excluded from the X chromosome. We suggest that OEF-1 might coordinate the timing of multiple germline processes as germ cells undergo critical regulatory transitions.

Materials and Methods

Strains

C. elegans strains were maintained by standard methods as described (Brenner 1974). Bristol N2 was used as the wild-type reference strain. All growth was performed at 20°, except for BA17, JK654, and YL312, which were maintained at 15° and shifted to 25° to induce sterility.

Antibody generation

Complementary DNA (cDNA) corresponding to the first 100 amino acids of OEF-1 was cloned into the pET-19b expression vector (Novagen) and transformed into BL21 cells. His-tagged OEF-1 N-terminal fragments were purified over Nickle-NTA resin (QIAGEN, Valencia, CA) under denaturing conditions, and injected into rabbits (Pocono Rabbit Farm and Laboratory). Affinity purification of the bleeds was performed as in Porter and Koelle (2010). Briefly, purified OEF-1 N-terminal protein was run on an acrylamide gel and transferred onto nitrocellulose. The bleeds were incubated with the membrane overnight at 4°, and bound antibodies were eluted off the membrane with 100 mM glycine (pH 2.5).

Apoptosis assays

Worms 16 hr beyond larval stage 4 (L4) were picked into 1 ml of 33 μM SYTO 12 (Molecular Probes, Eugene, OR) diluted in M9. After 4 hr of incubation at 23°, the worms were plated and fed for at least 30 min before visualization on agarose pads. The number of SYTO 12-positive cells per gonad arm was quantified. For checkpoint epistasis experiments, genotypes were blinded before quantification.

Brood size analyses

L4 worms were singly placed on plates seeded with the bacterial food source OP50, and moved to fresh plates each day until embryo production ceased. Unhatched embryos were scored as dead 24 hr after shifting. Larvae were counted 2 days after shifting and males were scored after 3 days. For him-8 brood size analyses, wild-type and him-8(e1489) L1s were grown on HT115 bacteria expressing empty L4440 vector or oef-1 dsRNA in L4440 on RNA interference (RNAi) plates, as in Fraser et al. (2000). F1 L4s were singly placed on fresh RNAi plates and broods were analyzed as above.

Chromatin immunoprecipitation and sequencing (ChIP-seq)

ChIP-seq on OEF-1::GFP young adults was performed as part of the modENCODE consortium project (Araya et al. 2014), and was performed as described (Niu et al. 2011; Kasper et al. 2014). Target calling analysis was performed as in Kasper et al. (2014).

Clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9

The following guide RNAs were designed to target the end of the second exon of oef-1 utilizing the fact that a 3′-GG enhances editing efficiency (Farboud and Meyer 2015): 5′-GTTTGAAGACATCTGAATGG-3′ and 5′-AGAAATATTAGAGAAATGGG-3′. The guides were cloned into p46149 (Addgene) as in Paix et al. (2014). Young adult worms were injected with Cas9 plasmid (Addgene p46168) at 50 ng/μl, each guide RNA at 75 ng/μl, and pRF4 rol-6 injection marker at 50 ng/μl. F1 rollers were screened for heterozygous deletions by PCR using primers to amplify a 719-bp region around the second exon of oef-1 (5′-AGACGAACAATCACTTGAATCAC-3′ and 5′-CATGGTGATTTCGACACAGG-3′). Sequencing confirmed a 56-bp deletion predicted to result in a stop codon after 134 amino acids. The resulting strain YL585 was backcrossed 4× prior to analysis.

DNA FISH

A probe to the 5S rDNA locus of chromosome V was prepared by PCR from genomic DNA as described (Dernburg et al. 1998). Probes were labeled using the FISH Tag DNA Green kit with Alexa Fluor 488 dye (Invitrogen, Carlsbad, CA) according to the manufacturer’s instructions, except that the isopropanol precipitation was performed overnight, the column purification steps were omitted, and the final labeled probe was resuspended in hybridization buffer. Dissection, fixation, and hybridization were performed as in Phillips et al. (2009). In the hybridization step, 100 ng of probe was used per slide. Images were acquired using a Zeiss Axioplan microscope with a 100× objective and a Zeiss AxioCam MRm camera (Zeiss, Thornwood, NY), and processed using Axiovision software.

5-ethynyl-2’-deoxyuridine (EdU) staining

The thymine-deficient Escherichia coli strain MG1693 was grown overnight at 37° in Luria broth, after which 2 ml of the overnight culture was added to 100 ml of M9 supplemented with 1% glucose, 1.25 μg/ml thiamine, 0.5 μM thymidine, 1 mM MgSO4, and 20 μM EdU. The 100 ml culture was grown for at least 24 hr at 37°, then was pelleted and resuspended in 1 ml of M9. Plates lacking peptone and containing 60 μg/ml carbenicillin were seeded with 250 μl of the resuspended bacteria and allowed to dry overnight. Worms 16 hr post-L4 were washed off plates seeded with OP50, and were washed once with M9 before being placed on MG1693-seeded plates. Worms were washed off MG1693-seeded plates after a 30-min incubation, washed twice with M9, and placed on OP50-seeded plates for 10 hr before dissection and fixation in 3% formaldehyde. For the L4 time point, arrested L1s were grown on OP50 for 36 hr before being placed on MG1693 for 30 min, and were chased on OP50 for 8 hr. Slides were freeze-cracked on a metal block embedded in dry ice and placed in −20° methanol. The next day, the slides were washed in PBST (1× PBS with 0.1% Tween) and stained with DAPI. EdU-positive cells were labeled using the Click-iT Plus EdU Alexa Fluor 488 Imaging Kit (Invitrogen) according to the manufacturer’s instructions, except that two sequential 250 μl reactions were performed per slide as in Crittenden et al. (2017). 0.375 μm Z stacks were acquired using a 40× objective, and the number of nuclei from the distal tip to the most proximal EdU-positive nucleus was counted. The file names were blinded before quantification.

Immunostaining

Adult worms 16–20 hr post-L4 were dissected to release gonads in 1.1× egg salts on silanized coverslips, and fixed with 1% formaldehyde for most antibodies. For anti-phospho-H3, anti-mCherry, and anti-SP56/anti-RME-2 staining, 3.7% formaldehyde was used. Poly-prep slides (Sigma-Aldrich, St. Louis, MO) were freeze-cracked and immediately placed in −20° methanol. Blocking was performed using 0.5–1% BSA in PBST for 30 min–1 hr at room temperature. Embryo samples were fixed with methanol and acetone, and washed with PBS as described (Strome and Wood 1983). All primary and secondary antibody incubations were performed overnight at 4°. The following antibodies and dilutions were used: 1:50 anti-OEF-1; 1:50 anti-OMA-1/2 (a gift of C. Eckmann); 1:50 anti-RME-2 (a gift of B. Grant); 1:50 SP56 (a gift of S. Strome); 1:100 anti-HIM-3 (a gift of M. Zetka); 1:200 anti-SYP-1 (a gift of A. Villeneuve); 1:1000 anti-H3K36me3 (ab9050; Abcam); 1:2000 anti-GFP (ab13970; Abcam); 1:200 anti-phospho-H3 (3H10; Upstate); 1:20 anti-PGL-1 (OIC1D4; Developmental Studies Hybridoma Bank); 1:100 anti-mCherry (PA5-34974; Pierce); and 1:500 anti-rabbit Alexa Fluor 488 or 568, anti-chicken Alex Fluor 488, anti-mouse Alexa Fluor 568, and anti-guinea pig Alexa Fluor 596 (Molecular Probes). Images were acquired as for DNA FISH experiments, except that a 40× objective was sometimes used.

