Antagonistic control of Caenorhabditis elegans germline stem cell proliferation and differentiation by PUF proteins FBF-1 and FBF-2
- Division of Biological Sciences, University of Montana, United States
Abstract
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 (Crittenden et al., 2006; Fox et al., 2011; Jaramillo-Lambert et al., 2007; Maciejowski et al., 2006).
Figure 1 with 2 supplements see all
FBF-1 and FBF-2 differentially regulate the extent of germline stem and progenitor cell (SPC) zone.
(A) Schematic of the distal germline of C. 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 μm. (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 Figure 1—figure supplement 1G, each analyzing 5–7 germlines per strain per time point (89–94 germlines per strain total). (B–F) All experiments were performed at 24°C. Plotted values are individual data points and means ± SD. Asterisks mark statistically significant differences (****, p<0.0001; **, p<0.01; *, p<0.05).
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 meiosis-promoting 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 (Butuči 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.
Slow germ cell proliferation together with a delay in meiotic entry enhance the penetrance of germline tumor formation in sensitized genetic backgrounds (Hubbard and Schedl, 2019; Killian and Hubbard, 2005). We tested whether fbf-2(lf) enhances the overproliferative phenotype of the weak glp-1 gain-of-function allele, glp-1(ar202). We found that fbf-2(lf) is a strong enhancer of glp-1(gf) since 97% fbf-2(lf); glp-1(gf) animals have tumorous germlines with 24% germlines showing complete tumors, even at the permissive temperature of 15°C (Table 1).
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.
| Genotype | Normal germline, % | Tum, % | Pro, % | N |
|---|---|---|---|---|
| fbf-2(q738lf) | 100 | 0 | 0 | many |
| fbf-1(ok91lf) | 100 | 0 | 0 | many |
| glp-1(ar202gf) | 100 | 0 | 0 | 231 |
| fbf-2(lf); glp-1(gf) | 3 | 24 | 74 | 104 |
| fbf-1(lf); glp-1(gf) | 99 | 0 | 1 | 414 |
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.
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 G2-phase 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 (Bernstein et al., 2005; 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 (Bernstein et al., 2005; 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.
Figure 2
FBF-mediated repression of cyclin B limits accumulation of germline progenitor cells.
(A) Schematic representation of transgenes encoding 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 α-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 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. (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 μm. (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.
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 differentiation-related 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 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).
Accelerated SPC cell cycle without a change in SPC meiotic entry rate is expected to result in accumulation of SPCs and an increase of SPC zone length. To test this prediction, we measured the extent of SPC zone of fbf-2(lf); 3xflag::cyb-2.1fbm and fbf-2(lf); 3xflag::cyb-2.1wt. We found that the SPC zone of fbf-2(lf); 3xflag::cyb-2.1fbm (~32 gcd, Figure 2Fiii) is significantly larger than that of the fbf-2(lf) (~26 gcd, Figure 2Fi, G, p<0.0001). By contrast, there is no significant difference in the length of SPC zone between the fbf-2(lf); 3xflag::cyb-2.1wt and fbf-2(lf) (Figure 2Fii, G). To test whether the expansion of SPC zone in fbf-2(lf); 3xflag::cyb-2.1fbm results from overexpression of cyb-2.1, we measured the extent of SPC zone following knockdown of cyb-2.1 by RNAi. We found that the SPC zone of fbf-2(lf); 3xflag::cyb-2.1fbm after cyb-2.1(RNAi) became significantly shorter (~26 gcd) compared to the control RNAi (~31 gcd; Figure 2H). Depletion of CYB-2.1 was confirmed by immunoblot for FLAG::CYB-2.1 after RNAi of cyb-2.1 compared to the control (Figure 2I).
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.
Figure 3 with 1 supplement see all
CCR4-NOT deadenylase complex promotes FBF-1 function in germline SPCs.
(A) Schematic of CCR4-NOT deadenylase complex in humans and C. elegans; highlighted are the scaffold (yellow) and catalytic (orange) subunits targeted by RNAi in this study. (B) 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 μm. (D) Percentage of germlines showing expression of GFP::H2B fusion extended to the distal end in the indicated genetic backgrounds and knockdown conditions. Plotted values are means ± SD. Data were collected from three independent experiments. N, the number of germlines scored. Efficiencies of RNAi treatments were confirmed by sterility (Figure 3—figure supplement 1B) or embryonic lethality (Supplementary file 3). (B–D) All experiments were performed at 24°C.
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 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.
Next, we tested whether CCR4-NOT knockdown disrupts FBF-1-mediated translational repression in SPCs. One FBF target mRNA associated with meiotic entry is htp-2, a HORMA domain meiotic protein (Merritt and Seydoux, 2010). Translational regulation of a transgenic reporter encoding GFP::Histone H2B fusion under the control of htp-2 3’UTR recapitulates FBF-mediated repression in germline SPCs, where GFP::H2B::htp-2 3’UTR production is inhibited in the wild type and both fbf-1 and fbf-2 single mutant gonads but is strongly derepressed in fbf-1 fbf-2 double mutant gonads (Merritt and Seydoux, 2010). If CCR4-NOT is required for fbf-1 activity, then fbf-2(lf) after CCR4-NOT subunit RNAi should show the same phenotype as fbf-1(lf) fbf-2(lf), or derepression of the reporter. We performed CCR4-NOT RNAi in the rrf-1(lf) background to preferentially direct the RNAi effects to the germline and avoid effects on the somatic cells (Kumsta and Hansen, 2012; Sijen et al., 2001). We observed derepression of the reporter in SPCs of 63–69% germlines of rrf-1(lf); fbf-2(lf) genetic background (Figure 3C,D). By contrast, derepression of the reporter was observed only in 3–5% of rrf-1(lf) and rrf-1(lf); fbf-1(lf) genetic backgrounds (Figure 3D; Figure 3—figure supplement 1A). These data suggest that the CCR4-NOT deadenylase complex is necessary for FBF-1-mediated repression of target mRNAs in germline SPCs, but is dispensable for FBF-2 regulatory function. fbf-1 fbf-2 double mutant hermaphrodites are sterile (Crittenden et al., 2002). We observed significantly increased sterility upon CCR4-NOT knockdown in rrf-1(lf); fbf-2(lf) compared to the rrf-1(lf) and rrf-1(lf); fbf-1(lf) (Figure 3—figure supplement 1B). Like fbf-1 fbf-2 double mutants, the majority of rrf-1(lf); fbf-2(lf) sterile germlines following CCR4-NOT knockdown failed to initiate oogenesis resulting in germline masculinization (data not shown). These observations suggest that CCR4-NOT is required for fbf-1 activity.
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 3—figure 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 colocalizes with CCR4-NOT in germline SPCs
Using co-immunostaining of endogenous FBF-1 or GFP::FBF-1 and 3xFLAG::CCF-1 followed by Pearson’s correlation coefficient analysis based on Costes’ automatic threshold (Costes et al., 2004), we found that both endogenous FBF-1 and GFP::FBF-1 foci colocalize with 3xFLAG::CCF-1 foci in SPC cytoplasm (Figure 4A,C; Figure 4—figure supplement 1A,B). By contrast, the colocalization between GFP::FBF-2 and 3xFLAG::CCF-1 is significantly less robust (Figure 4B,C). As an alternative metric of colocalization, we used proximity ligation assay (PLA) that can detect protein-protein interactions in situ at the distances < 40 nm (Fredriksson et al., 2002). PLA was performed in 3xflag::ccf-1; gfp::fbf-1, 3xflag::ccf-1; gfp::fbf-2, and 3xflag::ccf-1; gfp animals using the same antibodies and conditions for all three protein pairs. We observed significantly more dense PLA signals in 3xflag::ccf-1; gfp::fbf-1 than in the control (Figure 4D; p<0.0001, Table 2). By contrast, PLA foci density in mitotic germ cells of 3xflag::ccf-1; gfp::fbf-2 was not different from the control (Figure 4D; Table 2), although the expression of GFP::FBFs or GFP alone in mitotic germ cells appeared similar (Figure 4—figure supplement 1C). Together, these data suggest that FBF-1, but not FBF-2, colocalizes with CCR4-NOT in SPCs, in agreement with the dependence of FBF-1 function on CCR4-NOT.
Figure 4 with 1 supplement see all
FBF-1 colocalizes with CCR4-NOT complex in germline SPCs.
