Antagonistic control of Caenorhabditis elegans germline stem cell proliferation and differentiation by PUF proteins FBF-1 and FBF-2

Stem cells support tissue maintenance, but the mechanisms that coordinate the rate of stem cell self-renewal with differentiation at a population level remain uncharacterized. We find that two PUF family RNA-binding proteins FBF-1 and FBF-2 have opposite effects on Caenorhabditis elegans germline stem cell dynamics: FBF-1 restricts the rate of meiotic entry, while FBF-2 promotes both cell division and meiotic entry rates. Antagonistic effects of FBFs are mediated by their distinct activities toward the shared set of target mRNAs, where FBF-1-mediated post-transcriptional control requires the activity of CCR4-NOT deadenylase, while FBF-2 is deadenylase-independent and might protect the targets from deadenylation. These regulatory differences depend on protein sequences outside of the conserved PUF family RNA-binding domain. We propose that the opposing FBF-1 and FBF-2 activities serve to modulate stem cell division rate simultaneously with the rate of meiotic entry.


Introduction
Adult tissue maintenance relies on the activity of stem cells that self-renew and produce differentiating progeny (Morrison and Kimble, 2006). It is essential that self-renewal be balanced with differentiation to preserve the size of the stem cell pool over time. One simple mechamism achieving this balance is an asymmetric division that always produces a single stem cell daughter and a daughter destined to differentiate (Chen et al., 2016). Alternatively, tissue homeostasis can be controlled at a population level (Simons and Clevers, 2011), where some stem cells are lost through differentiation while others proliferate, with both outcomes occurring at the same frequency. Such population-level control of stem cell activity is observed in the Caenorhabditis elegans germline (Kimble and Crittenden, 2007). However, the mechanisms of population-level control of stem cell proliferation and differentiation in the adult tissues are largely unclear.
The C. elegans hermaphrodite germline is a robust system to explore the mechanisms coordinating stem cell proliferation and differentiation. It is maintained by a stem cell niche that supports about 200-250 mitotically dividing stem and progenitor cells at the distal end of the gonad (collectively called SPCs, Figure 1A,Bii). A single somatic distal tip cell serves as a stem cell niche and activates the GLP-1/Notch signaling necessary for SPC pool maintenance (Austin and Kimble, 1987), which in turn supports germline development (Hansen and Schedl, 2013). As germline stem cells move proximally away from the niche, they differentiate by entering meiotic prophase and eventually generate gametes near the proximal gonad end. Mitotic divisions of SPCs are not oriented and there doesn't appear to be a correlation between the position of cell divisions distributed throughout the SPC zone and the position of cells undergoing meiotic entry at the proximal end of the zone elegans adult hermaphrodite. In this and following images, germlines are oriented with their distal ends to the left. GLP-1/Notch signaling from the distal tip cell (blue) supports germline SPC proliferation. Progenitors enter meiosis in the transition zone. FBF-1 and FBF-2, downstream of GLP-1/ Notch, are required for SPC maintenance. Green circles, stem and progenitor cells; red diamonds, mitotically dividing cells. (B) Distal germlines dissected from adult wild type, fbf-1(lf), and fbf-2(lf) hermaphrodites and stained with anti-REC-8 (green) and anti-phospho-Histone H3 (pH3; red) to visualize the SPC zone and mitotic cells in M-phase. Germlines are outlined with the dashed lines and the vertical dotted line marks the beginning of transition zone as recognized by the 'crescent-shaped' chromatin and loss of REC-8. Scale bar: 10 mm. (C) SPC zone lengths of the wild type, fbf-1(lf) and fbf-2(lf) germlines were measured by counting germ cell diameters (gcd) spanning SPC zone. Genetic background is indicated on the X-axis and the extent of SPC zone on the Y-axis. Differences in SPC zone lengths were evaluated by one-way ANOVA with Dunnett's post-test. Data were collected from three independent experiments, with 10-15 germlines per strain per replicate. (D) Median SPC G2-phase length in different genetic backgrounds, as indicated on the X-axis. Difference in median G2 length was evaluated by one-way ANOVA with Dunnett's post-test. G2 length was estimated in three independent experiments as shown in Figure 1-figure supplement 1C, each replicate experiment involved analysis of 145-159 germlines per strain. (E) Larval germ cell doubling time in different genetic backgrounds (as indicated on the X-axis). Plotted values are individual data points and means ± SD. Difference in germ cell doubling time was evaluated by one-way ANOVA with Dunnett's post-test. Data were collected from four independent replicates as shown in Figure 1-figure supplement 1E,F, each analyzing 15-21 germlines per strain per time point (144-148 germlines per strain total). (F) Meiotic entry rate of germline progenitors in different genetic backgrounds indicated on the X-axis. Differences in meiotic entry rate between each fbf and the wild type were evaluated by one-way ANOVA with T-test with Bonferroni correction post-test. Meiotic entry rates were estimated in five independent experiments as shown in   (Crittenden et al., 2006;Fox et al., 2011;Jaramillo-Lambert et al., 2007;Maciejowski et al., 2006).
Analysis of C. elegans germline stem cell maintenance identified a number of genes affecting SPC self-renewal and differentiation (Hansen and Schedl, 2013). Genes essential for self-renewal include GLP-1/Notch and two highly similar Pumilio and FBF (PUF) family RNA-binding proteins called FBF-1 and FBF-2 (Austin and Kimble, 1987;Crittenden et al., 2002;Zhang et al., 1997). Genetic studies of stem cell maintenance led to a model where a balance of mitosis-and meiosispromoting activities maintains tissue homeostasis (Hubbard and Schedl, 2019), but the regulatory mechanism coordinating proliferative SPC activity with meiotic entry remained elusive.
Importantly, the SPC cell cycle is distinct from that of most somatic stem cells. One characteristic feature of C. elegans germline SPC cell cycle is a very short G1 phase (Fox et al., 2011;Furuta et al., 2018), reminiscent of the short G1 phase observed in mammalian embryonic stem cells (ESCs) (Becker et al., 2006;Kareta et al., 2015;White and Dalton, 2005). Mouse and human ESCs maintain robust cell division rates supported by a cell cycle with a short G1 phase, while the length of S and G2 phases is similar to that observed in differentiated somatic cells (Becker et al., 2006;Chao et al., 2019;Kareta et al., 2015;Stead et al., 2002). Despite the abbreviated G1 phase, ESCs maintain S and G2 checkpoints (Chuykin et al., 2008;Stead et al., 2002;White and Dalton, 2005). Similarly, C. elegans SPCs retain G2 checkpoints despite the shortened G1 phase (Butucˇi et al., 2015;Garcia-Muse and Boulton, 2005;Lawrence et al., 2015;Moser et al., 2009). This modified cell cycle may be due to a constant proliferative demand that both SPCs and ESCs are subject to. By contrast, this type of modified cell cycle is not observed in the adult stem cell populations that support regenerative response upon injury, such as adult mammalian bulge stem cells (hair follicle stem cells; Cotsarelis et al., 1990) or satellite cells (muscle stem cells; Schultz, 1974;Schultz, 1985;Snow, 1977) that remain in G0 or quiescent phase for the most of the adult life and only reenter cell cycle upon injury. Similarly, adult epidermal stem cells regulate their cell cycle by controlling the G1/S transition to maintain tissue homeostasis (Mesa et al., 2018).
Unlike somatic cells' G1 phase that is triggered and marked by increased amounts of cyclins E and D (Aleem et al., 2005;Guevara et al., 1999), germ cells and ESCs are characterized by a shortened G1 phase and maintain a constitutive robust expression of G1/S regulators Cyclin E and CDK2 throughout the cell cycle (Fox et al., 2011;Furuta et al., 2018;White and Dalton, 2005). Despite continuous proliferation of C. elegans SPCs, the SPC mitotic rate changes during development and in different mutant backgrounds (Kocsisova et al., 2019;Michaelson et al., 2010;Roy et al., 2016), and it is unknown how SPC division and meiotic entry rates might be altered while maintaining the cell cycle with an abbreviated G1 phase. Here, we report the mechanism through which PUF family RNA binding proteins FBF-1 and FBF-2 simultaneously change the rates of SPC cell cycle progression and meiotic entry.
PUF proteins are expressed in germ cells of many animals and are conserved regulators of stem cells (Salvetti, 2005;Wickens et al., 2002). C. elegans PUF proteins expressed in germline SPCs, FBF-1 and FBF-2, share the majority of their target mRNAs (Porter et al., 2019;Prasad et al., 2016) and are redundantly required for SPC maintenance (Crittenden et al., 2002;Zhang et al., 1997). Despite 89% identity between FBF-1 and FBF-2 protein sequences, several reports suggest that FBF-1 and FBF-2 localize to distinct cytoplasmic RNA granules and have unique effects on the germline SPC pool (Lamont et al., 2004;Voronina et al., 2012). FBF-1 and FBF-2 each support distinct numbers of SPCs (Lamont et al., 2004). Furthermore, FBF-1 inhibits accumulation of target mRNAs in SPCs, while FBF-2 primarily represses translation of the target mRNAs (Voronina et al., 2012). Some differences between FBF-1 and FBF-2 function might be explained by their association with distinct protein cofactors, as we previously found that a small protein DLC-1 is a cofactor specific to FBF-2 that promotes FBF-2 localization and function (Wang et al., 2016). Despite the fact that several repressive mechanisms have been documented for PUF family proteins (Quenault et al., 2011), it is relatively understudied how the differences between PUF homologs are specified. Here, we sought to take advantage of the distinct SPC numbers maintained by individual FBF proteins to understand how they regulate the dynamics of SPCs cell cycle and meiotic entry and to probe the functional differences between FBFs.
Elaborating on the general contribution of PUF proteins to stem cell maintenance, we describe here that FBF-1 and FBF-2 have antagonistic effects on the rates of germline SPC cell cycle and meiotic entry. We find that FBFs regulate core cell cycle machinery transcripts along with transcripts required for differentiation to coordinately affect both transcript classes. FBF-1 requires CCR4-NOT deadenylation machinery, while FBF-2 functions independently of CCR4-NOT and might protect target mRNAs from deadenylation. These distinct functions of FBFs are determined by the protein regions outside of the conserved PUF homology domain. The opposing regulation of SPC cell cycle and differentiation by FBFs allows stem cells to simultaneously modulate cell division rate and meiotic entry.