L1 PGC feeding assay

Gravid adult worms were bleached and the resulting embryos were hatched overnight in M9 in the absence of food. The synchronized L1s were plated on OP50 for 7 hr. The fed larvae were collected, washed, then freeze-cracked on slides pretreated with a 10 μl drop of 0.01% poly-L-lysine (Sigma). The larvae were fixed with −20° methanol for 10 min followed by −20° acetone for 5 min, and then stained for PGL-1 as above, except PBST was used for washes and the secondary antibody incubation was performed for 1 hr at room temperature.

Small molecule FISH (smFISH)

A set of Stellaris FISH probes to oef-1 coupled to CAL Fluor Red 610 was generated by Biosearch Technologies. Fixation and hybridization were performed on embryos and dissected adults as in Ji and van Oudenaarden (2012) and Campbell and Updike (2015).

SNPC-4 foci quantification

Dissection, fixation, and staining were performed as above using the mCherry antibody, and 0.375 μm Z stacks were taken of germ cells post-transition zone. The file names were blinded, and the number of pachytene nuclei with one SNPC-4:mCherry focus vs. two SNPC-4:mCherry foci was quantified as in Kasper et al. (2014).

Sperm counts

L4s were aged for 8–9 hr before fixation in Carnoy’s solution (300 μl 100% ethanol, 150 μl chloroform, and 50 μl acetic acid). Next, 2 μl of 1 mg/ml DAPI was added to 1 ml of M9, and 12 μl of the DAPI dilution was added to the slides. 0.375 μm Z stacks were acquired for 10 spermathecae per genotype. Sperm nuclei were counted using the Cell Counter plugin in Image J.

Statistical analyses

The statistical and graphing program GraphPad Prism (version 7) was used to perform Student's t-tests and Fisher’s exact tests, as well as to calculate SD. For gene set overlaps, hypergeometric probability tests were calculated using the web-based tool http://nemates.org/MA/progs/overlap_stats.html.

Data availability

Strains and reagents generated during this project are available upon request. The mutant alleles of oef-1 (F49E8.2) will also be deposited in the Caenorhabditis Genetics Center database. The OEF-1 ChIP-seq data generated by modENCODE/modERN are publicly available at the Gene Expression Omnibus under accession number GSE107190, and at http://encodeproject.org/ and http://epic.gs.washington.edu/modERN.

Results

OEF-1 is a novel, germline-specific factor

While screening transgenic lines of GFP-tagged transcription factors generated for the modENCODE consortium project, we identified F49E8.2 as an uncharacterized nuclear protein with highly germline-specific expression (Sarov et al. 2012; Araya et al. 2014). F49E8.2 is conserved in Caenorhabditis but has no known homologs in other species. It contains a single C2H2 zinc finger (Supplemental Material, Figure S1 in File S2), which is typically a DNA-binding domain (Wolfe et al. 1999). Examination of the GFP-tagged strain (referred to as F49E8.2::GFP) revealed nuclear expression restricted to the P lineage in embryos beginning in the P4 PGC (Figure 1A). In larval stages, F49E8.2::GFP is present in all germ cells (Figure 1, B and C). In adults, F49E8.2::GFP localizes to the nuclei of germ cells in the mitotic zone through entry into meiosis (Figure 1D). Strikingly, F49E8.2::GFP expression abruptly disappears in the late pachytene region and is absent from developing oocytes (Figure 1D). Based on these features and additional observations described below, we named F49E8.2 OEF-1.

Figure 1.

Figure 1

Open in a new tab

OEF-1::GFP exhibits germline-specific expression throughout development. (A) The strain OP383 expresses OEF-1 tagged with GFP beginning in ∼28-cell embryos in the nucleus of the primordial germ cell P4 (center), with no expression seen in the 4-cell stage embryo (left). OEF-1::GFP expression is seen in the nuclei of Z2 and Z3 in a ∼100-cell embryo (right). Merged DIC and GFP channels are shown. (B) OEF-1::GFP is present in the nuclei of Z2 and Z3 in a newly hatched L1 larva. White box indicates inset. Arrows indicate Z2 and Z3. DIC, GFP, and merged channels are shown. (C) OEF-1::GFP is present in all L4 germ cell nuclei, and (D) in mitotic and meiotic germ cell nuclei through late pachytene in adults. Arrows indicate maturing oocytes. Bar, 10 μm. L, larval stage; Sp, sperm.

OEF-1 is expressed early in embryogenesis and is maternally loaded as a transcript

To examine endogenous OEF-1 expression, we generated a polyclonal antibody to OEF-1. The endogenous antibody recapitulates the expression pattern of the OEF-1::GFP transgene in L4 and adult dissected gonads (Figure 3, A and B and Figure S2 in File S2), and is lost in animals lacking OEF-1 expression (Figure S2 in File S2). In embryos, endogenous OEF-1 protein expression is first visible even earlier than OEF-1::GFP, in the P2 blastomere rather than P4 (Figure 2A). The P lineage is transcriptionally silent in embryos (Seydoux et al. 1996), which makes it unlikely that oef-1 is newly transcribed during embryogenesis. However, OEF-1 protein expression is not detectable in maturing oocytes, implying that little to no OEF-1 is maternally loaded (Figure 1D and Figure 3B). Thus, we wondered whether oef-1 transcript was maternally loaded to explain this remarkably early protein expression in P2. Using smFISH, we detected oef-1 transcripts throughout wild-type dissected gonads, including in oocytes, as well as in early embryos prior to OEF-1 protein expression (Figure 2, B and D). In addition, oef-1 transcripts become entirely restricted to the P lineage by the formation of Z2 and Z3 (Figure 2E). Taken together, these results indicate that oef-1 is maternally loaded as a transcript, with post-transcriptional regulation accounting for the delay in protein expression until the 4-cell embryonic stage.

Figure 3.

Figure 3

Open in a new tab

OEF-1 exhibits distinct expression patterns in spermatogenesis and oogenesis. (A) Dissected L4 gonad stained with OEF-1 (green) and the spermatogenesis marker SP56 (red). DAPI is in blue. (B) Dissected adult gonad stained with OEF-1 (green) and the oogenesis marker OMA-1/2 (red). DAPI, blue. (C) fem-1(hc17); OEF-1::GFP L4s show a precocious loss of OEF-1::GFP expression in proximal germ cells (n = 50/50). Arrows indicate germ cells without OEF-1::GFP expression. (D) fem-3(q23); OEF-1::GFP adults exhibit extended OEF-1::GFP expression past the gonad bend (n = 44/50). (E) OEF-1::GFP is detected in pachytene-arrested germ cells in mpk-1(ga111) mutants. Bar, 10 μm. 1°, primary spermatocytes; L, larval stage; Sp, sperm.

Figure 2.