(A–B) Confocal images of SPCs co-immunostained for endogenous FBF-1 (A) or GFP-tagged FBF-2 (B, red) and 3xFLAG-tagged CCF-1 (green). DNA staining is in blue (DAPI). Arrows indicate complete overlap of FBF-1 and CCF-1 granules. Asterisks denote FBF-2 granules localizing close but not overlapping with CCF-1 granules. Scale bars in A and B: 5 μm. (C) Pearson’s correlation analysis quantifying the colocalization between FBF and CCF-1 granules in co-stained germline images. Plotted values are means ± SD. N, the number of analyzed germline images (single confocal sections through the middle of germline SPC nuclei including 5–8 germ cells). Asterisks mark statistically significant difference by Student’s t-test, p<0.01. (D) Confocal images of the distal germline SPC zones with PLA foci (grayscale) and DNA staining (blue). Germlines are outlined with dashed lines and vertical dotted lines indicate the beginning of the transition zone. Genotypes are indicated on top of each image group. Scale bar: 10 μm. (A–D) All experiments were performed at 24°C.
Table 2
Proximity ligation assay detects association of FBF-1 with CCR4-NOT complex component CCF-1.
PLA foci density was determined in maximal intensity projections of confocal image stacks encompassing germline SPC zones of the indicated strains. Reported values are mean ± SD derived from three independent biological replicates (3xflag::ccf-1; gfp::fbf-1 and 3xflag::ccf-1; gfp::fbf-2) or a single replicate (3xflag::ccf-1; gfp), all reared at 24°C. Differences in PLA density between 3xflag::ccf-1; gfp::fbf-1 or 3xflag::ccf-1; gfp::fbf-2 and the control 3xflag::ccf-1; gfp were analyzed by one-way ANOVA with Dunnett’s post-test. N, number of germline images analyzed.
| Genotype | PLA density in SPC zone (/um^2) x 10−2 | P value,vs. control | N |
|---|---|---|---|
| 3xflag::ccf-1; gfp::fbf-1 | 5.2 ± 2.4 | <0.0001 | 32 |
| 3xflag::ccf-1; gfp::fbf-2 | 1.1 ± 0.8 | ns | 27 |
| 3xflag::ccf-1; gfp | 0.6 ± 0.2 | n/a | 12 |
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 number of mitotic cells at the restrictive temperature (Pepper et al., 2003a; Figure 5—figure supplement 1A), thus allowing us to focus on the mRNAs in the mitotic cell population.
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).
Figure 5 with 1 supplement see all
Antagonistic effects of FBF-1 and FBF-2 on poly(A) tail of target mRNAs.
(A–C) Representative PAT-PCR amplification of the poly(A) tail of cyb-2.1 (A), htp-1 (B) and control myosin heavy chain unc-54 (C) in fbf-1(lf); glp-1(gf), glp-1(gf), and fbf-2(lf); glp-1(gf) genetic backgrounds at 25°C. The estimated lengths of poly(A) tails based on the PCR fragment sizes are indicated on the right. The areas boxed by dotted lines were quantified by densitometry in ImageJ. (D–F) Densitometric quantification of PAT-PCR amplification products (boxed in A-C). Y-axis, mean relative intensity represents the average intensities of normalized PAT-PCR reactions from three independent biological replicates. X-axis, estimated sizes of poly(A) tails. Values are means + SD. Vertical lines in (D–F) mark the peaks of PAT-PCR intensity profiles for each mRNA in each genetic background, dashed lines mark secondary peaks for htp-1. (A–F) Nematodes for all replicates were grown at 25°C.
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 5—figure 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.
Figure 6 with 1 supplement see all
Three variable regions of FBF-2 prevent its cooperation with CCR4-NOT.
(A) Schematics of FBF-1, FBF-2 and GFP::FBF-2(vrm) mutant transgene (Wang et al., 2016). Red and blue boxes indicate variable regions distinguishing FBF-1 and FBF-2 respectively, grey box indicates the RNA-binding domain, and green box indicates GFP tag. (B) Distal germlines of the indicated genetic backgrounds stained with anti-REC-8 (green) and anti-pH3 (red). Germlines are outlined with the dashed lines, and the vertical dotted line marks the beginning of transition zone. Scale bar: 10 μm. (C) The extent of SPC zone in the indicated genetic backgrounds (on the X-axis). FBF protein(s) present in each genetic background are noted above each data set. Plotted values are individual data points and means ± SD. Differences in SPC zone length between fbf-1(lf) fbf-2(lf); gfp::fbf-2(vrm) and the other strains were evaluated by one-way ANOVA test with Dunnett’s post-test; asterisks mark statistically significant differences (p<0.0001). Data were collected from three independent experiments and 24–28 germlines were scored for each genotype. (D) The extent of SPC zone was measured after crossing the GFP::FBF-2(vrm), GFP::FBF-1(wt) and GFP::FBF-2(wt) transgenes into fbf-1(lf) genetic background. As controls, SPC zone length was also measured in fbf-1(lf) and the wild type. Plotted values are individual data points and means ± SD. Data were collected from two independent experiments and 8–17 germlines were scored for each genotype per replicate. Differences in SPC zone length between fbf-1(lf) and all other strains were evaluated by one-way ANOVA test with Dunnett’s post-test; asterisks mark statistically significant differences (****, p<0.0001; **p<0.01). (E) The extent of SPC zone after knocking down CCR4-NOT subunits in the fbf-1(lf) fbf-2(lf); gfp::fbf-2(vrm) genetic background. RNAi treatments are indicated on the X-axis and average length of SPC zone ± SD on the Y-axis. Differences in SPC zone length between CCR4-NOT knockdowns and control were evaluated by one-way ANOVA (asterisks, p<0.01). Data were collected from three independent experiments. N, the number of independent germlines scored. (F) Confocal images of the distal germline SPC zones with PLA foci (grayscale) and DNA staining (blue). Germlines are outlined with dashed lines and vertical dotted lines indicate the beginning of the transition zone. Genotypes are indicated on top of each image group. Scale bar: 10 μm. (B–D): All experiments were performed at 24°C.
We first tested whether the three VRs are required for FBF-2-specific SPC zone length. To test this hypothesis, the extent of SPC zone was determined after crossing the GFP::FBF-2(vrm) transgene into fbf double mutant background. We found that the SPC zone maintained by GFP::FBF-2(vrm) (Figure 6Biv) is significantly longer than that maintained by GFP::FBF-2(wt) (Figure 6Biii) and the endogenous FBF-2 (p<0.01; Figure 6Bi, C) suggesting that the GFP::FBF-2(vrm) effect on SPC zone length is distinct from that of FBF-2. Western blot analysis indicated that expression of GFP::FBF-2(vrm) is comparable to that of GFP::FBF-2(wt), so their distinct effects on SPC zone length are likely due to functional differences (Figure 6—figure supplement 1A). To test whether GFP::FBF-2(vrm) can rescue either of fbf single mutants, we determined the extent of SPC zone after crossing GFP::FBF-2(vrm) into fbf-1(lf) and fbf-2(lf) genetic backgrounds. As controls, the lengths of SPC zones were also measured after crossing the wild type GFP::FBF-2(wt) and GFP::FBF-1(wt) transgenes into each fbf single mutant. As expected, the SPC zone length of fbf-2(lf); gfp::fbf-2(wt) is significantly shorter than fbf-2(lf) (p<0.01) while the SPC zone of fbf-2(lf); gfp::fbf-1(wt) is similar to fbf-2(lf) (Figure 6—figure supplement 1B), suggesting that GFP::FBF-2(wt), but not GFP::FBF-1(wt), rescues fbf-2(lf). Likewise, GFP::FBF-1(wt), but not GFP::FBF-2(wt), rescues fbf-1(lf) (p<0.01, Figure 6D). Interestingly, we found that the extent of SPC zone of fbf-2(lf); gfp::fbf-2(vrm) is similar to that of fbf-2(lf) (Figure 6—figure supplement 1B), suggesting that GFP::FBF-2(vrm) does not rescue fbf-2(lf). By contrast, the SPC zone of fbf-1(lf); gfp::fbf-2(vrm) is significantly longer than that of fbf-1(lf) (p<0.01, Figure 6D) and there is no significant difference in the SPC zone length between fbf-1(lf); gfp::fbf-2(vrm) and the wild type, suggesting that the GFP::FBF-2(vrm) rescues fbf-1(lf). We conclude that the three VRs outside of FBF-2 RNA-binding domain (VR1, 2, and 4) are important for FBF-2-specific effect on the extent of germline SPC zone and mutation or deletion of these VRs resulted in a mutant protein FBF-2(vrm) that functions similar to FBF-1.