Results
FBF-1 and FBF-2 differentially modulate cell division and meiotic entry of C. elegans germline SPCs During tissue maintenance, stem cells adjust their proliferative activity and differentiation rate through diverse regulatory mechanisms, including RNA-binding protein-mediated post-transcriptional regulation. We hypothesized that two paralogous RNA-binding proteins FBF-1 and FBF-2 differentially regulate germline stem cell mitotic rate and meiotic entry in C. elegans, resulting in distinct effects on the size of stem and progenitor cell (SPC) zone. We first determined how the extent of SPC zone was affected by loss-of-function mutations of each fbf. SPCs were marked by staining for the nucleoplasmic marker REC-8 ( Figure 1A,B; Hansen et al., 2004), and the extent of SPC zone was measured by counting the number of cell rows positive for REC-8 staining in each germline. Consistent with a previous report (Lamont et al., 2004), we observed that the SPC zone of fbf-1(ok91, loss-of-function mutation, lf) (~15 germ cell diameters, gcd; Figure 1Bi) is smaller than that of the wild type (~20 gcd, Figure 1Bii), whereas the SPC zone of fbf-2(q738, loss-of-function mutation, lf) (~25 gcd, Figure 1Biii) is larger than that of the wild type ( Figure 1B,C). The differences in the length of SPC zone between fbf single mutants and the wild type are consistently observed in animals through the first day of adulthood (Figure 1-figure supplement 1A).
To test whether the differences in the lengths of germline SPC zone between fbf mutants and the wild type result from changes in the rate of cell division, we compared cell cycle parameters in each genetic background. We started with measuring the M-phase index (the percentage of SPC zone cells in M phase) following immunostaining for the SPC marker REC-8 and the M-phase marker phospho-histone H3 (pH3, Figure 1B). We found that the mitotic index of fbf-1(lf) was significantly higher than that of the wild type (by 54%, Figure 1-figure supplement 1B). By contrast, the mitotic index of fbf-2(lf) was significantly lower than that of the wild type (by 42%; Figure 1-figure supplement 1B). These results suggested that loss of FBF-2 might reduce SPC proliferation. We also considered the possibility that the loss of FBF-1 might accelerate progression of SPCs through the cell cycle. However, as described below, this hypothesis was rejected. Since C. elegans germline stem cells have abbreviated G1 and extended G2 phases (Fox et al., 2011), we tested whether the G2-phase duration is affected differentially by loss of function mutation of each fbf. Using phospho-histone H3 immunostaining and 5-ethynyl-2'-deoxyuridine (EdU) pulse, we estimated a median G2 length by determining when 50% of pH3-positive cells become EdU-positive (Figure 1-figure supplement 1C). We found that the median G2 length of fbf-2(lf) is significantly greater than that of the wild type, suggesting that loss of FBF-2 results in slower progression through the G2-phase of the cell cycle (by 25%; Figure 1D). By contrast, the median G2 length of fbf-1(lf) is not significantly different from that of the wild type ( Figure 1D). We conclude that FBF-2 accelerates SPC cell cycle by facilitating the G2-phase progression.
Since mutation of fbf-1 did not affect the length of G2 phase, we tested whether percentage of SPCs in S phase is affected by this mutation. We determined percent SPCs labeled by EdU during a 30-min pulse (Fox et al., 2011) and found a minor increase in S-phase index in fbf-1(lf) compared to the wild type ( Figure 1-figure supplement 1D). These results refute the interpretation that fbf-1(lf) mutation causes faster cell cycle progression.
To directly estimate the rate of germ cell division in wild type and fbf mutants, we assayed germ cell proliferation during larval development before the onset of meiotic differentiation. In C. elegans, two primordial germ cells in L1 larvae proliferate to produce germline stem cell pools of 20-30 cells in L2 larval stage within 20 hr (Hansen et al., 2004;Hirsh et al., 1976;Pepper et al., 2003b;Figure 1-figure supplement 1E). We found that fbf-1(lf) did not affect the rate of germ cell division, while fbf-2(lf) dramatically reduced germ cell accumulation (Figure 1-figure supplement 1F). Exponential fits revealed that fbf-2(lf) significantly increased SPC doubling time from 6.1 hr to 8.1 hr ( Figure 1E). By contrast, there was no significant difference in germ cell proliferation rate between fbf-1(lf) and the wild type. We conclude that the cell division rate is decreased in fbf-2(lf) and unaffected in fbf-1(lf).
Despite the same SPC cell division rate, the SPC zone of fbf-1(lf) is smaller than that of the wild type, suggesting a possibility that fbf-1(lf) might result in faster meiotic entry. Conversely, compared to the wild type, fbf-2(lf) maintains a relatively larger SPC population but with slower proliferation, suggesting that the rate of meiotic entry in fbf-2(lf) might be slower than in the wild type. To test these possibilities, we determined the rate of meiotic entry in each genetic background. Animals were continuously EdU labeled and stained for EdU and REC-8 at three time points. The number of germ cells negative for REC-8 but positive for EdU were scored at each time point and the rate of meiotic entry was estimated from the slope of the plotted regression line as in Figure 1-figure supplement 1G. We found that fbf-1(lf) results in a significantly increased rate of meiotic entry compared to the wild type (by 31%; Figure 1F), whereas fbf-2(lf) results in a significantly reduced rate of meiotic entry (by 18%; Figure 1F). We conclude that FBF-2 stimulates meiotic entry while FBF-1 restricts meiotic entry.
In summary, mutations in fbf-1 and fbf-2 differentially influence both SPC cell cycle and meiotic entry rate, suggesting FBF proteins have antagonistic effects on SPC proliferation and differentiation. FBF-1 promotes a more quiescent stem cell state characterized by a slower rate of meiotic entry, while FBF-2 promotes a more activated stem cell state characterized by faster rates of both cell cycle and meiotic entry. While FBF's effects on transcripts regulating meiotic entry such as gld-1 have been documented previously (Crittenden et al., 2002;Brenner and Schedl, 2016), FBF regulation of the cell cycle has been unexplored. We hypothesized that FBFs control SPC progression through cell cycle by regulating B-type cyclin mRNAs. Table 1. fbf-2(lf) enhances the overproliferation phenotype of glp-1(gf) at 15˚C. All animals were maintained at 15˚C. For each genotype, after scoring sterility as the lack of embryos in the uterus, germlines of sterile animals were dissected and stained with DAPI, anti-REC-8 antibodies, and anti-phospho-histone H3 antibodies for evaluation of overproliferation. All animals were analyzed at 1 day after L4 stage. Tum, a complete tumorous germline. Pro, proximal overproliferation phenotype. PUF-mediated repression of cyclin B limits accumulation of germline SPCs in fbf-2 Cyclin B/Cdk1 kinase, also known as M-phase promoting factor, triggers G2/M transition in most eukaryotes (Lindqvist et al., 2009). Four cyclin B family genes provide overlapping as well as specific mitotic functions in C. elegans (van der Voet et al., 2009). We hypothesized that the slower G2phase and lower M-phase index of fbf-2(lf) SPCs results from translational repression and reduced steady-state levels of four cyclin B family transcripts mediated by the remaining germline-expressed PUF-family proteins, including FBF-1. We addressed this hypothesis in two ways. First, we tested whether mutation of FBF-binding elements (FBEs) in the 3'UTR of cyb-2.1 mRNA would result in translational derepression of cyb-2.1. Second, we assessed whether derepression of cyb-2.1 in fbf-2 (lf) would lead to accumulation of more SPCs by uncoupling PUF-mediated regulation of cell division and meiotic entry. FBFs repress their target mRNAs by binding to the FBF-binding elements (FBEs; UGUxxxAU) in the 3'UTRs Crittenden et al., 2002;Merritt and Seydoux, 2010). Four mRNAs encoding Cyclin B family members co-purify with FBF proteins and three of them contain predicted FBEs in their 3'UTRs (Porter et al., 2019;Prasad et al., 2016). Since cyb-2.1 mRNA is found in complex with FBFs across multiple experimental conditions and contains more canonical FBE sites in its 3'UTR than the other cyclin B transcripts, we selected cyb-2.1 for further analysis. If FBFs repress translation of cyb-2.1 by binding to FBEs, mutation of FBEs would cause derepression of CYB-2.1 protein. To test this prediction, we established a transgenic animal 3xflag::cyb-2.1(fbm), expressing 3xFLAG::CYB-2.1 under the control of 3'UTR with mutated FBEs (ACAxxxAU); as a control, a transgenic animal expressing 3xflag::cyb-2.1(wt) with wild-type FBEs was also established ( Figure 2A). Quantification of transgene transcript levels by qPCR suggested that steady-state transcript levels of 3xflag::cyb-2.1(fbm) were~4.5 fold greater than those of 3xflag::cyb-2.1(wt), suggesting that FBEs affect steady-state transcript levels ( Figure 2B). By immunoblotting, we found that the expression of 3xFLAG::CYB-2.1 protein was increased~1.4 fold in 3xflag::cyb-2.1(fbm) animals compared to 3xflag::cyb-2.1(wt), suggesting that the presence of FBEs decreases protein production from cyb-2.1 mRNA ( Figure 2C). The abundance of cyclin family proteins is subject to extensive post-translational control (Langenfeld et al., 1997;Peters, 2002), which might account for a larger difference observed at the level of transcript. C. elegans SPCs express five PUF-family proteins that cluster into three groups based on sequence similarity: FBF-1/-2, PUF-8, and PUF-3/-11 (Ariz et al., 2009;Crittenden et al., 2002;Haupt et al., 2020;Lamont et al., 2004;Stumpf et al., 2008). Each of the three PUF groups has a distinct RNA-binding specificity Koh et al., 2009;Opperman et al., 2005), so it is likely that FBEs in cyb-2.1 3'UTR are predominantly recognized and regulated by FBFs. However, we cannot exclude the possibility that FBEs in the cyb-2.1 3'UTR mediate association with PUFs other than FBF-1/-2. We conclude that cyb-2.1 expression in SPCs is downregulated by PUF proteins recruited to the FBEs.
Loss-of-function mutation in fbf-2 is associated with slower SPC cell division in conjunction with a slower SPC meiotic entry rate. We hypothesized that both these phenotypes might be mediated by reduced translation of key FBF target mRNAs that are required for cell cycle progression or meiotic entry respectively. We aimed to disrupt coordinate repression of cell cycle-and differentiationrelated transcripts in fbf-2(lf) by introducing 3xflag::cyb-2.1(fbm) transgene that produces increased levels of corresponding mRNA and protein. fbf-2(lf) with its slow cell division rate provides a sensitized background for testing the effects of cyclin B deregulation on cell cycle dynamics since it is not clear whether SPC cell cycle rate could be accelerated beyond that of the wild type. We hypothesized that the slower SPC cell cycle in fbf-2(lf) is caused by PUF-mediated repression of cyclin B-family mRNAs. If any cyclin B-family gene can promote SPC proliferation, disrupting translational repression of a single cyclin B-family transcript in fbf-2(lf) would rescue the slow cell cycle phenotype and accelerate SPC cell division. To test this hypothesis, we estimated the doubling time of larval germ cells after crossing the 3xflag::cyb-2.1fbm and 3xflag::cyb-2.1wt transgenes into fbf-2(lf) genetic background. We found that the SPC doubling time of fbf-2(lf); 3xflag::cyb-2.1fbm was significantly shorter than that of fbf-2(lf) ( Figure 2D). By contrast, there was no significant difference in the doubling time between fbf-2(lf); 3xflag::cyb-2.1wt and fbf-2(lf) ( Figure 2D). We expected that overexpression of 3xFLAG::CYB-2.1 in fbf-2(lf) genetic background would not affect SPC meiotic 3xFLAG-tagged CYB-2.1(wt) with wild type FBF binding elements (FBEs, UGUxxxAU) in 3'UTR and 3xFLAG-tagged CYB-2.1(fbm) with FBF binding elements mutated (ACAxxxAU). (B) qRT-PCR of 3xflag::cyb-2.1 and unc-54 transcripts in 3xflag::cyb-2.1(wt) and 3xflag::cyb-2.1(fbm) worms using actin (act-1) as a normalization control. Data represent two replicates, values are average ± SD. Differences in levels were evaluated by T-test; asterisks mark statistically significant difference (p<0.001). (C) Immunoblot analysis of 3xFLAG::CYB-2.1 protein levels in 3xflag::cyb-2.1(wt) and 3xflag::cyb-2.1(fbm) worms using a-tubulin as a loading control. Data represent three replicates, values are average ± SD. Differences in levels were evaluated by T-test; asterisk marks statistically significant difference (p<0.05). (D) Larval germ cell doubling time in different genetic backgrounds (as indicated on the Figure 2 continued on next page entry rate. Indeed, there was no significant difference in SPC meiotic entry among fbf-2(lf), fbf-2(lf); cyb-2.1wt, and fbf-2(lf); cyb-2.1(fbm) ( Figure 2E).
We conclude that disrupting PUF-mediated regulation of CYB-2.1 has uncoupled cell cycle dynamics from the rate of meiotic entry in the fbf-2(lf) background, supporting the model of coordinated regulation of cell cycle and meiotic entry by FBFs. These results further suggest that meiotic entry rate and cell cycle progression are regulated through distinct subsets of FBF targets, rather than meiotic entry rate being a direct consequence of how fast SPCs are generated by cell divisions.