Figure 2

Open in a new tab

OEF-1 is expressed early in embryogenesis and is maternally loaded as a transcript. (A) A 4-cell stage wild-type embryo immunostained with OEF-1 (green) and the germline-specific P granule component PGL-1 (red). OEF-1 is detected in the nucleus of the P2 blastomere. DAPI (DNA) is in blue. (B) oef-1 small molecule FISH (smFISH) probes hybridized to wild-type dissected adult gonads. oef-1 transcript is detected throughout the gonad, including in oocytes. White square indicates inset at right. (C) Dissected oef-1(vr25) adult gonads lose oef-1 signal. White square indicates inset at right. (D) Two-cell wild-type embryo with oef-1 smFISH signal (right) in both blastomeres. DAPI, left. (E) smFISH signal of an ∼100-cell wild-type embryo showing oef-1 (right) largely restricted to Z2 and Z3. DAPI, left. Bar, 10 μm.

OEF-1 exhibits distinct expression patterns in spermatogenesis and oogenesis

OEF-1 expression is detected in the proximal germ cells of L4 animals undergoing spermatogenesis, but not in the proximal germ cells of adult hermaphrodites undergoing oogenesis (Figure 1, C and D). Because wild-type hermaphrodites switch from spermatogenesis to oogenesis at the L4/adult transition, we hypothesized that OEF-1 expression is affected by gamete fate. We first costained dissected L4 gonads with OEF-1 and the sperm marker SP56 (Ward et al. 1986), and observed coexpression in the proximal gonad, indicating that OEF-1 is expressed throughout spermatogenesis, with the exception of mature sperm (Figure 1D and Figure 3A). By contrast, costaining of OEF-1 and the oocyte marker OMA-1/2 (Nousch et al. 2013) in adults confirmed that OEF-1 is not expressed during oogenesis (Figure 3B). Thus, OEF-1 is specifically excluded from oogenic germ cells, potentially through a protein degradation pathway.

To determine whether entry into oogenesis triggers loss of OEF-1 expression, we crossed the OEF-1::GFP line into gamete fate mutant backgrounds. fem-1(hc17) is a loss-of-function mutant resulting in feminized gonads that produce oocytes during L4 (Nelson et al. 1978), while fem-3(q23) is a gain-of-function mutant resulting in masculinized germlines that continue to produce sperm into adulthood (Barton et al. 1987). Notably, the fem-1; OEF-1::GFP strain showed a premature loss of OEF-1::GFP expression in proximal germ cells at L4 compared with wild-type hermaphrodite L4s undergoing spermatogenesis (n = 50/50; Figure 3C, compare to Figure 1C). Conversely, fem-3; OEF-1::GFP adults exhibited extended OEF-1::GFP expression past the gonad bend in presumptive primary spermatocytes (n = 44/50; Figure 3D, compare to Figure 1D). Finally, we utilized mpk-1(ga111) mutants, in which germ cells arrest at the late pachytene stage of meiosis I and do not initiate oogenesis (Church et al. 1995). In mpk-1; OEF-1::GFP adults, OEF-1::GFP was present in pachytene-arrested germ cells, indicating that reduction of OEF-1 expression occurs upon or after pachytene exit (Figure 3E). Together, these data indicate that the abrupt loss of OEF-1 expression at the pachytene-to-diplotene transition depends upon progression into oogenesis, and not on other temporal or spatial gonadal signals.

oef-1 mutants have an accelerated rate of germ cell progression

We next examined mutants of oef-1 to understand its potential function in the germline. Two alleles were previously available from consortia, and we generated a third using CRISPR-Cas9 gene editing (Figure S1 in File S2). The tm4563 allele contains a complex substitution (a 304-bp deletion plus a 19-bp insertion) spanning almost the entirety of exon 1 to the beginning of exon 2, while the gk205699 allele contains a point mutation that alters a proline to a serine in the sole zinc finger of OEF-1 (Figure S1 in File S2). The CRISPR allele, vr25, contains a 56-bp frameshift deletion in exon 2. The OEF-1 antibody could not detect OEF-1 in the tm4563 or the vr25 mutant background (Figure S2 in File S2). Notably, the epitope targeted by the OEF-1 antibody overlaps almost completely with the deleted amino acids in tm4563, but not entirely in vr25 (Figure S1 in File S2). Therefore, because we could be confident that vr25 was a protein null, we subsequently used it as the primary mutant for phenotypic analysis. Intriguingly, OEF-1 was detected in the gk205699 background (Figure S2 in File S2), suggesting that this point mutant retains normal expression.

As OEF-1 is expressed early in embryonic development (Figure 1A and Figure 2A), we first examined oef-1(vr25) PGCs for potential defects. After P4 divides into Z2 and Z3 during late embryogenesis, the two PGCs arrest in G2 until hatched L1s are exposed to food (Fukuyama et al. 2006). After 7 hr of feeding, only 11.1% of PGCs in wild-type larvae had undergone an initial cell division (Figure 4A). However, in 42.4% of oef-1(vr25) mutant larvae, PGCs had divided once or even twice (Figure 4A). This result was reproducible in the two other oef-1 mutant alleles (Figure S3A in File S2), and introduction of the OEF-1::GFP transgene into the oef-1(vr25) mutant background rescued early PGC divisions to wild-type levels (Figure S3B in File S2).

Figure 4.

Figure 4

Open in a new tab

oef-1 mutants have an accelerated rate of germ cell progression. (A) Starved and synchronized wild-type and oef-1(vr25) L1s were fed for 7 hr, then fixed and stained with PGL-1 to mark primordial germ cells (PGCs). The percentages of larvae with one, two, three, or four PGCs are shown. n ≥ 63 L1s per genotype. (B) Wild-type and oef-1(vr25) adult worms were fed 5-ethynyl-2’-deoxyuridine (EdU)-labeled bacteria followed by a 10-hr chase on unlabeled bacteria. The number of germ cell diameters from distal tip to the most proximal EdU-labeled nucleus was quantified. n ≥ 16 gonad arms per genotype. **** P < 0.0001, Student’s t-test. DTC, distal tip cell. Error bars represent SD. (C) Brood sizes of wild-type and oef-1(vr25) mutants at 20°. n ≥ 13 parental animals. **** P < 0.0001, Student’s t-test. Error bars represent SD. (D) The number of sperm in wild-type and oef-1(vr25) spermathecae. n = 10 spermathecae per genotype. **** P < 0.0001, Student’s t-test. Error bars represent SD.

We then assayed oef-1(vr25) adult hermaphrodites to ask whether germ cells continued to exhibit any cell cycle defects at later developmental stages. We did not observe any difference in the length of the proliferative or transition zones in oef-1(vr25) mutants by DAPI staining (Figure S4, A and B in File S2), suggesting no changes in the number of cells in mitosis or entering meiosis. We did detect a slight increase in the number of germ cells in M phase in oef-1(vr25) adults by phospho-H3 staining (8.65 vs. 7.22 positive nuclei, n ≥ 37 gonad arms per genotype, P < 0.01, Student’s t-test) (Figure S4C in File S2). These conflicting observations suggested that oef-1 mutant germ cells either exhibit a slight delay in M phase or an increased rate of progression through mitosis and into meiosis.