Since FBF-1 function requires the CCR4-NOT complex and FBF-2(vrm) appears similar to FBF-1, we hypothesized that CCR4-NOT is required for FBF-2(vrm) function. To test this hypothesis, we measured SPC zone length after knockdown of CCR4-NOT subunits in fbf-1(lf) fbf-2(lf); gfp::fbf-2(vrm) animals by RNAi. We found that SPC zone of fbf-1(lf) fbf-2(lf); gfp::fbf-2(vrm) after RNAi of CCR4-NOT subunits becomes significantly shorter than the control (p<0.01, Figure 6E), suggesting that GFP::FBF-2(vrm) function requires CCR4-NOT. Furthermore, if GFP::FBF-2(vrm) cooperates with CCR4-NOT, we expect that it might associate with CCF-1 by proximity ligation assay. Indeed, PLA foci density in the mitotic cells of 3xflag::ccf-1; gfp::fbf-2(vrm) was significantly greater than in the control (Figure 6F; Table 3; p<0.0001). We conclude that the VRs outside of FBF-2 RNA-binding domain are required for FBF-2-specific effect on the extent of SPC zone and to prevent FBF-2 from cooperating with CCR4-NOT.
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.
| Genotype | PLA density in SPC zone (/um^2) x 10−1 | p value,vs. control | N |
|---|---|---|---|
| 3xflag::ccf-1; gfp::fbf-2vrm | 2.3 ± 0.7 | <0.0001 | 58 |
| 3xflag::ccf-1; gfp | 1.1 ± 0.5 | n/a | 48 |
The variable region 4 (VR4) of FBF-2 is sufficient to prevent cooperation with CCR4-NOT
To test whether one of the three VRs outside of FBF-2 RNA-binding domain (VR1, 2, and 4) is sufficient to support FBF-2-specific effects on the length of SPC zone, we established a transgenic FBF-1 chimera with VR4 swapped from FBF-2 (GFP::FBF-1(FBF-2vr4); Figure 7A) and crossed it into fbf double mutant. Since VR3 residing in FBF-2 RNA-binding domain was not sufficient for FBF-2-specific function, gfp::fbf-1(fbf-2vr3) (with VR3 swapped from FBF-2; Figure 7A) chimeric transgene was made for comparison. SPC zone length assessment showed that the SPC zone maintained by GFP::FBF-1(FBF-2vr4) (Figure 7Biii) is significantly shorter than that maintained by GFP::FBF-1(wt) (Figure 7Bv) and endogenous FBF-1 (p<0.0001; Figure 7Bii, C). By contrast, the SPC zone maintained by GFP::FBF-1(FBF-2vr3) (Figure 7Biv) is similar to that maintained by the GFP::FBF-1(wt) (Figure 7Biv, C). This finding suggested that GFP::FBF-1(FBF-2vr4) might function similarly to FBF-2. Western blot analysis revealed that the protein expression levels of GFP::FBF-1, GFP::FBF-1(FBF-2vr3), and GFP::FBF-1(FBF-2vr4) were comparable, so the differential effects on SPC zone length are likely due to functional differences (Figure 7—figure supplement 1A). To test whether GFP::FBF-1(FBF-2vr4) rescues FBF-1- or FBF-2-specific function, we measured the extent of SPC zones after crossing GFP::FBF-1(FBF-2vr4) into fbf-1(lf) and fbf-2(lf) genetic backgrounds. For comparison, GFP::FBF-1(FBF-2vr3) was also crossed into each fbf single mutant. We found that the SPC zone of fbf-1(lf); gfp::fbf-1(fbf-2vr4) is similar to that of fbf-1(lf) (Figure 7—figure supplement 1B), suggesting that GFP::FBF-1(FBF-2vr4) does not rescue fbf-1(lf). Interestingly, SPC zone of fbf-2(lf); gfp::fbf-1(fbf-2vr4) is significantly shorter than that of fbf-2(lf) (p<0.01, Figure 7—figure supplement 1C), suggesting that GFP::FBF-1(FBF-2vr4) rescues fbf-2(lf). By contrast, GFP::FBF-1(FBF-2vr3) rescues fbf-1(lf), but not fbf-2(lf) (Figure 7—figure supplement 1B,C). We conclude that the presence of VR4 from FBF-2 in a chimeric GFP::FBF-1(FBF-2vr4) protein is sufficient to impart FBF-2-specific effect on the extent of SPC zone.
Figure 7 with 1 supplement see all
Variable region 4 (VR4) from FBF-2 is sufficient to prevent FBF-1 chimera from cooperation with CCR4-NOT.
(A) Schematics of FBF-1, FBF-2, transgenic GFP::FBF-1(FBF-2vr4) chimera (with VR4 swapped from FBF-2), and transgenic GFP::FBF-1(FBF-2vr3) chimera (with VR3 swapped from FBF-2). Red and blue boxes, variable regions distinguishing FBF-1 and FBF-2 respectively; grey box, RNA-binding domain; green box, GFP tag. (B) Distal germlines dissected from the indicated genetic backgrounds stained with anti-REC-8 (green) and anti-pH3 (red). Germlines are outlined with the dashed lines and the vertical dotted line marks the beginning of the transition zone. Scale bar: 10 μm. (C) The extent of SPC zone in the indicated genetic backgrounds (on the X-axis). FBF protein present in each genetic background is noted above each data set. Plotted values are individual data points and means ± SD. Differences in SPC zone length between fbf-1(lf) fbf-2(lf); gfp::fbf-1(fbf-2vr4) and the other strains were evaluated by one-way ANOVA test with Dunnett’s post-test; asterisks mark statistically significant differences (p<0.0001). Data were collected from three independent experiments and 31–60 germlines were scored for each genotype. (D) SPC zone length after knocking down CCR4-NOT subunits in the fbf-1(lf) fbf-2(lf); gfp::fbf-1(fbf-2vr4) and fbf-1(lf) fbf-2(lf); gfp::fbf-1(fbf-2vr3) genetic backgrounds (as indicated on the X-axis). Plotted values are means ± SD. Asterisks mark the group with significant changes in SPC zone length after CCR4-NOT knockdown vs control RNAi by one-way ANOVA (p<0.01). Data were collected from two independent experiments. N, the number of hermaphrodite germlines scored. (B–D) All experiments were performed at 24°C.
To test whether VR4 is sufficient to inhibit cooperation of GFP::FBF-1(FBF-2vr4) with CCR4-NOT, we measured the length of SPC zone after knockdown of CCR4-NOT subunits in fbf-1(lf) fbf-2(lf); gfp::fbf-1(fbf-2vr4) animals by RNAi. As a control, CCR4-NOT knockdown was also performed on fbf-1(lf) fbf-2(lf); gfp::fbf-1(fbf-2vr3). We found that the SPC zone of fbf-1(lf) fbf-2(lf); gfp::fbf-1(fbf-2vr4) after RNAi of CCR4-NOT subunits is similar to the control (Figure 7D), suggesting that GFP::FBF-1(FBF-2vr4) function in SPCs does not rely on CCR4-NOT. By contrast, the SPC zone of fbf-1(lf) fbf-2(lf); gfp::fbf-1(fbf-2vr3) is significantly shortened after RNAi of CCR4-NOT subunits compared to the control (p<0.01, Figure 7D), indicating that GFP::FBF-1(FBF-2vr3) maintains dependence on CCR4-NOT. We conclude that FBF-2 VR4 in a chimeric GFP::FBF-1(FBF-2vr4) protein is sufficient to support FBF-2-specific effect on the extent of germline SPC zone and to prevent the chimera’s cooperation 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).
Figure 8
A model of antagonistic effects of FBF-1 and FBF-2 on germline SPC dynamics and target mRNAs.
(A) FBFs regulate the rate of stem and progenitor (green) cell division and the rate of entry into meiosis (grey). Both FBFs negatively regulate each other’s expression (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.
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 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 G2-phase 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 (Bernstein et al., 2005; 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 co-bind 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 co-regulation. 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).