FBF-1 function requires CCR4-NOT deadenylase complex
One mechanism of PUF-dependent destabilization of target mRNAs is through recruitment of CCR4-NOT deadenylase that shortens poly(A) tails of the targets (Quenault et al., 2011). CCR4-NOT deadenylase is a complex that includes three core subunits: two catalytic subunits CCR-4/CNOT6/6L and CCF-1/CNOT-7/8 and a scaffold subunit LET-711/CNOT1, which are highly conserved in C. elegans and humans ( Figure 3A; Nousch et al., 2013). Although multiple PUF family proteins, including FBF homologs in C. elegans, interact with a catalytic subunit of CCR4-NOT in vitro, the contribution of CCR4-NOT to PUF-mediated repression in vivo is still controversial (Suh et al., 2009;Weidmann et al., 2014). We hypothesized that the enlarged germline SPC zone in fbf-2(lf) mutant results from FBF-1-mediated destabilization and translational repression of target mRNAs required for meiotic entry achieved through the activity of CCR4-NOT deadenylase. If so, knockdown of CCR4-NOT in fbf-2(lf) genetic background would lead to derepression of target mRNAs in SPCs and a decrease of the length of SPC zone.
First, we measured the extent of SPC zone after RNAi-mediated knockdown of core CCR4-NOT subunits, and found that CCR4-NOT RNAi dramatically shortened the SPC zone in fbf-2(lf) compared to the control RNAi (p<0.01; Figure 3B). By contrast, the lengths of SPC zones in the wild type and fbf-1(lf) animals were not significantly affected by CCR4-NOT knockdown ( Figure 3B). We note that the observed effects of CCR4-NOT knockdown are milder than those reported by a recent publication Nousch et al., 2019; these differences might result from the later developmental stage affected . Plotted values are individual data points and means ± SD. Difference in germ cell doubling time was evaluated by one-way ANOVA with Dunnett's post-test. (E) Meiotic entry of progenitors in different genetic backgrounds. Time course of accumulating EdU-labeled, REC-8 negative germ cells in different genetic backgrounds in one biological replicate (the data are representative of two biological replicates, each analyzing 6-7 germlines per strain per time point, up to 41-42 germlines per strain total). X-axis displays time points when animals were dissected for staining for EdU and REC-8. Y-axis indicates the number of EdU-positive cells that are negative for REC-8. Plotted values are means ± SD. (F) Distal germlines dissected from the fbf-2(lf), fbf-2(lf); cyb-2.1(fbm) and fbf-2(lf); cyb-2.1(wt) animals and stained with anti-REC-8 (green) and anti-pH3 (red). Germlines are outlined with dashed lines and the vertical dotted line marks the beginning of transition zone. Scale bar: 10 mm. (G) The extent of SPC zone in the fbf-2(lf), fbf-2(lf); cyb-2.1(fbm) and fbf-2(lf); cyb-2.1(wt) genetic backgrounds. Plotted values are individual data points and means ± SD. Differences in SPC zone lengths were evaluated by one-way ANOVA with Dunnett's post-test; asterisks mark statistically significant difference (p<0.0001). Data were collected from two independent experiments and 14-19 germlines were scored for each genotype per replicate. (H) The extent of SPC zone in the fbf-2(lf); cyb-2.1(fbm) after cyb-2.1(RNAi) compared to the empty vector RNAi control. Plotted values are individual data points and means ± SD. Differences in SPC zone lengths were evaluated by T-test; asterisks mark statistically-significant difference (p<0.0001). Data were collected from two independent experiments and 44 independent germlines were scored for each condition. (I) Immunoblot analysis of 3xFLAG::CYB-2.1 protein levels in 3xflag::cyb-2.1fbm after cyb-2.1(RNAi) compared to the empty vector RNAi control. Tubulin was used as a loading control. (B-I) All experiments were performed at 15˚C. The extent of SPC zone after knocking down CCR4-NOT subunits in the wild type, fbf-1(lf) and fbf-2(lf) genetic backgrounds. Genetic backgrounds and RNAi treatments are indicated on the X-axis and the average size of SPC zone ± SD is plotted on the Y-axis. Differences between CCR4-NOT RNAi and the empty vector RNAi control were evaluated by one-way ANOVA. Asterisks mark the group with significant changes in SPC zone length after CCR4-NOT knockdown, p<0.01. Data were collected from three independent experiments. N, the number of hermaphrodite germlines scored. (C) Distal germlines of rrf-1(lf); fbf-2(lf) expressing a GFP::Histone H2B fusion under the control of the htp-2 3'UTR after the indicated RNAi treatments. Germlines are outlined with dashed lines and vertical dotted lines indicate the beginning of the transition zone. All images were taken with a standard exposure. Scale bar: 10 mm.  by our knockdown approach. Our findings suggest that CCR4-NOT is required for FBF-1-mediated regulation of germline SPC zone length, but does not significantly contribute to FBF-2 function.
CCR4-NOT knockdown might disrupt FBF-1 regulatory function or FBF-1 protein expression and localization. To distinguish between these possibilities, we determined the abundance of endogenous FBF-1 after ccf-1(RNAi) by immunoblot using tubulin as a loading control. We found that FBF-1 protein abundance does not decrease after CCF-1 knockdown compared to the control (Figure 3figure supplement 1C,D). Immunostaining for the endogenous FBF-1 showed that in control germlines FBF-1 localized in foci adjacent to perinuclear P granules (Figure 3-figure supplement 1E) as previously reported (Voronina et al., 2012). Upon CCF-1 knockdown, FBF-1 foci were still observed next to P granules ( Figure 3-figure supplement 1F). Therefore, we conclude that CCR4-NOT is not required for FBF-1 expression and localization, and CCR4-NOT knockdown specifically disrupts FBF-1 function.
In summary, we conclude that CCR4-NOT is required for FBF-1, but not FBF-2-mediated regulation of target mRNA and germline SPC zone length. We further predicted that FBF-1 localizes together with CCR4-NOT to the same RNA-protein complex in SPCs.