We therefore determined whether overall germ cell progression was altered in oef-1(vr25) mutants. We performed pulse-chase experiments using the thymidine analog EdU. EdU is incorporated into germ cells undergoing S phase, and thus labels proliferative cells (Crittenden et al. 2006). Wild-type and oef-1(vr25) mutants were exposed to EdU-labeled bacteria for 30 min followed by a 10-hr chase on unlabeled bacteria. Relative to wild-type, oef-1(vr25) mutants exhibited a significant increase in the number of cell diameters from the distal tip to the most proximally EdU-labeled germ cell, suggesting that oef-1 mutant germ cells progress at a faster rate (44.1 vs. 36.1 cell diameters, n ≥ 16 gonad arms per genotype, P < 0.0001, Student’s t-test) (Figure 4B and Figure S5A in File S2). A similar difference was also observed at the L4 stage (Figure S5B in File S2).

Consistent with an accelerated rate of germ cell progression, oef-1(vr25) mutants exhibit a 33% increase in brood size relative to wild-type (Figure 4C) (340.0 vs. 255.3, n ≥ 13 animals per genotype, P < 0.0001, Student’s t-test). The other two oef-1 mutant alleles also had increased brood sizes (Figure S6 in File S2). Since brood size is limited by sperm number, we quantified the number of sperm present in the spermathecae of young adults. We found that oef-1(vr25) mutants had a significant increase in the number of sperm generated per gonad arm compared to wild-type (Figure 4D) (156.7 vs. 113.4, n = 10 spermathecae per genotype, P < 0.0001, Student’s t-test). A faster rate of germ cell progression could explain an increase in brood size as more germ cells would be specified to sperm during the spermatogenesis window. Increased sperm production might also occur if spermatogenesis is extended due to a delay in the transition to oogenesis (Hodgkin and Barnes 1991). However, we saw no detectable alteration in the onset of oogenesis between L4 and adulthood in oef-1(vr25) mutants compared to wild-type by staining with the oogenesis marker RME-2 (Grant and Hirsh 1999) (Table S1 in File S2).

Finally, we asked whether the rate of germ cell progression was altered in maturing oocytes. We allowed wild-type and oef-1(vr25) adults to lay eggs for 4 hr and quantified the number of eggs laid. We found that more eggs were laid by oef-1(vr25) mutants, suggesting that oocyte progression is also accelerated relative to wild-type (Figure S5C in File S2). We conclude that germ cells progress more rapidly through all stages of development and differentiation in oef-1 mutants.

oef-1 mutants exhibit increased germline apoptosis that depends upon the synapsis checkpoint

To ask whether a faster rate of germ cell progression affected cell viability, we stained wild-type and oef-1 mutant adults for apoptotic germ cells using the dye SYTO 12. We detected a significant increase in the number of positively-stained germ cells in oef-1(vr25) mutants compared to wild-type (11.0 vs. 6.91, n = 23 gonad arms per genotype, P < 0.0001, Student’s t-test) (Figure 5A). This defect was also present in the other two oef-1 mutant alleles (Figure 5B), and was rescued by the OEF-1::GFP transgene (Figure S7 in File S2). The activation of either of two meiotic checkpoints can increase apoptosis above levels of physiological germline apoptosis: the synapsis checkpoint or the DNA damage checkpoint (Gartner et al. 2008). In hus-1(op241) mutants, the DNA damage checkpoint does not occur (Hofmann et al. 2002), whereas the synapsis checkpoint is not activated in pch-2(tm1458) mutants (Bhalla and Dernburg 2005). We performed an epistasis test to determine whether one of these pathways accounted for the increased apoptosis in oef-1 mutants. Apoptosis levels were restored to wild-type in pch-2; oef-1 mutants, but remained high in hus-1; oef-1 mutants (Figure 5C). This result indicates that the increased apoptosis in oef-1 mutants is due to activation of the synapsis checkpoint.

Figure 5.

Figure 5

Open in a new tab

oef-1 mutants exhibit increased germline apoptosis depending on the synapsis checkpoint. (A) Wild-type and oef-1(vr25) adults were stained with SYTO 12 to mark apoptotic germ cells and the number of positively-stained germ cells per gonad arm was quantified. n = 23 gonad arms per genotype. **** P < 0.0001, Student’s t-test. Error bars represent SD. (B) SYTO 12 quantification in other oef-1 mutant alleles. n ≥ 19 gonad arms per genotype. *** P < 0.001 and **** P < 0.0001, Student’s t-test. Error bars represent SD. (C) SYTO 12 quantification in apoptosis checkpoint mutants. n ≥ 35 gonad arms per genotype. **** P < 0.0001, Student’s t-test. Error bars represent SD. (D) Tagged SNPC-4 foci (green) were counted in pachytene nuclei in wild-type (left) and oef-1(vr25) (right) dissected gonads. SNPC-4 binds strongly to a specific region of chromosome IV in the germline and forms visible foci, making it a highly specific chromosomal marker (Kasper et al. 2014). Arrows indicate nuclei with two foci in oef-1(vr25). DAPI, blue. The percentages of nuclei with one focus or two foci are shown. n = 10 germlines per genotype, at least 20 nuclei per germline (total n ≥ 440 nuclei per genotype). Bar, 10 μm. n.s., not significant.

To determine whether oef-1 mutants had defects in chromosome synapsis, we used two chromosomal markers to assess pairing, a DNA locus near the pairing center (PC) on chromosome V (Phillips and Dernburg 2006) and a transcription factor that localizes to a domain at the opposite end from the PC on chromosome IV (Kasper et al. 2014) (see Materials and Methods). Intriguingly, the marker on chromosome IV showed delayed pairing in 40.5% of oef-1(vr25) pachytene nuclei compared to 28.2% of wild-type pachytene nuclei (n ≥ 440 nuclei per genotype, P < 0.0001, Fisher’s exact test) (Figure 5D), while the marker on chromosome V did not exhibit any detectable pairing defect (Figure S8 in File S2). Consistent with this latter result, we did not detect defects in the colocalization of the meiotic axis protein HIM-3 and the synaptonemal complex component SYP-1 (Figure S9 in File S2), and did not observe univalent chromosomes during diakinesis in oef-1 mutants (data not shown), indicating completed chromosome synapsis (Dombecki et al. 2011). Moreover, when we eliminated all germline apoptosis using ced-4(n1162) mutants, we did not observe an increased incidence of male offspring or embryonic lethality in ced-4; oef-1 double-mutants (Table S2 in File S2), as would be expected if pairing or synapsis were defective (Bhalla and Dernburg 2005). We suggest that a faster rate of progression through the proliferative zone in oef-1 mutants results in germ cell nuclei entering meiosis before they are fully prepared to undergo pairing and synapsis, and/or that accelerated progression through pachytene slightly perturbs the fidelity of these meiotic events (see Discussion).

OEF-1 associates with germline-expressed genes and localizes to autosomes

To probe the molecular function of OEF-1, we analyzed ChIP-seq data sets for the OEF-1::GFP strain in young adults that were generated by the modENCODE project (Araya et al. 2014). Significant OEF-1-binding sites were assigned to 1998 protein-coding target genes (File S1). Of these, 86.5% were germline-expressed genes (Ortiz et al. 2014) (1729 genes; 1.7× enriched, P < 6.32 × 10−253, hypergeometric probability test) (Figure 6A). Protein-coding targets were also enriched for genes with germline-biased expression (the combined data set of germline-intrinsic and oogenesis-enriched genes) (Reinke et al. 2004) (748 genes; 2.7× enriched, P < 1.74 × 10−184, hypergeometric probability test) (Figure 6A). Notably, individual gene targets display unusual OEF-1-binding profiles, with prominent association with gene bodies rather than promoter or intergenic regions (Figure 6B). In particular, target genes often displayed OEF-1 peaks at the 5′ and 3′ ends, with distributed binding above input signal along gene bodies (Figure 6B).