Table 4
Variable regions outside of the RNA-binding domain regulate FBF function.
| Transgene | Mutated variable region (VR) sequence | Rescues fbf-1(lf)? | Rescues fbf-2(lf)? | Dependent on CCR4-NOT |
|---|---|---|---|---|
| GFP::FBF-2wt | N/A | No | Yes | No* |
| GFP::FBF-1wt | N/A | Yes | No | Yes* |
| GFP::FBF-2(vrm) | mutated VR1, 2; VR4 deleted | Yes | No | Yes† |
| GFP::FBF-1(FBF-2vr4) | VR4 swapped with FBF-2 | No | Yes | No† |
| GFP::FBF-1(FBF-2vr3) | VR3 swapped with FBF-2 | Yes | No | Yes† |
-
Rescue assays were performed by crossing transgenic GFP::FBFs into loss of function mutant of each fbf, followed by SPC zone length measurement (Figure 6, Figure 6—figure supplement 1, and Figure 7—figure supplement 1). Dependence on CCR4-NOT was defined as a decrease in the length of SPC zone after knocking down CCR4-NOT subunits.
* – analyzed in single fbf loss-of-function mutants, Figure 3B.
-
† – analyzed in the strains containing GFP::FBF transgenes in fbf-1 fbf-2 double-mutant background, Figures 6E and 7D.
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 non-canonical binding sites.
mRNA deadenylation and PUF-mediated repression
Multiple studies suggest that deadenylation contributes to PUF-mediated translational repression (Goldstrohm et al., 2006; Kadyrova et al., 2007; Van Etten et al., 2012; Weidmann et al., 2014). CCR4-NOT deadenylation machinery is conserved in evolution from yeast to humans (Collart et al., 2017; Wahle and Winkler, 2013). Although deadenylation is required for germline stem cell maintenance in flies, nematodes and mice (Berthet et al., 2004; Fu et al., 2015; Joly et al., 2013; Nakamura et al., 2004; Nousch et al., 2019; Shan et al., 2017; Suh et al., 2009), the contribution of deadenylation to PUF translational repression in vivo is still controversial (Weidmann et al., 2014). Previous studies of CCR4-NOT component CCF-1 in C. elegans suggested that paralogous 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.
Materials and methods
Key resources table
| Reagent type (species) or resource | Designation | Source or reference | Identifiers | Additional information |
|---|---|---|---|---|
| Strain, strain background (C. elegans) | JK3022 | Crittenden et al., 2002 | fbf-1(ok91) II | |
| Strain, strain background (C. elegans) | JK3101 | Lamont et al., 2004 | fbf-2(q738) II | |
| Strain, strain background (C. elegans) | GC833 | Pepper et al., 2003a | glp-1(ar202) III | |
| Strain, strain background (C. elegans) | JH3269 | Putnam et al., 2019 | pgl-1::gfp(ax3122) IV | |
| Strain, strain background (C. elegans) | JH2766 | Merritt and Seydoux, 2010 | axIs1922[Ppie-1::GFP::H2B::htp-2 3’UTR] | |
| Strain, strain background (C. elegans) | UMT433 | This paper | mntSi33 [Pgld-1::3xFLAG::CYB-2.1::cyb-2.1 3'UTR] unc-119(ed3) III | |
| Strain, strain background (C. elegans) | UMT394 | This paper | mntSi29 [Pgld-1::3xFLAG::CYB-2.1::cyb-2.1 3'UTR (fbm)] unc-119(ed3) III | |
| Strain, strain background (C. elegans) | UMT360 | This paper | mntSi23 (Pgld-1::3xFLAG::CCF-1::ccf-1 3’UTR) II; unc-119(ed3) III | |
| Strain, strain background (C. elegans) | UMT338 | This paper | mntSi21 [Pgld-1::patcGFP::fbf-1 3'UTR] unc-119(ed3) III | |
| Strain, strain background (C. elegans) | UMT389 | This paper | mntSi28 [Pgld-1::patcGFP::fbf-1::fbf-1 3'UTR] unc-119(ed3) III | |
| Strain, strain background (C. elegans) | UMT378 | This paper | mntSi27 [Pgld-1::patcGFP::fbf-2::fbf-2 3'UTR] unc-119(ed3) III | |
| Strain, strain background (C. elegans) | UMT441 | This paper | mntSi32 [Pgld-1::patcGFP::FBF-2vrm::fbf-2 3'UTR] unc-119(ed3) III | |
| Strain, strain background (C. elegans) | UMT373 | This paper | mntSi26 [Pgld-1::patcGFP::fbf-1(fbf-2vr3)::fbf-1 3'UTR] unc-119(ed3) III | |
| Strain, strain background (C. elegans) | UMT411 | This paper | mntSi31 [Pgld-1::patcGFP::fbf-1(fbf-2vr4)::fbf-1 3'UTR] unc-119(ed3) III | |
| Antibody | Mouse monoclonal anti-FLAG M2 | Sigma-Aldrich Cat# F1804 | RRID:AB_262044 | (1:1,000) |
| Antibody | Rabbit monoclonal anti-GFP | Thermo-Fisher Cat# G10362 | RRID:AB_2536526 | (1:200) |
| Antibody | Mouse monoclonal anti-phospho-Histone H3 pSer10 6G3 | Cell Signaling Technology Cat# 9706 | RRID:AB_331748 | (1:400) |
| Antibody | Rabbit polyclonal anti-REC-8 | Novus Biologicals Cat# 29470002 | RRID:AB_2178279 | (1:500) |
| Antibody | Mouse monoclonal anti-PGL-1 K76 | DSHB Cat# K76 | RRID:AB_531836 | (5.2 μg/ml) |
| Antibody | Rabbit polyclonal anti-FBF-1 | Voronina et al., 2012; PA2388 | (3.5 μg/ml) | |
| Antibody | Mouse monoclonal anti-Tubulin DM1A | Sigma-Aldrich Cat# T6199 | RRID:AB_477583 | (1:300) |
| Antibody | Goat anti-mouse IgG (H+L) 594 | Jackson ImmunoResearch Cat# 115-585-146 | RRID:AB_2338881 | (1:500) |
| Antibody | Goat anti-rabbit IgG (H+L) 594 | Jackson ImmunoResearch Cat# 111-585-144 | RRID:AB_2307325 | (1:500) |
| Antibody | Goat anti-rabbit IgG (H+L) 488 | Jackson ImmunoResearch Cat# 111-545-144 | RRID:AB_2338052 | (1:500) |
| Antibody | Goat anti-mouse IgM 594 | Jackson ImmunoResearch Cat# 115-585-020 | RRID:AB_2338874 | (1:500) |
| Antibody | Goat anti-mouse HRP | Jackson ImmunoResearch Cat# 115-035-003 | RRID:AB_10015289 | (1:5,000) |
| Antibody | Goat anti-rabbit HRP | Jackson ImmunoResearch Cat# 111-035-003 | RRID:AB_2313567 | (1:5,000) |
| Recombinant DNA reagent | Plasmid: control RNAi | Timmons and Fire, 1998 | pL4440 | |
| Recombinant DNA reagent | Plasmid: let-711 RNAi | Source BioScience Ahringer RNAi Collection | ||
| Recombinant DNA reagent | Plasmid: ccr-4 RNAi | Source BioScience Ahringer RNAi Collection | ||
| Recombinant DNA reagent | Plasmid: ccf-1 RNAi | Sönnichsen et al., 2005 | ||
| Recombinant DNA reagent | Plasmid: cyb-2.1 RNAi | This paper | cyb-2.1 genomic CDS in pL4440 | |
| Sequence-based reagent | act-1.qF | Merritt and Seydoux, 2010 | qPCR primers | GGCCCAATCCAAGAGAGGTATC |
| Sequence-based reagent | act-1.qR | Merritt and Seydoux, 2010 | qPCR primers | CAACACGAAGCTCATTGTAGAAGG |
| Sequence-based reagent | unc-54.qF | This paper | qPCR primers | agagagcaggttttggaggat |
| Sequence-based reagent | unc-54.qR | This paper | qPCR primers | ttgagggtgacctcatttcc |
| Sequence-based reagent | him-3.qF | Merritt and Seydoux, 2010 | qPCR primers | AGAGATTTTCGTATCTCTAAATAACGGAATC |
| Sequence-based reagent | him-3.qR | Merritt and Seydoux, 2010 | qPCR primers | GGGTGTATAGTCTTTTGGAGCTTTTTC |
| Sequence-based reagent | htp-1.qF | Merritt and Seydoux, 2010 | qPCR primers | ATTCGGAGGACAGTGACACAA |
| Sequence-based reagent | htp-1.qR | Merritt and Seydoux, 2010 | qPCR primers | GTGCTTTCTCGAGAGACTCAGTTATATC |
| Sequence-based reagent | htp-2.qF | Merritt and Seydoux, 2010 | qPCR primers | ATCGTTCAATTCGGAGGACAC |
| Sequence-based reagent | htp-2.qR | Merritt and Seydoux, 2010 | qPCR primers | GTGTTTTCTCGAGAGAATCGGTTATATT |
| Sequence-based reagent | cyb-1.qF | This paper | qPCR primers | CCAACAACAGACGAACATCG |
| Sequence-based reagent | cyb-1.qR | This paper | qPCR primers | CTGGATTGGATGGCTTGAGT |
| Sequence-based reagent | cyb-2.1.qF | This paper | qPCR primers | AAACCACGAAAAATGCCGT |
| Sequence-based reagent | cyb-2.1.qR | This paper | qPCR primers | TGAAGCTGTCGTCAAGAACA |
| Sequence-based reagent | cyb-2.2.qF | This paper | qPCR primers | CAAGAATCAACATGAAAACGGAT |
| Sequence-based reagent | cyb-2.2.qR | This paper | qPCR primers | TCAGCCATGCAATTGAACTC |
| Sequence-based reagent | cyb-3.qF | This paper | qPCR primers | ACACCATTCAGAAGCTTGCAT |
| Sequence-based reagent | cyb-3.qR | This paper | qPCR primers | AGCGATCTCCGGAAAGGTAG |
| Sequence-based reagent | cyb-2.1.PAT | This paper | PCR primers | tacgttcctgtgttctgctt |
| Sequence-based reagent | htp-1.PAT | This paper | PCR primers | tggcatgttctatatcatcgtgt |
C. elegans culture and strains