FBF-1 and FBF-2 differentially impact target mRNAs polyadenylation
Since a knockdown of CCR4-NOT deadenylase produces distinct effects on FBF-1-and FBF-2-mediated target repression, we hypothesized that FBFs differentially affect deadenylation of target mRNAs. To test this hypothesis, we compared the length of the poly(A) tail of two FBF target mRNAs among the wild type, fbf-1(lf), and fbf-2(lf) genetic backgrounds by Poly(A) tail (PAT)-PCR. We selected the targets associated with both cell cycle (cyb-2.1; Kershner and Kimble, 2010;Porter et al., 2019;Prasad et al., 2016) and meiotic entry (htp-1; Merritt and Seydoux, 2010) and used an mRNA not associated with FBFs (unc-54) as a control. RNA samples were extracted from animals of glp-1 (gain-of-function, gf) mutant background, which produces germlines with a large PAT-PCR assays using RNA samples extracted from fbf-1(lf); glp-1(gf), glp-1(gf), and fbf-2(lf); glp-1(gf) revealed that the poly(A) tail distributions of cyb-2.1 and htp-1 mRNAs in fbf-2(lf) were both shifted to shorter lengths compared to the wild type background ( Figure 5A,B,D,E). Conversely, in fbf-1(lf) background, the poly(A) tail profiles of cyb-2.1 and htp-1 mRNAs were shifted to longer lengths ( Figure 5A,B,D,E). By contrast, the poly(A) tail of the control unc-54 mRNA did not decrease in fbf-2(lf) background or increase in fbf-1(lf) compared to the wild type ( Figure 5C,F). We conclude that FBF-1 promotes deadenylation of FBF target mRNAs and FBF-2 protects the targets from deadenylation. This is consistent with weaker effects of CCR4-NOT knockdown on FBF target regulation in the genetic backgrounds where FBF-2 is present (wild type and fbf-1(lf), Figure 3).
Cytoplasmic deadenylation of mRNA frequently leads to its decay (Mugridge et al., 2018). To test whether differential polyadenylation of FBF targets in fbf-1(lf) and fbf-2(lf) resulted in changes in their steady-state amounts relative to the wild type, we compared the mRNA abundance of several FBF targets among fbf-1(lf); glp-1(gf), glp-1(gf), and fbf-2(lf); glp-1(gf) genetic backgrounds by qPCR ( Figure 5-figure supplement 1B). We determined steady-state levels of both meiotic entry-associated transcripts him-3, htp-1, and htp-2 (Merritt and Seydoux, 2010) and cell cycle regulators cyb-1, cyb-2.1, cyb-2.2 and cyb-3 (Kershner and Kimble, 2010;Porter et al., 2019;Prasad et al., 2016), and used unc-54 as a control. All transcript levels were normalized to a housekeeping gene actin (act-1). The steady-state abundance of FBF targets in fbf-2(lf) decreased relative to the wild type and the decrease was significant for all transcripts except him-3 ( Figure 5-figure supplement  1B). The levels of most FBF targets in the fbf-1(lf) were not significantly different from those in the wild-type background, except for cyb-2.1 that accumulated to significantly lower levels ( Figure 5figure supplement 1B). By contrast, the abundance of unc-54 mRNA did not decrease in either genetic background. We conclude that shorter poly(A) tails of FBF targets in fbf-2(lf) are associated with lower mRNA accumulation for many targets.
Three variable regions outside of FBF-2 RNA-binding domain are necessary to prevent cooperation with CCR4-NOT Our findings suggest that FBF-1-mediated SPC maintenance depends on the CCR4-NOT deadenylase complex, while FBF-2 can function independent of CCR4-NOT. Since FBF proteins are very similar in primary sequence except for the four variable regions (VRs, Figure 6A), we next investigated whether the VRs were necessary for FBF-2-specific maintenance of germline SPCs and prevented FBF-2 dependence on CCR4-NOT. We previously found that mutations/deletions of the VRs outside of FBF-2 RNA-binding domain (VR1, 2 and 4, Figure 6A) produced GFP::FBF-2(vrm) protein with a disrupted localization and compromised function (Wang et al., 2016). We hypothesized that these three VRs might contribute to FBF-2-specific effects on the extent of SPC zone as well as prevent FBF-2 from cooperating with CCR4-NOT.