Figure 6.

Figure 6

Open in a new tab

OEF-1 associates with germline-expressed genes and localizes to autosomes. (A) The overlap between OEF-1::GFP coding targets, the set of germline-expressed genes (Ortiz et al. 2014), and the combined set of germline-intrinsic/oocyte-enriched genes (Reinke et al. 2004). P < 6.32 × 10−253 and P < 1.74 × 10−184, respectively, hypergeometric probability test. (B) OEF-1::GFP ChIP-seq signal along the germline-expressed genes csr-1, pgl-1, and rme-2. OEF-1 associates with the 5′ and 3′ ends of genes, as well as with gene bodies. meg-1 is an example of an X-linked germline-expressed gene for which OEF-1 signal is reduced. OEF-1::GFP ChIP-seq signal is in green, control “input” signal is in black. (C) A genome-wide view of OEF-1::GFP ChIP-seq signal (green) and input control (black). OEF-1 ChIP-seq signal is reduced along the X chromosome compared to autosomes. (D) Pachytene nuclei of the OEF-1::GFP strain stained with GFP (green) and H3K36me3 (red). Arrows indicate DAPI (blue) regions not stained with GFP or H3K36me3. Bar, 10 μm. ChIP-seq, chromatin immunoprecipitation and sequencing; Chr, chromosome.

Few germline-expressed genes reside on the X chromosome (Reinke et al. 2004), and accordingly OEF-1 targets were markedly depleted for X-linked genes (28 genes; 0.1× enriched, P < 1.585 × 10−95, hypergeometric probability test) (Figure 6C). Indeed, even the few X-linked genes that are expressed in the germline show reduced OEF-1 binding relative to autosomal germline-expressed genes, as exemplified by meg-1 (Figure 6B). To determine whether this autosomal-specific binding pattern was distinguishable in vivo, we examined dissected gonads costained with OEF-1 and the autosome-specific histone modification H3K36me3 (Rechtsteiner et al. 2010; Gaydos et al. 2012). A DAPI-stained chromosome that did not stain for H3K36me3 or OEF-1 was indeed distinguishable in pachytene nuclei (Figure 6D). We conclude that OEF-1 is an autosomal-enriched factor with preferential association with germline-expressed loci.

oef-1 partially rescues the Him phenotype of him-5, xnd-1, and him-8 mutants

Because of this striking chromosomal bias, we asked whether OEF-1 functioned in the same pathway as two other autosomal-specific factors that also lose expression in late pachytene: HIM-5 (High Incidence of Males-5) and XND-1 (X chromosome Nondisjunction-1) (Wagner et al. 2010; Meneely et al. 2012). These two germline-specific proteins act in the same genetic pathway to regulate crossover frequency on the X chromosome (Wagner et al. 2010; Meneely et al. 2012). Reduced crossover events in him-5 and xnd-1 mutants lead to broods with high percentages of embryonic lethality and males (Wagner et al. 2010; Meneely et al. 2012). Since HIM-5 and XND-1 exhibit the same autosomal localization and loss at late pachytene as OEF-1, we analyzed the broods of oef-1; him-5 and xnd-1; oef-1 double-mutants. Given that xnd-1 mutants have pleiotropic germline defects and show stochastic sterility in later generations (McClendon et al. 2016), we focused our analyses on the broods of xnd-1 M+Z- parents. We found that oef-1 partially rescued the him-5 percentage of males from 37.53 to 30.24% (P < 0.0001, Fisher’s exact test) (Table 1). In xnd-1; oef-1 double-mutants, we found that oef-1 again partially rescued the percentage of males from 6.31 to 4.11% (P = 0.0045, Fisher’s exact test) (Table 2).

Table 1. oef-1 partially rescues him-5 percentage of males.

Genotype Percentage of males (%) Total progeny
Wild-type 0.00 2576
oef-1(vr25) 0.07 4238
him-5(e1490) 37.53 2775
oef-1; him-5 30.24**** 2811
Open in a new tab

n = 13–14 parental animals per genotype.

****

P < 0.0001, Fisher’s exact test.

Table 2. oef-1 partially rescues xnd-1 percentage of males.

Genotype Percentage of males (%) Total progeny
Wild-type 0.06 3574
oef-1(vr25) 0.00 4420
xnd-1(ok709) M+Z- 6.31 1474
xnd-1 M+Z-; oef-1 4.11** 1945
Open in a new tab

n = 13–21 parental animals per genotype.

**

P < 0.005, Fisher’s exact test.

To determine whether the effect of oef-1 on the HIM-5/XND-1 pathway was specific, we analyzed him-8 mutants. HIM-8 works independently of HIM-5/XND-1 to mediate pairing and synapsis specifically of the X chromosome, and localizes to the X chromosome PC (Phillips et al. 2005). Thus, if the effect of oef-1 were specific to him-5 and xnd-1, oef-1 would not have an effect on the him-8 percentage of males. However, when we performed oef-1 RNAi on him-8(e1489) mutants and examined the percentage of males in the progeny, we found that oef-1 also partially rescued the percentage of males in him-8 broods (P < 0.0001, Fisher’s exact test) (Table 3). We conclude that OEF-1 alters the frequency of nondisjunction through a mechanism that affects both the XND-1/HIM-5 and HIM-8 pathways, and that is genetically distinct from both pathways.

Table 3. oef-1 partially rescues him-8 percentage of males.

Genotype RNAi treatment Percentage of males (%) Total progeny
Wild-type Empty vector 0.12 1691
oef-1 0.21 1933
him-8(e1489) Empty vector 40.77 1869
oef-1 34.79**** 2018
Open in a new tab

n = 6–7 parental animals per genotype.

****

P < 0.0001, Fisher’s exact test. RNAi, RNA interference.

Discussion

Here, we have characterized the expression of the highly tissue-specific novel factor OEF-1 and gained insight into its role in the C. elegans germline. OEF-1 is detectably expressed only in the germline, and is continually expressed from the P2 blastomere until the onset of oogenesis, with the exception of mature sperm. oef-1 mutants exhibit accelerated germ cell progression throughout germ cell development, which likely leads to increased apoptosis due to activation of the synapsis checkpoint. Finally, we found that OEF-1 associates broadly with many germline-expressed genes and predominantly localizes to autosomes. Together, these observations suggest that OEF-1 might play a subtle but important role in coordinating germline-specific events, and ensuring robust or buffered progression through germ cell development.