Request a detailed protocolAll C. elegans hermaphrodite strains (Supplementary file 1) used in this study were cultured on NNGM plates seeded with OP50 as per standard protocols (Brenner, 1974). All GFP-tagged transgenic animals were cultured at 24°C to avoid GFP silencing. Temperature sensitive allele glp-1(ar202) is a gain-of-function (gf) mutant and is referred to as glp-1(gf) in this study. glp-1(gf) is fertile at 15°C, but sterile at 25°C and produces tumorous germlines. glp-1(gf) was crossed with each single fbf loss-of-function (lf) mutant, fbf-1(ok91) and fbf-2(q738), to generate fbf-1(lf); glp-1(gf) and fbf-2(lf)/mIn1; glp-1(gf). Double mutant strains and glp-1(gf) single mutant were maintained at 15°C. Propagation of fbf-2(lf); glp-1(gf) for large-scale sample collection is detailed below. Synchronized L1 larvae of glp-1(gf) strains were cultured at 25°C until early adulthood. RNA was extracted from tumorous worms and was subsequently used for qPCR and poly(A) tail length analysis.
Propagation of fbf-2(q738); glp-1(ar202gf) nematodes for bulk sample collection
Request a detailed protocolSterility of fbf-2(lf); glp-1(gf) nematodes was partially rescued to 50–66% fertility by a mild glp-1(RNAi). The cultures of fbf-2(lf); glp-1(gf) were propagated on glp-1(RNAi) bacteria at 15°C. Due to heritable RNAi, the rescued fertility was retained up to the third generation after transfer to E. coli food, OP50. To collect large amounts of fbf-2(lf); glp-1(gf) worms with tumorous germlines, the culture of fbf-2; glp-1(ar202); glp-1(RNAi) was expanded on OP50 plates at 15°C before synchronizing by bleach. Then, synchronous L1 larvae were cultured on OP50 at 25°C.
Transgenic animals
Request a detailed protocolAll transgene constructs were generated by Gateway cloning (Thermo Fisher Scientific). GFP::FBF-1 and GFP::FBF-2 constructs were generated with the gld-1 promoter, patcGFP containing introns (Frøkjær-Jensen et al., 2016), fbf-1 or fbf-2 genomic coding and 3’UTR sequences in pCFJ150 (Frøkjaer-Jensen et al., 2008). GFP::FBF-2(vrm) was generated with gld-1 promoter, patcGFP, fbf-2 genomic coding sequences with variable regions 1, 2, and four mutated (P28A, S136A, K137A, Δ139–140, Δ607–632; Wang et al., 2016), and fbf-2 3’UTR in pCFJ150. GFP::FBF-1(FBF-2vr4) and GFP::FBF-1(FBF-2vr3) constructs were generated with gld-1 promoter, patcGFP, fbf-1 genomic coding sequences with swapped variable regions 4 or three from fbf-2, and fbf-1 3’UTR sequences in pCFJ150. 3xFLAG::CCF-1 construct contains gld-1 promoter, ccf-1 genomic coding and 3’ UTR sequences in pCFJ150. 3xFLAG::CYB-2.1wt and 3xFLAG::CYB-2.1fbm constructs contain gld-1 promoter, cyb-2.1 genomic coding and 3’ UTR sequences with either wild type (wt, 5’ UGUxxxAU 3’) or mutated (fbm, 5’ ACAxxxAU 3’) FBF-binding sites in pCFJ150.
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 Cas9-induced 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
Request a detailed protocolC. 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
Request a detailed protocolSynchronous 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-8-positive SPCs. Percent differences in mitotic indices were calculated as for G2-phase length or differentiation rate.
EdU labeling
Request a detailed protocolS-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 MgSO4, 5 μg/mL thymine, 6 μM thymidine and 20 μM 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 K2HPO4 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 × 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 × 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 × 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 × 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
Request a detailed protocolSynchronous 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
Request a detailed protocolTo 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 EdU-positive 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 t50 value of the sigmoidal dose-response 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
Request a detailed protocolGerm 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
Request a detailed protocolThe following RNAi constructs were used: ccr-4, let-711 (Kamath and Ahringer, 2003), ccf-1 (cenix:341 c12; Sönnichsen 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 anti-phospho-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 −80°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 μg total RNA. During PCR amplification, 1 μl 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
Request a detailed protocolThe 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 μg RNA template per each 20 μg 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 ΔΔCt 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 post-tests. 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
Request a detailed protocolFor 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)
Request a detailed protocolPLA 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 × 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
Request a detailed protocolReporter 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
Request a detailed protocolSynchronous 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 μm 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).
Data availability
All data generated or analysed during this study are included in the manuscript and supporting files.
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Decision letter
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Yukiko M YamashitaReviewing Editor; University of Michigan, United States
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Marianne E BronnerSenior Editor; California Institute of Technology, United States
In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.
Acceptance summary:
This study provides a significant advance in our understanding of how stem cell population is regulated in C. elegans germline. It has been known that two PUF proteins FBF-1 and FBF-2 have redundant roles in this process, but there have been unexplained differences in their phenotypes and protein localization. This study highlights functional differences of FBF-1 and FBF-2, explaining how these two PUF proteins together regulate germline stem cell population. Specifically they showed that, although they target the same set of genes, their mechanisms of action is different: FBF-1 (but not FBF-2) relies on CCR4-NOT deadenylase for its function to down regulate target mRNA. Overall, the reviewers felt that this study provides an important advance in our understanding of how C. elegans germline stem cells are maintained, with generalizable knowledge that is likely broadly applicable to other systems.
Decision letter after peer review:
Thank you for submitting your article "PUF family proteins FBF-1 and FBF-2 regulate germline stem and progenitor cell homeostasis in C. elegans" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Marianne Bronner as the Senior Editor. The reviewers have opted to remain anonymous.
The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.
Summary:
This study provides a significant advancement in our understanding of how stem cell population is regulated in C. elegans germline. It has been known that two PUF proteins FBF-1 and FBF-2 have redundant roles in this process, but there have been unexplained differences in their phenotypes and protein localization. This study highlights functional differences of FBF-1 and FBF-2, explaining how these two PUF proteins together regulate germline stem cell population. Specifically they showed that, although they target the same set of genes, their mechanisms of action is different: FBF-1 (but not FBF-2) relies on CCR4-NOT deadenylase for its function to down regulate target mRNA.
Overall, the reviewers felt that this study provides an important advance in our understanding of how C. elegans germline stem cells are maintained, with generalizable knowledge that is likely broadly applicable to other systems. The reviewers provided suggestions to improve the accuracy/clarity of the manuscript.