Discussion
Our results support three main conclusions that advance our understanding of how PUF family FBF proteins modulate rates of cell cycle progression and meiotic entry in C. elegans germline stem and progenitor cells. First, FBF proteins simultaneously adjust SPC cell cycle and meiotic entry rates through regulation of FBF target mRNAs affecting each process. Second, FBF-mediated repression of cyclin B affects SPC cell division rate. Third, distinct effects of FBF homologs on their target mRNAs and SPC development are mediated by differential cooperation of FBFs with deadenylation machinery. In turn, activation of deadenylation machinery by FBFs depends on the protein sequences outside of the conserved PUF RNA-binding domain. Collectively, our results support a model where two paralogous FBF proteins achieve complementary effects on SPC cell division and meiotic entry through distinct regulatory mechanisms (Figure 8). Table 3. Proximity ligation assay detects association of FBF-2(vrm) with CCR4-NOT complex component CCF-1. PLA foci density was determined in maximal intensity projections of confocal image stacks of germline SPC zones of the indicated strains. Reported values are mean ± SD derived from three independent biological replicates reared at 24˚C. PLA densities were compared by Student's t-test. N, number of germline images analyzed.

FBFs affect the rates of both stem cell mitotic divisions and meiotic entry
Here, we provide evidence that loss-of-function mutation of fbf paralogs change the rates of both cell cycle and meiotic entry in C. elegans germline SPC. We find that slow proliferation of SPCs in fbf-2(lf) is associated with a slower rate of progenitor meiotic entry (differentiation), while the progenitors of fbf-1(lf) mutant have a faster rate of meiotic entry ( Figure 1D-F). We propose that cell  (Lamont et al., 2004, not pictured). In fbf-2(lf), FBF-1 is overexpressed, and both cell division and meiotic entry rates are slow. This results in an increased total number of SPCs in the adult (larger green circle). In fbf-1(lf), FBF-2 is overexpressed and meiotic entry rate is enhanced, reducing overall numbers of SPCs (smaller green circle). In the double fbf-1(lf) fbf-2(lf) mutant (not pictured), adult SPCs are lost to differentiation. (B) FBF-1 and FBF-2 bind the same target mRNAs, and each can promote SPC maintenance. FBF-1 cooperates with CCR4-NOT deadenylase and robustly represses target mRNAs (Crittenden et al., 2002;Brenner and Schedl, 2016) to restrict the rate of germline stem cell meiotic entry. FBF-2 protects mRNAs from deadenylation; FBF-2-dependent regulation is associated with leaky repression of protein synthesis and sustains the wild-type rates of SPC cell division and meiotic entry. Differential cooperation of FBFs with CCR4-NOT is determined by their variable regions outside of the RNA-binding domain.
cycle and meiotic entry are coordinately affected by FBF-mediated control of target mRNAs relevant to each process (Figure 8). Antagonistic post-transcriptional regulation by FBF-1 and FBF-2 promotes the wild-type coordination of cell cycle progression and meiotic entry. Loss of either fbf changes the stem cell system dynamics, which in turn results in changes of SPC zone size in the individual fbf mutants compared to the wild type ( Figure 8A). Nevertheless, each individual FBF is able to maintain germline stem cells, in a state that is either closer to quiescent (FBF-1), or primed for differentiation (FBF-2). Although this model is likely oversimplified, it provides a framework for future exploration of factors controlling SPC dynamics. Previous research identified SYGL-1 and LST-1 as key transcriptional targets of GLP-1/NOTCH that promote germline stem cell fate (Kershner et al., 2014). SYGL-1 and LST-1 are cofactors of both FBFs that promote FBF-dependent regulation (Haupt et al., 2019;Shin et al., 2017). The abundance of SYGL-1 and LST-1 impacts the number of uncommitted stem cells and thus the overall size of SPC zone (Haupt et al., 2019;Shin et al., 2017). Remarkably, both SYGL-1 and LST-1 are also FBF targets (Kershner and Kimble, 2010;Kershner et al., 2014;Prasad et al., 2016), and it would be interesting to investigate whether changes in SPC zone length in fbf mutants are associated with altered levels or pattern of expression of SYGL-1 and/or LST-1.
Slow meiotic entry rate in fbf-2(lf) likely results from enhanced translational repression of FBF targets that regulate differentiation; indeed, slower accumulation of the FBF target GLD-1 has been documented in this genetic background (Brenner and Schedl, 2016). In a similar fashion, mutations of FBF targets gld-2 and gld-3 lead to a decrease in meiotic entry rate and to accumulation of excessive numbers of SPCs (Eckmann et al., 2004;Fox and Schedl, 2015). Conversely, higher meiotic entry rate of fbf-1(lf) SPCs might be explained by partial derepression of FBF targets. FBF-2 represses FBF target transcripts in fbf-1(lf) background, while sequestering them in large cytoplasmic aggregates (Voronina et al., 2012). However, this repression is less effective than that exerted by FBF-1 since partial derepression of GLD-1 has been previously observed in fbf-1(lf) (Brenner and Schedl, 2016;Crittenden et al., 2002).
We find that FBF-2 promotes SPC cell division by facilitating progression through the G2-phase of cell cycle ( Figure 1D). Thus, SPCs of the fbf-2(lf) mutant are characterized by longer median G2phase length. By contrast, the G2-phase of fbf-1(lf) SPCs is the same as in the wild type, even though this genetic background shows an increase in the mitotic index ( Figure 1D, Figure 1-figure supplement 1B). There are several possible explanations for a higher mitotic index in fbf-1(lf) background. One is slow progression through the M-phase of the cell cycle. We tested for activation of mitotic checkpoints in fbf-1(lf) genetic background, but found no difference with the wild type in the prevalence of inactivated NCC-1/CDK-1 (pTyr15) (data not shown). Another possibility for the increase in mitotic index is the difference in the proportion of cycling-competent versus non-cycling cells in fbf-1(lf). The proximal SPC zone contains non-cycling cells in meiotic S-phase, estimated to comprise 30-40% of total SPC zone cells in the wild-type germlines (Crittenden et al., 2006;Jaramillo-Lambert et al., 2007;Fox et al., 2011). Faster meiotic entry rate of fbf-1(lf) SPCs might be associated with faster progenitor transit through the meiotic S-phase. As a result, the number of non-cycling premeiotic cells (and consequently the total number of SPC zone cells) would be lower, leading to an inflated SPC mitotic index. We could not address whether fbf-1(lf) germlines have a lower number of progenitors in meiotic S-phase since there are no specific molecular markers for this developmental stage. Finally, we find that disruption of FBF-mediated regulation of a single B-type cyclin in slowly proliferating and slowly differentiating fbf-2(lf) SPCs is sufficient to disturb stem cell homeostasis, to promote faster cell cycle, and to result in excessive SPC accumulation (Figure 2). This observation is consistent with a model where FBFs regulate the rates of SPC cell division and meiotic entry by affecting separate sets of target mRNAs.
In vitro and in vivo studies have revealed that FBFs recognize the same FBE 3'UTR motif and bind largely same mRNAs Prasad et al., 2016;Porter et al., 2019). Based on this, we speculate that antagonistic regulation of FBF target mRNAs results from FBFs competing for the same transcripts. Alternatively, since some FBF targets have multiple FBEs, FBFs might cobind the transcripts thus subjecting mRNAs to combinatorial regulation. Despite FBF localization to distinct RNA granules, diffuse cytoplasmic fraction of FBF proteins might be sufficient for such coregulation. Either mechanism would result in FBF target mRNAs differentially responding to the presence of a single or both FBFs, and would explain how change-of-function mutants such as FBF-2vrm and FBF-1(FBF-2vr4) are able to rescue loss-of-function of the non-cognate fbf ( Table 4).