OEF-1 is expressed early and specifically in the germline

We identified OEF-1 as a germline-specific protein with strikingly early embryonic expression restricted to the P lineage. We detect endogenous OEF-1 in the nucleus of the P2 blastomere (Figure 2A) and GFP-tagged OEF-1 in the nucleus of P4 (Figure 1A). This delay between endogenous and transgene protein expression could be due to the additional time required for folding of the GFP tag (Heim et al. 1994). Because the oef-1 transcript is present throughout the gonad and in early embryonic stages (Figure 2, B and D), we suggest that a post-transcriptional repression mechanism prevents OEF-1 protein expression during oogenesis and early embryogenesis. OMA-1 and OMA-2 are two RNA-binding proteins that are critical for the oocyte-to-embryo transition and are expressed during oogenesis (Detwiler et al. 2001). Previously, the oef-1 transcript was found to immunoprecipitate with OMA-1/2 by RNA immunoprecipitation and sequencing (RIP-seq) (Spike et al. 2014), suggesting that OMA-1/2 may bind to and inhibit the translation of oef-1 during oogenesis. The transcripts of xnd-1 and him-5 also immunoprecipitated with OMA-1/2 in the same study (Spike et al. 2014), suggesting a potentially common translational repression mechanism between factors with oocyte-excluded protein expression.

Gamete fate program and the link to pachytene

Notably, we showed that the loss of OEF-1 protein expression in the late pachytene region in adult germlines occurs at and depends upon the onset of oogenesis (Figure 3). During spermatogenesis, OEF-1 is detected in primary spermatocytes, and expression does not cease until late in differentiation, when many proteins are eliminated en masse from maturing sperm (L’Hernault 2006) (Figure 1D and Figure 3A). By contrast, in the oogenic germline, OEF-1 expression abruptly becomes undetectable in late pachytene, concomitant with the onset of protein expression of early oogenesis factors and before any morphological signs of differentiation (Figure 1D and Figure 3B). The loss of other proteins (like HIM-5 and XND-1) at pachytene during oogenesis may also depend on the gamete fate program. We hypothesize that a proteosomic pathway degrades nonmaternally-loaded proteins at the pachytene stage. This degradation pathway is likely tied to the proper exit from pachytene, as inhibition of pachytene exit prevents the degradation of OEF-1 (Figure 3E). This transition point in germ cells may be critical to the proper development of oocytes; the consequence of continued expression of oocyte-excluded factors is not known. Attempts to drive OEF-1 in the proximal germline through 3′-UTR regulation (Merritt et al. 2008) did not result in detectable OEF-1 expression (data not shown), which is consistent with regulation of OEF-1 at the protein level.

OEF-1 may limit the rate of germ cell progression through development

Germ cells in oef-1 mutants progress faster than in wild-type, beginning with a precocious first division of PGCs (Figure 4A). In wild-type L1 larvae, the Z2/Z3 division occurs when a prolonged G2 arrest is released (Fukuyama et al. 2006). This release is triggered by the presence of food and depends upon activation of the genome-wide transcriptional program (Butuci et al. 2015). The checkpoint kinase CHK-1 is implicated in the timing of Z2/Z3 cell cycle reentry; in chk-1 mutants, the Z2/Z3 division occurs precociously (Butuci et al. 2015), similarly to oef-1 mutants, suggesting that OEF-1 may act in or affect this checkpoint. Even as late larvae and adults, oef-1 mutants continue to show accelerated germ cell progression, as shown by EdU labeling (Figure 4B), likely manifesting in more sperm generation during the spermatogenesis window and an increase in brood size (Figure 4, C and D). The observation that oef-1(vr25) mutants lay eggs at a faster rate (Figure S5C in File S2) suggests that even differentiated oocytes progress faster compared to wild-type.

In addition to changes in the rate of germ cell progression, oef-1 mutants exhibited an increase in germ cell death, which was eliminated upon removal of the synapsis checkpoint by pch-2 mutation (Figure 5). This checkpoint is triggered by delays or defects in synapsis (Bhalla and Dernburg 2005). Synapsis occurs progressively, originating from a specific site, the PC, on one end of each chromosome, followed by elongation of the synaptonemal complex down chromosome pairs (Rog and Dernburg 2015). We observed defective pairing in oef-1 mutants when tracking a locus far from a PC (Figure 5D), but not for a locus close to a PC (Figure S8 in File S2), suggesting that pairing along the length of homologous chromosomes is mildly delayed. However, synapsis ultimately occurs successfully in oef-1 mutants, as we found no defects in the colocalization of HIM-3 and SYP-1 (Figure S9 in File S2), bivalent chromosome number in oocytes (data not shown), or embryonic viability and incidence of males even if apoptosis is blocked (Table S2 in File S2).

From these phenotypes, we hypothesize that oef-1 mutant germ cells exhibit accelerated transit through the proliferative zone and enter meiosis before pairing can be completed, leading to a slight delay in completed pairing and synapsis of homologous chromosomes, and subsequent activation of the synapsis checkpoint. Additionally, the accelerated rate of progression seems to continue through pachytene in oef-1 mutants, and may directly affect meiotic events and thus trigger the synapsis checkpoint. Because apoptosis only occurs during oogenesis (Gartner et al. 2008), we propose that in the L4 stage, spermatogenic germ cells rapidly progress unchecked through meiosis, leading to increased sperm number and brood sizes in oef-1 mutants. In oef-1 mutant adults, the elevated apoptosis might eliminate more germ cells with unsynapsed X chromosomes upon loss of him-5, xnd-1, or him-8 activity, leading to fewer XO male progeny (Table 1, Table 2, and Table 3). Consistent with this possibility, we found that apoptosis levels in oef-1; him-5 mutants remain elevated, as in oef-1 mutants, relative to wild-type and him-5 (data not shown). Together, these phenotypes are consistent with a possible role of OEF-1 in limiting or controlling the rate at which germ cells progress through various stages of development or differentiation.

The molecular function of OEF-1

How might OEF-1 have such a subtle but influential effect on coordinating germ cell processes? Through analysis of modENCODE ChIP-seq data, we found that OEF-1 primarily localizes to germline-expressed genes (Figure 6A). However, it does not bind like most “point-source” transcription factors; OEF-1 peaks are distributed over gene bodies, rather than localized to promoters. Similar distributed binding profiles have been observed for splicing factors (Fontrodona et al. 2013) and for histone modifications (Evans et al. 2016). Moreover, we did not detect statistically significant changes in transcript abundance of OEF-1 ChIP-seq targets upon loss of oef-1 in preliminary analyses of RNA-seq data on dissected gonads (data not shown). We therefore theorize that OEF-1 does not act to modulate transcription of specific target genes, but might have some other molecular role, perhaps in RNA processing or chromatin biology. Notably, even though C2H2 zinc fingers are most commonly associated with DNA binding, they can also bind RNA or mediate protein–protein interactions (Hall 2005; Gamsjaeger et al. 2007). The C2H2 zinc finger is clearly critical to OEF-1 function, as a single damaging point mutation in the zinc finger domain is sufficient to phenocopy a protein null (Figure 5B, Figure S3A, and Figure S6 in File S2). In the future, additional studies will further delineate the molecular mechanisms that underlie the enhanced progression of germ cells in oef-1 mutants.

Supplementary Material

Supplemental material is available online at http://www.genetics.org/lookup/suppl/doi:10.1534/genetics.117.1123/-/DC1.

Click here for additional data file. (91.3KB, xlsx)
Click here for additional data file. (12MB, pdf)

Acknowledgments

We would like to thank the modENCODE and modERN consortium projects; members from the University of Chicago Institute for Genomics and Systems Biology for ChIP-seq data processing; LaDeana Hillier for target-calling analysis; and Christian Eckmann, Barth Grant, Susan Strome, Anne Villeneuve, and Monique Zetka for antibodies. Some strains were provided by the Caenorhabditis Genetics Center [National Institutes of Health (NIH) P40 OD-010440]. C.E.M. was in part supported by the NIH (T32 GM-007499) and by the K.S. & Feili Lo Foundation Graduate Fellowship for Excellence in Stem Cell Research. This work was also supported by NIH R01 GM-108663, awarded to V.R.