Essential revisions:
1) The following is summary of required changes/revisions (suggested by multiple reviewers)
– Cell cycle phenotypes for fbf(-)s should be added to/improved
– RT-PCR data needs clarification (the tbb-2 negative control in particular and the statistical analysis), and potentially the addition of complementary CCR4-NOT data
– CYB-2.1 with mutated FBEs should include a wild-type control and more thorough characterization of the effect on cell cycle progression
– Deadenylation of CYB-2.1 needs additional controls (wild-type = glp-1(gf) alone) and to look at additional targets that might show a similar trend.
– While the specific suggestions differ, a little more from the domain swapping experiments would add to the manuscript – showing changes in co-localization, addressing the implications of their different rescue experiments, and/or experiments addressing readouts other than PZ/Rec8+ zone size.
– All think Figure 8 (the model) needs clarification/improvement to add to the manuscript
2) In addition, individual reviewers provided suggestions/comments, which can be addressed by textual changes (individual comments can be found at the bottom of this letter). Although these may not require additional experiments, we would like the authors to address these points by editing to improve accuracy/clarity of the manuscript.
Reviewer #1:
Wang et al. provide significant contributions to the interesting story of differing roles for two very similar PUF proteins FBF-1 and FBF-2 in the C. elegans germ line with implications for PUF proteins in general. First, based on cell cycle measurements, they define different and interesting roles for the FBFs in regulating rates of mitotic cell cycle progression and meiotic entry ("proliferation" and "differentiation"), and they identify a key cyclin. Second, they distinguish the role of FBF-1 as dependent on CCR4-NOT deadenylase, likely leading to degradation of target mRNAs. Third, they define sequences in FBF-1 that mediate CCR4-NOT dependence.
Comments and Suggestions:
1) Figure 5. A major conclusion is that FBF-1 is promoting deadenylation of its target mRNAs via CCR4-NOT. Demonstration of PolyA length differences with additional FBF-1 targets would bolster this conclusion.
2) mRNA quantification after CCR4-NOT RNAi as in and compared with Figure 1G would also further bolster the degradation conclusion.
3) Does the FBF-2 VR mutant now associate with CCF-1 in the proximal ligation assay? If so, this would strengthen the conclusion that the function has been shifted to FBF-1-CCR4-NOT.
4) Figure 8. Are you proposing a competition model? Can you clarify how your molecular model will play out in the control of the spatial extent of the PZ?
5) "Proliferation" and its control should be clarified where possible as control of "rate of mitotic cell cycle progression". In the past, there has been confusion in the field regarding "proliferation" as the alternative cell fate to "differentiation". For "meiotic entry", clarify where possible spatial position versus rate. Again, to help with the past confusion in the field, several places in the manuscript could clarify by substituting "differentiation" with position or rate of "meiotic entry".
6) Several comments regarding the term "SPC zone size", throughout the manuscript:
First, best to call the pool of REC-8-positive cells "progenitor cells", and the zone as the "progenitor zone", abbreviating as "PZ", rather than defining a new term and acronym.
Second, the "zone size" should be (1) standardized to numbers of cells in the key genotypes to account for possible differences in gonad width and (2) defined more clearly as a distance measurement since the term "size" has been used in the literature to mean cell numbers, and (3) justified better as a readout.
Third, since this distance measurement is a key phenotypic readout throughout the manuscript, define better how the position of the dotted lines indicating the edge of the PZ was determined: the distance between the end of the REC-8 positive cells and the dotted line is not uniform in the figures (compare, for example Figure 1C and 6B). It is especially unclear for Figure 7B.
7) Introduction paragraph two. The position of cell divisions is not uniform (e.g., Maciejowski et al.). Please restate.
8) Subsection “FBF-1 and FBF-2 differentially regulate mRNA abundance of target genes controlling proliferation and differentiation”. Early adult glp-1(ar202) shifted to 25º as L1 does not contain "only mitotic cells" (Pepper et al., 2003). Please restate.
9) I found the naming of the vr mutants and swaps made this section hard to follow. Perhaps you could name them based on the donor and target proteins?
10) Discussion paragraph two. This point is subtle and a bit confusing. Perhaps clarify that you mean the proportions of cycling-competent versus non-cycling-competent pre-meiotic S that normally contribute to the PZ?
11) In the same paragraph add "specific", "no specific molecular markers…"
Reviewer #2:
Wang et al. have proposed that FBF proteins function differentially in regulating the same set of mRNA targets. Using a set of FBF targets they demonstrated that mRNA is downregulated by FBF-1 but not FBF-2. They provided evidence that FBF-1, but not FBF-2, functions together with CCR-4/NOT to de-adenylated cyb-2.1 and possibly other FBF targets. They argued that 3 variable regions (VR), present in FBF-2, are critical for their distinct molecular identity, removing and swapping these regions would make them each behave like other. This work furthers the mechanistic understanding on how puf proteins function in regulating their targets, the logic is straightforward and the manuscripts clearly deliver the above findings to readers. However, the data provided by the Wang et al. suffers from improper statistical analysis, imprecise interpretation of result and needs additional data to support the claims in the manuscript.
1) FBF-1 and FBF-2 differentially modulate proliferation and meiotic entry of C. elegans germline SPCs.
The authors have found that fbf mutants have differential mitotic index and concluded that they have different rate of proliferation. For the fbf-2 mutant, a longer cell cycle is supported by slightly longer G2 phase. However, for the fbf-1 mutant G2 length is the same as wild type, which is inconsitent with the higher mitotic index (Lara-Gonzalez et al., 2019). Higher M-phase index could instead be due to cells spending more time in M-phase (which is possible due to overexpression of cell cycle regulators such as cyclin B proteins, which is downregulated to trigger M-phase exit). Additional data will need to be presented before making the conclusion that the fbf-1 mutant has a shorter cell cycle. Determining the S-phase index could provide support for a shorter cell cycle. The S-phase index data could also bolster the point that fbf-2 has a longer cell cycle, as the G2 lengthening is very modest and thus unlikely to explain the decreased meiotic output.
Importantly, the result that meiotic output of fbf-1 and fbf-2 mutants is higher and lower, respectively, than wild type is sufficient to explain the differences in the SPC zone length and FBF proteins role in meiotic entry. However, the data suffered from low sample size.
2) FBF-1 and FBF-2 differentially regulate mRNA abundance of target genes controlling proliferation and differentiation.
The authors hypothesized that "FBF-1 and FBF-2 might be acting differentially on the same set of mRNAs". They chose a set of mRNA targets of FBF (3 meiotic genes and 4 cell cycle regulators). Based on qPCR in fbf mutants and wild type (all in glp-1(gf) background, Figure 1G), authors argue that FBF-1 destabilize these target mRNAs, while FBF-2 promotes their accumulation. This data lays the foundation for their subsequent analysis. However, the data need improvement in several ways:
i) The authors have chosen linear trend analysis to show that, for all the mRNA tested, levels are higher in fbf-1(-) > wild type > fbf-2. While trend looks to be true, but given that control, tbb-2 mRNA levels are reduced by almost half in fbf-2 mutant compared to wild type, with a possible exception cyb-2.2, none of the mRNAs tested can be different in fbf-2 mutant compared to wild type. Based on the graph one can only argue that FBF-1 destabilize mRNA levels but FBF-2 does not have any appreciable effect.
ii) The statistical labels in Figure 1G is inconsistent with the statement made in the result. The authors claimed that other than cyb-1, all the other targets mRNA expression are increased in fbf-1(-), but reduced in fbf-2(-). While in Figure 1G, the statistical significance label for htp-2 and cyb-1 are EXACTLY the same. In addition, the authors are encouraged to explain why one-way ANOVA was chosen for their statistical analysis in the Materials and methods section. A pairwise comparison between wild type and each fbf single mutant should be performed for all the mRNAs tested; if there is a significant difference then one can make a conclusion.
iii) The authors indicate that they are studying these targets' regulation in germline tumor, however the RNA is from whole worms. The authors are encouraged to explain their approach.
3) Repression of cyclin B by FBF limits accumulation of germline SPCs.
The authors have mutated all three FBEs in cyb-2.1 and have successfully shown that CYB-2.1 protein levels have become higher in immunoblot. They also show that this mutation causes an increase of SPC size in fbf-2 mutant and this can be reverted by depleting cyb-2.1. Based on this the authors conclude that "levels of B-type cyclins limit SPC proliferation rate in fbf-2(lf) and disruption of FBF-1-mediated repression of a single cyclin B gene is sufficient to affect the size of germline SPC zone".