Regulation of cyclin B by PUF-family proteins in stem cells
PUF mRNA targets have been studied in multiple organisms including C. elegans, mouse, and human identifying thousands of target mRNAs (Chen et al., 2012;Galgano et al., 2008;Kershner and Kimble, 2010;Morris et al., 2008;Porter et al., 2019;Prasad et al., 2016). One highly conserved group of PUF regulatory targets is related to the control of cell cycle progression. In several developmental contexts, stem cells undergo rapid G1/S transitions and spend an extended time in G2, as observed for C. elegans germline stem cells as well as for mouse and human embryonic stem cells (Fox et al., 2011;Lange and Calegari, 2010;Orford and Scadden, 2008). Human, mouse, and C. elegans PUF proteins repress Cip/Kip family cyclin-dependent kinase inhibitors (Kalchhauser et al., 2011;Kedde et al., 2010;Lin et al., 2019). This repression was found to be important for cell cycle progression of human and mouse cells through G1 (Kedde et al., 2010;Lin et al., 2019). Additionally, mitotic cyclins B and A are among the core targets of PUF proteins across species including nematode FBFs (Kershner and Kimble, 2010;Porter et al., 2019;Prasad et al., 2016), Drosophila Pumilio (Asaoka-Taguchi et al., 1999), human and mouse PUM1 and PUM2 (Chen et al., 2012;Galgano et al., 2008;Hafner et al., 2010;Morris et al., 2008), and yeast Puf proteins (Gerber et al., 2004;Wilinski et al., 2015). Cyclin B regulation by PUFs contributes to cell cycle control of Drosophila embryonic cell divisions (Asaoka-Taguchi et al., 1999;Vardy and Orr-Weaver, 2007) and to the control of meiotic resumption during Xenopus and zebrafish oocyte maturation (Kotani et al., 2013;Nakahata et al., 2003;Ota et al., 2011). Here, we find that PUF-mediated regulation of mitotic cyclins affects cell cycle dynamics in the germline stem cells of C. elegans. A recent report suggests that regulation of cyclin B by PUFs is also observed in mouse embryonic stem cells (Uyhazi et al., 2020). It would be interesting to investigate whether PUF-family proteins might affect cell cycle of embryonic stem cells through control of cyclin B expression. The slow cell cycle phenotype of the fbf-2(lf) mutant SPCs was rescued by introduction of a CYB-2.1 transgene with mutated FBE elements ( Figure 2D), suggesting that the levels of B-type cyclins are limiting for SPC progression through cell cycle in this genetic background. Among the B-type cyclins, CYB-3 has emerged as a major G2 regulator in C. elegans germline stem cells (Lara-Gonzalez et al., 2019). Since cyb-3 lacks canonical FBE elements in its 3'UTR, testing whether FBFs might regulate CYB-3 levels to control SPC division rate will require identification of the relevant noncanonical binding sites.
PUF proteins FBF-1 and FBF-2 might employ both CCF-1-dependent and CCF-1-independent regulatory modes (Suh et al., 2009). Here, we find that FBF-1 and FBF-2 differentially cooperate with CCR4-NOT deadenylation machinery in C. elegans germline SPCs (Figure 3). Multiple lines of evidence suggest that FBF-1's function in vivo is supported by the CCR4-NOT deadenylation. First, the size of germline SPC zone maintained by FBF-1 in the absence of FBF-2 is significantly reduced by a knock-down of CCR4-NOT deadenylase components ( Figure 3B). Second, FBF-1-mediated repression of FBF target reporter in vivo requires CCR4-NOT deadenylase ( Figure 3C,D). By contrast, SPC zone maintained by FBF-2 and repression of reporter by FBF-2 in the absence of FBF-1 are not affected by CCR4-NOT component knock down. Taken together, these observations provide genetic evidence that CCR4-NOT promotes FBF-1 function in germline SPCs. The increase in FBF-1 protein levels that we observed after knocking down the CCR4-NOT subunit ccf-1 (Figure 3-figure supplement 1C) might result from the relief of FBF-1 auto-regulation (Lamont et al., 2004). Third, both endogenous FBF-1 and GFP::FBF-1 colocalize with a core CCR4-NOT subunit 3xFLAG::CCF-1 in vivo by co-immunostaining ( Figure 4A,C). Additionally, an in vivo test of protein interaction between GFP::FBF-1 and 3xFLAG::CCF-1 using proximity ligation assay detects positive signal suggesting that these proteins reside in the same complex ( Figure 4D). By contrast, there's significantly less in vivo colocalization and proximity between GFP::FBF-2 and 3xFLAG::CCF-1. These data are consistent with the model that FBF-1 and FBF-2 form distinct RNP complexes, of which FBF-1 complex preferentially includes CCR4-NOT deadenylase. Finally, we assessed the FBF target poly(A) tail length in the nematodes mutant for each fbf, and found that the poly(A) tail length of FBF targets cyb-2.1 and htp-1 was relatively shorter in fbf-2(lf) background than in wild type and fbf-1(lf). By contrast, fbf-1(lf) mutation resulted in a shift of poly(A) tail distributions to longer lengths compared to the wild type ( Figure 5). We conclude that FBF-1 selectively cooperates with deadenylation machinery to promote translational repression of target mRNAs ( Figure 8B). Conversely, FBF-2 protects the target mRNAs from deadenylation and promotes repression of target mRNAs by a deadenylation-independent mechanism.
The two FBF proteins are 91% identical in primary sequence (Zhang et al., 1997). If FBFs have distinct abilities to engage deadenylation machinery, what are the features of FBF-2 that prevent it from cooperating with CCR4-NOT? PUF RNA-binding domain is sufficient for a direct interaction with the CCF-1 subunit of CCR4-NOT and its homologs in multiple species, including C. elegans (Goldstrohm et al., 2006;Hook et al., 2007;Kadyrova et al., 2007;Suh et al., 2009;Van Etten et al., 2012). However, protein sequences outside of the well-structured RNA-binding domain can promote PUF-induced deadenylation and are hypothesized to function either through improved recruitment of CCR4-NOT complex or through allosteric activation of CCR4-NOT (Webster et al., 2019). We find that the Variable Region (VR) sequences outside of the RNA-binding domain of FBF-1 and FBF-2 play a key role in determining whether these proteins are able to cooperate with CCR4-NOT ( Table 4). Mutations of three VRs (VR1, 2, and 4) in FBF-2 result in a protein that now cooperates with CCR4-NOT, suggesting that these regions are necessary to prevent the wild-type FBF-2 from engaging with the deadenylase (Figure 6E,F). Conversely, swapping the VR4 of FBF-2 onto FBF-1 renders the chimeric protein FBF-1(FBF-2vr4) insensitive to CCR4-NOT knockdown, indicating that VR4 of FBF-2 is sufficient to prevent cooperation with CCR4-NOT ( Figure 7D). By contrast, swapping VR3 residing within FBF-2 RNA-binding domain into FBF-1 does not affect the FBF-1(FBF-2vr3) chimera's cooperation with CCR4-NOT, supporting the importance of protein sequences outside of the RNA-binding domain for cooperation with CCR4-NOT. Overall, we conclude that the protein regions outside of the conserved PUF RNA-binding domain regulate the repressive action mediated by each PUF protein homolog. As a result, distinct sequences flanking the RNA-binding domain may lead to differential preference of regulatory mechanisms exerted by individual PUF-family proteins ( Figure 8B). Identifying the sequences outside of FBF-1 RNA-binding domain that promote its cooperation with CCR4-NOT remains a subject for future studies.
In conclusion, our results suggest a new mechanism regulating stem cell mitotic activity in conjunction with meiotic entry rate or differentiation in the C. elegans germline through antagonistic regulation of key mRNA targets by PUF family FBF proteins. Complementary activities of FBF-1 and FBF-2 combine to fine tune SPC proliferation and meiotic entry coordinately regulating both processes. PUF proteins are conserved stem cell regulators in a variety of organisms, and their control of target mRNAs that affect proliferation and differentiation is widespread as well. The future challenge will be to determine whether PUF-dependent RNA regulation in other stem cell systems might be modulated to adjust stem cell division rate along with changing the rate of differentiation.
A single-copy insertion of each GFP-tagged FBF transgene and CYB-2.1 transgenes was generated by homologous recombination into universal Mos1 insertion site on chromosome III after Cas9induced double-stranded break (Dickinson et al., 2013;Wang et al., 2016). Similarly, single-copy insertion of 3xFLAG-tagged CCF-1 was generated by targeting universal Mos1 insertion site on chromosome II. Transgene insertion into universal Mos1 insertion sites was confirmed by PCR.