Author contributions: C.E.M. and V.R. conceived and designed the experiments. C.E.M. performed the experiments. C.E.M. and V.R. analyzed the data. C.E.M. and V.R. wrote the paper.

Footnotes

Communicating editor: M. Sundaram

Literature Cited

  1. Araya C. L., Kawli T., Kundaje A., Jiang L., Wu B., et al. , 2014.  Regulatory analysis of the C. elegans genome with spatiotemporal resolution. Nature 512: 400–405. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Barton M. K., Schedl T. B., Kimble J., 1987.  Gain-of-function mutations of fem-3, a sex-determination gene in Caenorhabditis elegans. Genetics 115: 107–199. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bhalla N., Dernburg A. F., 2005.  A conserved checkpoint monitors meiotic chromosome synapsis in Caenorhabditis elegans. Science 310: 1683–1686. [DOI] [PubMed] [Google Scholar]
  4. Brenner S., 1974.  The genetics of Caenorhabditis elegans. Genetics 77: 71–94. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Butuci M., Williams A. B., Wong M. M., Kramer B., Michael W. M., 2015.  Zygotic genome activation triggers chromosome damage and checkpoint signaling in C. elegans primordial germ cells. Dev. Cell 34: 85–95. [DOI] [PubMed] [Google Scholar]
  6. Campbell A. C., Updike D. L., 2015.  CSR-1 and P granules suppress sperm-specific transcription in the C. elegans germline. Development 142: 1745–1755. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Church D. L., Guan K. L., Lambie E. J., 1995.  Three genes of the MAP kinase cascade, mek-2, mpk-1/sur-1 and let-60 ras, are required for meiotic cell cycle progression in Caenorhabditis elegans. Development 121: 2525–2535. [DOI] [PubMed] [Google Scholar]
  8. Crittenden S. L., Leonhard K. A., Byrd D. T., Kimble J., 2006.  Cellular analyses of the mitotic region in the Caenorhabditis elegans adult germ line. Mol. Biol. Cell 17: 3051–3061. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Crittenden S. L., Seidel H. S., Kimble J., 2017.  Analysis of the C. elegans germline stem cell pool. Methods Mol. Biol. 1463: 1–33. [DOI] [PubMed] [Google Scholar]
  10. Dernburg A. F., McDonald K., Moulder G., Barstead R., Dresser M., et al. , 1998.  Meiotic recombination in C. elegans initiates by a conserved mechanism and is dispensable for homologous chromosome synapsis. Cell 94: 387–398. [DOI] [PubMed] [Google Scholar]
  11. Detwiler M. R., Reuben M., Li X., Rogers E., Lin R., 2001.  Two zinc finger proteins, OMA-1 and OMA-2, are redundantly required for oocyte maturation in C. elegans. Dev. Cell 1: 187–199. [DOI] [PubMed] [Google Scholar]
  12. Dombecki C. R., Chiang A. C., Kang H. J., Bilgir C., Stefanski N. A., et al. , 2011.  The chromodomain protein MRG-1 facilitates SC-independent homologous pairing during meiosis in Caenorhabditis elegans. Dev. Cell 21: 1092–1103. [DOI] [PubMed] [Google Scholar]
  13. Evans K. J., Huang N., Stempor P., Chesney M. A., Down T. A., et al. , 2016.  Stable Caenorhabditis elegans chromatin domains separate broadly expressed and developmentally regulated genes. Proc. Natl. Acad. Sci. 118: E7020–E7029. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Farboud B., Meyer B. J., 2015.  Dramatic enhancement of genome editing by CRISPR-Cas9 through improved guide RNA design. Genetics 199: 959–971. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Fontrodona L., Porta-de-la-Riva M., Moran T., Niu W., Diaz M., et al. , 2013.  RSR-2, the Caenorhabditis elegans ortholog of human spliceosomal component SRm300/SRRM2, regulates development by influencing the transcriptional machinery. PLoS Genet. 9: e1003543. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Fraser A. G., Kamath R. S., Zipperlen P., Martinez-Campos M., Sohrmann M., et al. , 2000.  Functional genomic analysis of C. elegans chromosome I by systematic RNA interference. Nature 408: 325–330. [DOI] [PubMed] [Google Scholar]
  17. Fukuyama M., Rougvie A. E., Rothman J. H., 2006.  C. elegans DAF-18/PTEN mediates nutrient-dependent arrest of cell cycle and growth in the germline. Curr. Biol. 16: 773–779. [DOI] [PubMed] [Google Scholar]
  18. Gamsjaeger R., Liew C. K., Loughlin F. E., Crossley M., Mackay J. P., 2007.  Sticky fingers: zinc-fingers as protein-recognition motifs. Trends Biochem. Sci. 32: 63–70. [DOI] [PubMed] [Google Scholar]
  19. Gartner A., Boag P. R., Blackwell T. K., 2008.  Germline survival and apoptosis (September 4, 2008) WormBook, ed. The C. elegans Research Community, WormBook, /10.1895/wormbook.1.145.1, http://www.wormbook.org. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Gaydos L. J., Rechtsteiner A., Egelhofer T. A., Carroll C. R., Strome S., 2012.  Antagonism between MES-4 and Polycomb repressive complex 2 promotes appropriate gene expression in C. elegans germ cells. Cell Rep. 2: 1169–1177. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Grant B., Hirsh D., 1999.  Receptor-mediated endocytosis in the Caenorhabditis elegans oocyte. Mol. Biol. Cell 10: 4311–4326. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Hall T. M., 2005.  Multiple modes of RNA recognition by zinc finger proteins. Curr. Opin. Struct. Biol. 15: 367–373. [DOI] [PubMed] [Google Scholar]
  23. Heim R., Prasher D. C., Tsien R. Y., 1994.  Wavelength mutations and posttranslational autoxidation of green fluorescent protein. Proc. Natl. Acad. Sci. USA 91: 12501–12504. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Hillers, K. J., V. Jantsch, E. Martinez-Perez, and J. L. Yanowitz, 2017 Meiosis (May 4, 2017), WormBook, ed. The C. elegans Research Community, WormBook, /10.1895/wormbook.1.178.1, http://www.wormbook.org. [DOI] [PMC free article] [PubMed]
  25. Hodgkin J., Barnes T. M., 1991.  More is not better: brood size and population growth in a self-fertilizing nematode. Proc. Biol. Sci. 246: 19–24. [DOI] [PubMed] [Google Scholar]
  26. Hofmann E. R., Milstein S., Boulton S. J., Ye M., Hofmann J. J., et al. , 2002.  Caenorhabditis elegans HUS-1 is a DNA damage checkpoint protein required for genome stability and EGL-1-mediated apoptosis. Curr. Biol. 12: 1908–1918. [DOI] [PubMed] [Google Scholar]
  27. Ji N., van Oudenaarden A., 2012.  Single molecule fluorescent in situ hybridization (smFISH) of C. elegans worms and embryos (December 13, 2012), WormBook, ed. The C. elegans Research Community, WormBook, /10.1895/wormbook.1.153.1, http://www.wormbook.org. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Kasper D. M., Wang G., Gardner K. E., Johnstone T. G., Reinke V., 2014.  The C. elegans SNAPc component SNPC-4 coats piRNA domains and is globally required for piRNA abundance. Dev. Cell 31: 145–158. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Kelly W. G., Schaner C. E., Dernburg A. F., Lee M. H., Kim S. K., et al. , 2002.  X-chromosome silencing in the germline of C. elegans. Development 129: 479–492. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Kimble J., Crittenden S. L., 2007.  Controls of germline stem cells, entry into meiosis, and the sperm/oocyte decision in Caenorhabditis elegans. Annu. Rev. Cell Dev. Biol. 23: 405–433. [DOI] [PubMed] [Google Scholar]