There are multiple issues with this result:
i) C. elegans has ~12 puf genes (Prasad et al., 2016) and most of them have not been fully characterized; thus it is possible that mutated FBEs might be recognized by one or more of these puf proteins rather than FBF. Immunostaining and quantification of CYB-2.1 in extruded germlines of the fbf-1 mutant and the fbf-2 mutant would be required to claim that there is reciprocal regulation by the FBF proteins, rather than other puf proteins.
ii) Mutating FBEs in cyb-2.1, results in its higher expression. Rationale for checking the effect of its overexpression in fbf-2(-) background, instead of wild type, is not clear. Since there are no FBEs in cyb-2.1fbm, its effects are expected to be fbf-agnostic.
iii) Subsection “Repression of cyclin B by FBF limits accumulation of germline SPCs” paragraph three authors have concluded that increase in SPC zone size in fbf-2; cyb-2.1fbm vs fbf-2 is due to increased proliferation and also the meiotic entry rate is unchanged between two mutants. They have not provided any evidence for either of these two possibilities, i.e. proliferation rate has increased in fbf-2; cyb-2.1fbm and meiotic entry rate is same in fbf-2; cyb-2.1fbm as compared to fbf-2.
4) FBF-1 promotes deadenylation of its target mRNA.
The authors moved back to cyb-2.1 to show that poly A tails are affected in fbf mutants without giving any rationale. The assay should be performed on a few more genes if any generalized statement statement is made, otherwise it must be restricted to cyb-2.1. Also wild type (glp-1(gf)) is missing in the assay, which makes it hard to interpret the data. Is FBF-1 alone or both FBFs responsible for deadenylation of cyb-2.1 mRNA or if FBF-1 and FBF-2 have opposing effect on cyb-2.1 mRNA?
5) While the authors present some solid lines of evidence that FBF-1 and FBF-2 are different (meiotic entry rate, FBF-1 acts with CCR-4/NOT to de-adenylated cyb-2.1 and there are specific sequences in FBF-2 that block this interaction), they are clearly redundant with each other for the stem cell fate and repression of htp-2. Thus, Figure 8 would be more useful to the community if their redundant function was also included.
6) FBF-1 and FBF-2 do not explain the spatial control of expression in the SPC zone and thus balance between proliferation and differentation, which is through glp-1 targets lst-1 and sygl-1. It is surprising that there is no mention of lst-1 and sygl-1-1 in the Discussion, particularly since they are co-factors of FBF-1 and FBF-2 (Haupt et al., 2019, Shin et al., 2017).
Reviewer #3:
The C. elegans germline is a well-established model for investigating how signaling from a niche regulates the balance between stem cell self-renewal and differentiation. In the germline, the stem cell population is maintained as a pool, i.e. rather than undergoing invariant asymmetric divisions that always produce one stem cell and one differentiating daughter cell, the proliferative potential of any stem cell is determined by its distance from the niche and/or the level of niche signaling that it receives. The germline niche produces a Notch ligand which signals via the Notch receptor GLP-1 to promote the stem or mitotic cell fate. Once the level of Notch signaling drops below a threshold, cells enter meiosis. Downstream of Notch signaling, the FBF proteins, FBF-1 and FBF-2, act to maintain the stem cell pool and in fbf-1, fbf-2 double mutants, at least at the adult stage, the stem cell population is lost and the germline resembles one in which Notch signaling has been removed entirely (Crittenden et al., 2002). This phenotype, in combination with the observation that FBF-1/2 share 89% identity and the majority of mRNA targets, has led to a simplified model in which they serve largely redundant roles. However, a fully redundant function is not supported by observations that single loss of function fbf-1 or fbf-2 germlines have distinct phenotypes (Lamont et al., 2004) and that FBF-1 and FBF-2 proteins have different localization patterns (Voronina et al., 2012).
Here the authors attempt to dissect what underlies this difference. Their work addresses two fundamental questions of broad impact/interest (1) how does niche signaling maintain a balance of proliferation and differentiation in a stem cell pool and (2) how can different modes of action be specified in largely similar proteins. As the first is broadly relevant to the stem cell field and the second is addressed within the context of two highly conserved proteins that play a role in stem cell maintenance in many organisms, this work is likely to be interesting to a wide audience. It should also be noted that because of the complexity of the system (the large number of FBF-1/2 targets and the number of other factors that also impinge upon stem cell dynamics), these questions are quite hard to address, and I commend the authors for trying to do so.
My critique of this manuscript can be broken into two categories: (1) it would benefit from some changes in writing to increase clarity and accessibility for non-experts and (2) there are a few questions that I had regarding experimental issues, some related to interpretation, which could be addressed in the text (points D, and F-H below) and some related to additional controls/experiments that would substantiate the authors' main conclusions (points A-C, E, and F). The suggested experiments are not substantially outside of those that the authors have already conducted and I think that most of them (enough to satisfy the majority of my major comments) could be performed within a reasonable time frame.
Major comments:
1) Writing/interpretation:
As someone who works in the germline, but not on this pathway in particular, I found that this paper was often hard to digest. It would be very helpful if the authors could make sure that their rationale for each experiment and its interpretation is accessible to a reader not intimately familiar with the field. For example, the difference in proliferative zone (PZ) size in fbf-1(lf) and fbf-2(lf), I think pointing out here how a change in PZ size shows a shift in the balance between proliferation and differentiation, as if both were slowed down or sped up in complete coordination, the size of PZ would likely be constant. In addition, explaining why experiments were designed in a certain way and why the (perhaps) easier experiment is not feasible would be very helpful.
I also believe that the authors need to revise their language in several instances to avoid conflating two different questions/fields. For example, I found the use of "differentiation demands" confusing, because, for me, "differentiation demands" implies reproductive demand, i.e. the need for mature gametes. In both the C. elegans germline (e.g. Narbonne et al., 2015) and in other stem cell populations (e.g. Hsu, Li and Fuchs, 2014), signaling from differentiated/differentiating progeny to their stem cell parents plays a significant role in regulating stem cell proliferation, such that proliferative output matches the demand for differentiated progeny. I do not think that the authors are addressing this type of "feedback" signaling – i.e. they do not show that the rate of meiotic entry regulates the rate of proliferation, but rather that these two things are coordinated. I think that it is important to clarify their questiion to avoid confusion and/or erroneous expectations from people, such as myself, who are more used to thinking about feedback mechanisms. There are several instances of this throughout, including: "Complementary activities of FBF-1 and FBF-2 combine to fine tune SPC proliferation and differentiation to respond to proliferative demands of the tissue."
2) Experimental issues:
A) Figure 1. The cell cycle phenotype of fbf-1(lf) is puzzling. Since there are no quiescent cells in the germline, an increase in mitotic index (MI) ought to reflect either a faster passage through the mitotic cell cycle or a longer duration of mitosis. The authors report no change in the length of G2, which leads me to wonder whether S phase is accelerated or mitosis is delayed (I believe there are assays to assess both (S phase, Fox and Schedl, 2011) and M phase (Gerhold et al., 2015)). This result is also odd in light of their observation that the transcript levels of most of the Cyclin Bs (except for CYB-1), are highly elevated – i.e. why is the duration of G2 unaffected despite higher levels? The authors explanation in the discussion that a higher MI could be due to a smaller proliferative population, due to accelerated meiotic entry, would predict (I think) that the stem cell pool should eventually be lost – i.e. in older fbf-1(lf) adults, do they see a progressive loss of proliferative cells? As this interpretation (i.e. FBF-2, rather than FBF-1, being primarily responsible for cell cycle regulation) is key to their model figure at the end, it bears a closer look.
B) Also Figure 1. I am unsure as to how to interpret the RT-qPCR data. The authors state that their negative control (tbb-2) is unaffected, yet in fbf-2(lf) transcript levels appear to be at least a strongly depressed as those of the genes that they argue are maintained by FBF-2. It is possible that I am missing something here, but it would be good if the authors could address/explain this, as it is visually striking/distracting in their data/graph and if it is accurate, i.e. that in fbf-2(lf) transcript levels are generally lower, it is problematic for the interpretation that FBF-1 and FBF-2 have reciprocal effects on shared target transcripts. It is also not entirely clear to me what the linear trend test adds to the data interpretation, some clarification/discussion would be helpful.