Germline SPC zone measurement
C. elegans were synchronized by bleaching, and hatched L1 larvae were plated on NNGM plates with OP50 bacteria or RNAi culture, grown at specified temperatures and harvested at varying time points depending on the experiment. For the time course of SPC zone length, L1 larvae of fbf-1(lf), fbf-2(lf) and the wild type (N2) were cultured at 24˚C and dissected at 46 hr (early adults that have initiated oogenesis), 52 hr (adults) and 63 hr (older adults) time points. L1 larvae of fbf-2(lf); cyb-2.1fbm, fbf-2(lf); cyb-2.1wt and fbf-2(lf) were grown at 15˚C for 5 days until adult stage. In all other SPC zone quantification assays, L1 larvae of all worm strains were cultured at 24˚C and dissected for staining at 52 hr time point. Gonads were dissected and stained for mitotic marker REC-8 (Hansen et al., 2004), and the length of SPC zone in each germline was measured by counting the number of germ cell rows positive for REC-8 staining before transition zone, ending with the last row fully occupied by REC-8-positive cells. Measuring the extent of progenitor zone by counting the number of cell rows positive for mitotic marker REC-8 provides a reliable estimate of progenitor cell numbers and correlates with progenitor cell numbers in the key genotypes including wild type, fbf-1 (lf), and fbf-2(lf) (R2 = 0.779; Figure 1-figure supplement 2).

M phase index measurement
Synchronous cultures of wild type (N2), fbf-1(lf) and fbf-2(lf) L1 larvae were cultured at 24˚C for 52 hr. Gonads were dissected and stained for a mitotic marker REC-8 and an M phase marker phospho-Histone H3 (pH3). Primary and secondary antibodies are described in Supplementary file 2. M phase index was calculated by dividing the number of pH3-positive SPCs by the number of REC-8positive SPCs. Percent differences in mitotic indices were calculated as for G2-phase length or differentiation rate.

EdU labeling
S-phase index, G2-phase length and differentiation rate of germ cells were measured by feeding C. elegans EdU-labeled bacteria for varying amounts of time at 24˚C (Crittenden et al., 2006;Fox et al., 2011;Kocsisova et al., 2018), with three or four replicates. EdU bacteria plates were prepared by diluting an overnight culture of thymine deficient MG1693 E. coli (The Coli Genetic Stock Center; Yale University) 1/25 in 1% glucose, 1 mM MgSO 4 , 5 mg/mL thymine, 6 mM thymidine and 20 mM EdU in M9 minimal media. This culture was grown at 37˚C for 24 hr, pelleted by centrifugation, resuspended in 10 mL M9 minimal media and plated on 60 mm NNGM plates. Worm strains were synchronized by bleaching, hatched overnight and L1 larvae were cultured on OP50 plates at 24˚C for~50 hr to reach young adult stage, when they were exposed to EdU-labeled bacteria. After feeding for specified time, worms were picked off EdU plates, dissected on poly-L-lysine treated slides, frozen on dry ice and fixed in ice-cold 100% methanol for 1 min followed by 2% paraformaldehyde/100 mM K 2 HPO 4 pH 7.2 for 5 min. Next, slides were blocked in PBS/0.1% BSA/0.1% Tween-20 (PBS-T/BSA) for 30 min at room temperature. Samples were incubated with primary antibodies against either phospho-Histone H3 or REC-8 (Supplementary file 2). After overnight incubation with primary antibody slides were washed 3 Â 10 min with PBS-T/BSA and incubated with secondary antibody for 1.5 hr at room temperature. Secondary antibodies were either Alexa Fluor 594-conjugated goat anti-mouse IgG (H+L) or Alexa Fluor 594-conjugated goat anti-rabbit IgG (H+L), respectively (Supplementary file 2). After incubation with secondary antibody slides were washed 4 Â 15 min with PBS-T/BSA. Next, the Click-iT reaction was performed according to the manufacturer's instructions (Molecular Probes) with the exception that 2 Â 30 min Click-iT reactions were performed to increase the signal of the Alexa 488 dye. After incubation with the second Click-iT reaction, slides were washed 4 Â 15 min with PBS-T/BSA. Vectashield with DAPI (Vector Laboratories) was added to each sample before cover-slipping. Germline images were captured as z-stacks spanning the thickness of each germline using a Leica DM5500B microscope. For each replicate time point 7-14 germlines were scored and the data represent three or five biological replicates. Nuclei were manually counted using the Cell Counter plug-in in Fiji (Schindelin et al., 2012) and the Marks-to-Cells R script (Seidel and Kimble, 2015) was used to remove multiply-counted nuclei.

S-phase index
Synchronous cultures of young adult nematodes were fed EdU-labeled bacteria for 30 min at 24˚C. Germ cells were co-labeled with anti-REC-8 antibody. The S-phase index was calculated as the fraction of REC-8-positive cells that were also EdU-positive.

G2 length and differentiation rate analysis
To calculate the median duration of G2-phase animals were fed EdU and collected at 30 min intervals from 0 to 3.5 hr. Germ cells were co-labeled with anti-pH3 antibody and the fraction of M-phase nuclei that have also completed G2-phase was determined by dividing the number of pH3 and EdUpositive nuclei by the total number of pH3 positive nuclei. The percent pH3 and EdU-positive nuclei was plotted on the y-axis against the duration of the EdU label on the x-axis and the data were fit to a sigmoidal varying slope curve using GraphPad Prism software, with top and bottom constrained at 100 and 0 respectively (Figure 1-figure supplement 1B). The t 50 value of the sigmoidal doseresponse model was taken as the median duration of G2-phase, or the time at which 50% of pH3 positive cells are also EdU positive.
The rate of meiotic entry was calculated by feeding the worms EdU-labeled bacteria for 3, 6 or 10 hr and co-labeling the germ cells with anti-REC-8 antibody. The number of nuclei that had entered meiosis in the time since EdU exposure based on being REC-8 negative and EdU positive were counted for each time point. The number of nuclei that entered meiosis was plotted on the y-axis and the duration of the EdU label was plotted on the x-axis in GraphPad Prism software. Linear regression analysis was used to calculate the slope, which corresponds to the number of cells that have entered meiosis per hour. Percent differences in G2-phase length or differentiation rate were calculated through subtracting the mean value of median G2-phase length or differentiation rate of each fbf mutant from that of the wild type followed by dividing by the value of the wild type.

Larval germ cell proliferation rate
Germ cell proliferation assays of fbf mutants were performed using strains where germ cells were identified by expression of CRISPR-tagged PGL-1::GFP. The nematodes were synchronized by bleaching and hatched L1s were fed on OP50 NNGM plates. The pgl-1::gfp, fbf-1(ok91); pgl-1::gfp and fbf-2(q738); pgl-1::gfp strains were grown at 24˚C. At 17 and 21 hr time points after the start of feeding samples were taken to image germ cell accumulation in L2 larvae. The data represent four biological replicates, and 15-21 germlines of each strain were scored per time point in each replicate. To analyze CYB-2.1 effect on larval germ cell proliferation, the fbf-2(q738); pgl-1::gfp, fbf-2 (q738); cyb-2.1(wt); pgl-1::gfp, and fbf-2(q738); cyb-2.1(fbm); pgl-1::gfp strains were grown at 15˚C. At 41 and 46 hr time points after the start of feeding germ cells were imaged in L2 larvae, and the data represent five biological replicates with 13-20 germlines scored per time point in each replicate. The number of germ cells were scored in each germline by counting cells containing P granules. The doubling rate of larval germ cells was estimated by exponential fits performed independently for each biological replicate.