  31. Lesch B. J., Page D. C., 2012.  Genetics of germ cell development. Nat. Rev. Genet. 13: 781–794. [DOI] [PubMed] [Google Scholar]
  32. L’Hernault, S. W., 2006 Spermatogenesis (February 20, 2006), WormBook, ed. The C. elegans Research Community, WormBook, /10.1895/wormbook.1.85.1, http://www.wormbook.org.
  33. McClendon T. B., Mainpal R., Amrit F. R., Krause M. W., Ghazi A., et al. , 2016.  X chromosome crossover formation and genome stability in Caenorhabditis elegans are independently regulated by xnd-1. G3 (Bethesda) 6: 3913–3925. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Meneely P. M., McGovern O. L., Heinis F. I., Yanowitz J. L., 2012.  Crossover distribution and frequency are regulated by him-5 in Caenorhabditis elegans. Genetics 190: 1251–1266. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Merritt C., Rasoloson D., Ko D., Seydoux G., 2008.  3′ UTRs are the primary regulators of gene expression in the C. elegans germline. Curr. Biol. 18: 1476–1482. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Nelson G. A., Lew K. K., Ward S., 1978.  Intersex, a temperature-sensitive mutant of the nematode Caenorhabditis elegans. Dev. Biol. 66: 386–409. [DOI] [PubMed] [Google Scholar]
  37. Niu W., Lu Z. J., Zhong M., Sarov M., Murray J. I., et al. , 2011.  Diverse transcription factor binding features revealed by genome-wide ChIP-seq in C. elegans. Genome Res. 21: 245–254. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Nousch M., Techritz N., Hampel D., Millonigg S., Eckmann C. R., 2013.  The Ccr4-Not deadenylase complex constitutes the main poly(A) removal activity in C. elegans. J. Cell Sci. 126: 4274–4285. [DOI] [PubMed] [Google Scholar]
  39. Ortiz M. A., Noble D., Sorokin E. P., Kimble J., 2014.  A new dataset of spermatogenic vs. oogenic transcriptomes in the nematode Caenorhabditis elegans. G3 (Bethesda) 4: 1765–1772. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Paix A., Wang Y., Smith H., Lee C., Calidas D., et al. , 2014.  Scalable and versatile genome editing using linear DNAs with micro-homology to Cas9 sites in C. elegans. Genetics 198: 1347–1356. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Phillips C. M., Dernburg A. F., 2006.  A family of zinc-finger proteins is required for chromosome-specific pairing and synapsis during meiosis in C. elegans. Dev. Cell 11: 817–829. [DOI] [PubMed] [Google Scholar]
  42. Phillips C. M., Wong C., Bhalla N., Carlton P. M., Weiser P., et al. , 2005.  HIM-8 binds to the X chromosome pairing center and mediates chromosome-specific meiotic synapsis. Cell 123: 1051–1063. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Phillips C. M., McDonald K. L., Dernburg A. F., 2009.  Cytological analysis of meiosis in Caenorhabditis elegans. Methods Mol. Biol. 558: 171–195. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Porter M. Y., Koelle M. R., 2010.  RSBP-1 is a membrane-targeting subunit required by the Gαq-specific but not the Gαo-specific R7 regulator of G protein signaling in Caenorhabditis elegans. Mol. Biol. Cell 21: 232–243. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Rechtsteiner A., Ercan S., Takasaki T., Phippen T. M., Egelhofer T. A., et al. , 2010.  The histone H3K36 methyltransferase MES-4 acts epigenetically to transmit the memory of germline gene expression to progeny. PLoS Genet. 6: e1001091. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Reinke V., Gil I. S., Ward S., Kazmer S., 2004.  Genome-wide germline-enriched and sex-biased expression profiles in C. elegans. Development 131: 311–323. [DOI] [PubMed] [Google Scholar]
  47. Robert V. J., Garvis S., Palladino F., 2015.  Repression of somatic cell fate in the germline. Cell. Mol. Life Sci. 72: 3599–3620. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Rog O., Dernburg A. F., 2015.  Direct visualization reveals kinetics of meiotic chromosome synapsis. Cell Rep. 10: 1639–1645. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Sarov M., Murray J. I., Schanze K., Pozniakovski A., Niu W., et al. , 2012.  A genome-scale resource for in vivo tag-based protein function exploration in C. elegans. Cell 150: 855–866. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Seydoux G., Mello C. C., Pettitt J., Wood W. B., Priess J. R., et al. , 1996.  Repression of gene expression in the embryonic germ lineage of C. elegans. Nature 382: 713–716. [DOI] [PubMed] [Google Scholar]
  51. Spike C. A., Coetzee D., Nishi Y., Guven-Ozkan T., Oldenbroek M., et al. , 2014.  Translational control of the oogenic program by components of OMA ribonucleoprotein particles in Caenorhabditis elegans. Genetics 198: 1513–1533. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Strome S., Wood W. B., 1983.  Generation of asymmetry and segregation of germline granules in early C. elegans embryos. Cell 35: 15–25. [DOI] [PubMed] [Google Scholar]
  53. Wagner C. R., Kuervers L., Baillie D. L., Yanowitz J. L., 2010.  xnd-1 regulates the global recombination landscape in Caenorhabditis elegans. Nature 467: 839–843. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Wang J. T., Seydoux G., 2013.  Germ cell specification. Adv. Exp. Med. Biol. 757: 17–39. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Ward S., Roberts T. M., Strome S., Pavalko F. M., Hogan E., 1986.  Monoclonal antibodies that recognize a polypeptide antigenic determinant shared by multiple Caenorhabditis elegans sperm-specific proteins. J. Cell Biol. 102: 1778–1786. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Wolfe S. A., Nekludova L., Pabo C. O., 1999.  DNA recognition by Cys2His2 zinc finger proteins. Annu. Rev. Biophys. Biomol. Struct. 3: 183–212. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Click here for additional data file. (91.3KB, xlsx)
Click here for additional data file. (12MB, pdf)

Data Availability Statement

Strains and reagents generated during this project are available upon request. The mutant alleles of oef-1 (F49E8.2) will also be deposited in the Caenorhabditis Genetics Center database. The OEF-1 ChIP-seq data generated by modENCODE/modERN are publicly available at the Gene Expression Omnibus under accession number GSE107190, and at http://encodeproject.org/ and http://epic.gs.washington.edu/modERN.

Articles from Genetics are provided here courtesy of Oxford University Press

RESOURCES