RNAi treatment
The following RNAi constructs were used: ccr-4, let-711 (Kamath and Ahringer, 2003), ccf-1 (cenix:341 c12; Sö nnichsen et al., 2005) and cyb-2.1 (genomic CDS) in pL4440 (Timmons and Fire, 1998). Empty vector pL4440 was used as a control in all RNAi experiments. All RNAi constructs were verified by sequencing. RNAi plates were prepared as previously described (Wang et al., 2016) and synchronously hatched L1 larvae were plated directly on RNAi plates, except for let-711 and ccf-1(RNAi), where L1 larvae were initially grown on OP50 plates and transferred to RNAi plates at the L3/L4 stage. For CCR4-NOT knockdown, L1 larvae were grown at 24˚C for 52 hr before analysis; for cyb-2.1 knockdown, worms were grown at 15˚C for 120 hr before analysis. The effect of cyb-2.1(RNAi) was confirmed by western blot of 3xFLAG::CYB-2.1. The effectiveness of CCR4-NOT RNAi treatments was assessed by scoring sterility (Figure 3-figure supplement 1) or embryonic lethality (Supplementary file 3) in the F1 progeny of the fed animals.
RNA extraction and PAT-PCR glp-1(gf) and fbf-1(lf); glp-1(gf) C. elegans were synchronized using bleach, hatched L1s were cultured at 25˚C and worms were harvested after 52 hr. A subset of animals from each batch were dissected and germlines were stained with DAPI, anti-REC-8, and antiphospho-histone H3 to assess abnormal proliferation. Although the documented phenotype of glp-1 (ar202) at 25˚C is ectopic germline proliferation in the proximal region, we have often observed formation of full germline tumors in all three genotypes in our culture conditions ( Figure 5-figure supplement 1A). The RNA samples were prepared from the cultures with >77% full germline tumors. Worm pellets were washed twice with 1x M9 to remove OP50 bacteria, weighed, flash-frozen using dry ice/ethanol slurry and stored at À80˚C. Three biological replicates were analyzed for each genetic background. Total RNA was isolated using Trizol (Invitrogen) and Monarch Total RNA miniprep kit (NEB). RNA concentration was measured using Qubit (Thermo Fisher). PAT-PCR for the FBF target cyb-2.1 and control tbb-2 was performed using a Poly(A) Tail-Length Assay Kit (Thermo Fisher). Briefly, G/I tailing, reverse transcription, PCR amplification and detection were performed following the kit protocol. Each G/I tailing reaction used 1 mg total RNA. During PCR amplification, 1 ml of diluted RT sample was used in each PCR reaction and a two-step PCR program was used: 94˚C for 2 min, (94˚C for 10 s, 60˚C for 1 min 30 s) x 35 cycles, 72˚C for 5 min. PCR products were assessed using 6 or 8% polyacrylamide gels (made with 29:1 Acrylamide/Bis Solution, Bio-Rad) electrophoresis. PCR products were visualized with SYBR Gold stain (Invitrogen) and recorded using ChemiDoc MP Imaging System (Bio-Rad). Poly(A) tail length distributions were compared using densitometry analysis in ImageJ following background subtraction and normalization. The peak locations were identified as maxima of average normalized intensity profiles.

qPCR
The qPCR data represent three biological replicates of glp-1(gf), four biological replicates of fbf-2 (lf); glp-1(gf), and six biological replicates of fbf-1(lf); glp-1(gf). cDNA was synthesized using the SuperScript IV reverse transcriptase (Thermo Fisher) using 2 mg RNA template per each 20 mg cDNA synthesis reaction. Quantitative PCR reactions were performed in technical triplicates per each input cDNA using iQ SYBR Green Supermix (Bio-Rad) with cDNA diluted 1:10 as template. Primers for htp-1, htp-2, him-3, and act-1 were as described (Merritt and Seydoux, 2010). Primers for cyb-1, cyb-2.1, cyb-2.2, cyb-3, and unc-54 were designed to span exon-exon boundaries to avoid amplification of residual genomic DNA. Abundance of each mRNA in two fbf mutants relative to the wild type was calculated using the comparative DDCt method (Pfaffl, 2001) with actin act-1 as a reference gene. After the mRNA abundance of each tested gene was normalized to act-1, the fold change values from replicates were averaged. Finally, fold change values of each tested gene in glp-1(gf); fbf-1(lf) and glp-1(gf); fbf-2(lf) genetic backgrounds were scaled to the average value in glp-1 (gf) in which the mRNA abundance was set to 1. Differences in mRNA abundance between glp-1(gf) and fbf-2(lf); glp-1(gf) were evaluated by one-way ANOVA statistical tests with Student's t-test posttests. Since all FBF targets analyzed in this paper are germline-enriched in the adult nematodes, the mRNA abundances in whole worm lysates reflect their abundances in the germline.

Immunolocalization and image analysis
For all immunostaining experiments, C. elegans hermaphrodites were dissected and fixed as previously described (Wang et al., 2016). All primary antibody incubations were overnight at 4˚C and all secondary antibody incubations were for 1.5 hr at room temperature. For colocalization analysis of endogenous FBF-1 and 3xFLAG::CCF-1, dissected gonads of flag::ccf-1 were stained with anti-FBF-1 (Rabbit) and anti-FLAG primary antibodies (Mouse) (Supplementary file 2). For colocalization analysis of GFP::FBFs and 3xFLAG::CCF-1, dissected gonads of 3xflag::ccf-1; gfp::fbf-2 and 3xflag::ccf-1; gfp::fbf-1 were stained with rabbit anti-GFP and mouse anti-FLAG primary antibodies (Supplementary file 2). Secondary antibodies were Goat anti-Mouse or Goat anti-Rabbit. Germline images were acquired using Zeiss 880 confocal microscope. Localization of FBF granules relative to CCF-1 granules were analyzed in a single confocal section per germline with four to six germ cells in SPC zone by Pearson's correlation coefficient analysis using the JACoP plugin of ImageJ. For each worm strain, four to eight independent germline images were analyzed and Pearson's correlation coefficient values were averaged.

Proximity ligation assay (PLA)
PLA was performed on dissected C. elegans gonads following a modified Duolink PLA Protocol as described (Day et al., 2020). Fixation was as previously described (Wang et al., 2016). Blocking steps included incubation in 1xPBS/0.1% Triton-X-100/0.1% BSA for 2 Â 15 min at room temperature, in 10% normal goat serum for 1 hr at room temperature, and in Duolink blocking buffer for 1 hr at 37˚C. Primary anti-GFP and anti-FLAG antibodies were diluted in Duolink diluent (Supplementary file 2). After overnight incubation with primary antibodies at 4˚C, 1:5 dilutions of PLUS and MINUS Duolink PLA Probes were added to each slide and incubated at 37˚C for 1 hr. Next, slides were incubated at 37˚C for ligation (for 30 min) and amplification (for 100 min) steps and finally mounted with Duolink Mounting medium with DAPI. Images were acquired using Zeiss 880 confocal microscope. The ImageJ 'Analyze Particles' plugin was used to quantify PLA foci in germline images.

FBF target reporter regulation assay
Reporter transgene with GFP fused to Histone H2B and the 3' untranslated region (UTR) of htp-2 (Merritt et al., 2008;Merritt and Seydoux, 2010) was crossed into rrf-1(lf), rrf-1(lf)/hT2; fbf-1(lf) and rrf-1(lf); fbf-2(lf) genetic backgrounds. RNAi targeting let-711 and ccf-1 were conducted on these reporter strains as described above. The effectiveness of RNAi treatments was assessed by scoring F1 embryo lethality. RNAi-treated worms were dissected and fluorescent germline images were acquired on a Leica DFC300G camera attached to a Leica DM5500B microscope with a standard exposure. Percentage of germlines that exhibited target reporter derepression in the SPC zone was scored for each strain.

Immunoblotting
Synchronous cultures of C. elegans were collected at the adult stage by washing in 1xM9 and centrifugation and worm pellets were lysed by sonication. Proteins from worm lysates were separated using SDS-PAGE gel electrophoresis and transferred to a 0.45 mm PVDF membrane (EMD Millipore) as previously described (Ellenbecker et al., 2019). Primary and secondary antibodies are described in Supplementary file 2. Blots were developed using Luminata Crescendo Western HRP substrate (EMD Millipore) and visualized using ChemiDoc MP Imaging System (Bio-Rad).