C) In Figure 2, the authors address the hypothesis that slow G2 and low MI in fbf-2(lf) is due to FBF-1-mediated translational repression of cyb-2.1. The authors state that protein levels for CYB-2.1 with mutated FBF binding elements (FBEs) are elevated, but they cannot detect protein levels for the WT version, which could imply that it is not expressed at all. It would be helpful if the authors could should that both transgenes are expressed by looking at transcript levels. I also wonder if this result predicts that in wildtype germlines, expression of cyb-2.1 with mutated FBEs, but not wild-type cyb-2.1 would affect cell cycle progression? Finally, this hypothesis also seems to predict that a partial RNAi of FBF-1 in a fbf-2(lf) background, should rescue, at least to some extent, the fbf-2(lf) phenotype. If this experiment is technically feasible, it would greatly strength the authors argument.
D) In Figure 3, the authors use a translational reporter for the htp-2 3'UTR, which their RT-qPCR results suggest should be derepressed in fbf-1(lf), but they do not seem to see this and I am unsure as to why not? In addition, their results strongly suggest that the change in PZ size in fbf-2(lf) requires CCR4-NOT complex members, which supports the hypothesis that FBF-1 activity with CCR4-NOT is leading to transcript loss. However, if FBF-1 acts with CCR4-NOT to regulate transcript levels, shouldn't RNAi of CCR4-NOT complex members phenocopy fbf-1(lf)? It would be helpful if the authors could address these questions. This may be another instance of where the data/results are not fully accessible to a non-expert.
E) In Figure 5, the authors quantify the polyA tail length of cyb-2.1 and show that it is shorter in fbf-2(lf), suggesting that it is being deadenylated, presumably by FBF-1/CCR4-NOT. It would strengthen the authors conclusion substantially, if they could show that the change in polyA tail length was rescued by depletion of CCR4-NOT complex members. It is also unclear to me why the authors compared fbf-1(lf) to fbf-2(lf) rather than to wildtype? I also wonder whether tbb-2 is the best control as the detection levels seem very low which could make slight changes in peak detection challenging.
F) Figure 6: I think that this a key place where the authors interpretation could be clarified somewhat. I believe that in Figure 6, the fact that FBF-2 without variable regions (VRs) 1, 2 and 4 cannot fully rescue the fbf-2(lf) phenotype in either a fbf-2(lf) or a fbf-1(lf) fbf-2(lf) background suggests that either this version of FBF-2 is only partially functional or it is acting like FBF-1. The authors then provide support for the latter interpretation by showing that the activity of FBF-2(vrm) requires CCR4-NOT and can also partially rescue fbf-1(lf). This last piece of data is in Figure 6—figure supplement 1, but, in my opinion, should be moved into the main figure panel. This also suggests that FBF-1 may be able to act with CCR4-NOT without any of its own variable regions, which (I think?) is somewhat surprising. The authors could test this by using a similar approach.
G) Figure 7: For the domain swapping experiments here, I believe the main conclusions are that VR3 (in the RNA-binding domain) is not required for FBF-1 activity, and that VR4 from FBF-2 is sufficient to prevent FBF-1 from acting with CCR4-NOT to regulate polyadenylation, as FBF-1 with FBF-2's VR4 cannot rescue fbf-1(lf) in a fbf-1(lf) or a fbf-1(lf),fbf-2(lf) background, while FBF-1 with FBF-2's R3 can. However, what I do not understand is why FBF-1 with FBF-2's VR4 rescues fbf-2(lf) (Figure 7—figure supplement 1)? Does this mean that it is acting as a dominant negative, since presumably wild-type FBF-1 is still present in this background?
H) Figure 8: The model figure as currently drawn doesn't really help me understand the authors' mains conclusions/findings. It would be great if this could be re-worked to reflect the effects of FBF-1 and FBF-2 on the stem cell population (i.e. perhaps in relation to PZ size which is their phenotypic readout for most experiments) and how these activities lead to a balance between stem cell maintenance and differentiation.
https://doi.org/10.7554/eLife.52788.sa1Author response
Essential revisions:
1) The following is summary of required changes/revisions (suggested by multiple reviewers).
– Cell cycle phenotypes for fbf(-)s should be added to/improved
As suggested by the reviewers, we now report new experiments to provide additional analysis of cell cycle in the three genotypes, including the S-phase index (Figure 1—figure supplement 1D). We additionally have determined cell doubling rate during larval development (as in Lara-Gonzalez et al., 2019) and found that the cell doubling rates in fbf-1 mutant and wild type are similar, while the doubling time of fbf-2 mutant is significantly longer (Figure 1E and Figure 1— figure supplement 1E, F).
– RT-PCR data needs clarification (the tbb-2 negative control in particular and the statistical analysis), and potentially the addition of complementary CCR4-NOT data
In response to the reviewers’ suggestions we have made the following changes:
1) We have replaced the negative control with unc-54 myosin, and report that its abundance is not affected similarly to the FBF target abundance.
2) Upon repeating the qPCR experiments, we found that the increase of FBF target transcript levels in fbf-1(lf) was not reproducible. We have increased the number of biological replicates analyzed for each fbf mutant to gain confidence in the conclusions. The decrease in abundance of FBF targets in the fbf-2(lf) mutant background has been validated.
3) For statistical analysis, we have replaced linear trend analysis by a comparison between fbf-2 mutant and the wild type background, since the target levels in fbf-1(lf) did not appear to change appreciably. This analysis revealed significant downregulation of multiple target mRNAs in fbf-2(lf) mutant (Figure 5—figure supplement 1B).
While we agree that it would be informative to test whether the decrease in FBF targets in fbf-2(lf) background was dependent on CCR4-NOT, we were not able to perform the analysis due to technical difficulties of growing large-scale cultures of fbf-2(lf); glp-1(gf) animals while achieving effective RNAi knockdown of CCR4-NOT.
– CYB-2.1 with mutated FBEs should include a wild-type control and more thorough characterization of the effect on cell cycle progression
We have included the following new experiments addressing these suggestions:
1) We have generated a new wild type cyb-2.1 control transgene, and now document its expression at the mRNA and protein levels (Figure 2B, C).
2) We now include estimation of larval SPC proliferation rate comparing fbf-2(lf) with the strains expressing wild type or mutant 3xflag::cyb-2.1 transgenes. These new experiments suggest that cell cycle rate is accelerated in fbf-2(lf); 3xflag::cyb-2.1fbm compared to fbf-2(lf), but not significantly changed in fbf-2(lf); 3xflag::cyb-2.1wt (Figure 2D).
3) We have documented meiotic entry in the fbf-2(lf) compared to the strains expressing wild type or mutant 3xflag::cyb-2.1 transgenes. These new experiments suggest that meiotic entry rate is not affected by the expression of 3xflag::cyb-2.1 transgenes (Figure 2E).
– Deadenylation of CYB-2.1 needs additional controls (wild-type = glp-1(gf) alone) and to look at additional targets that might show a similar trend.
We have addressed these suggestions by the following modifications of the experiment reflected in the updated Figure 5:
1) We have analyzed an additional FBF target, htp-1, and now find that its poly(A) tail similarly shortens in fbf-2(lf) and lengthens in fbf-1(lf) (Figure 5B, E)
2) We have included glp-1(gf) control to compare with fbf mutants, and find that the poly(A) tails of FBF targets in glp-1(gf) appear to be of intermediate lengths, in between the distributions in fbf-1(lf) and fbf-2(lf).
3) As suggested by reviewer 3, we have replaced tbb-2 by a different control, myosin heavy chain unc-54 (Figure 5C, F)
– While the specific suggestions differ, a little more from the domain swapping experiments would add to the manuscript – showing changes in co-localization, addressing the implications of their different rescue experiments, and/or experiments addressing readouts other than PZ/Rec8+ zone size.
We have addressed these reviewer suggestions as follows:
1) As suggested by reviewer 1, we have tested whether FBF-2(VRM) variable region mutant associates with CCF-1. We found that proximity ligation assay detects significant enrichment of signal over background suggesting that FBF-2vrm RNP now incorporates CCR4-NOT deadenylase. These data are reported in Figure 6F and Table 3.
2) As suggested by reviewer 3, we moved fbf-1 rescue experiment to the main Figure 6 (panel D).
3) As suggested by reviewer 3, we elaborated the discussion of rescue experiments (in the Discussion section).
– All think Figure 8 (the model) needs clarification/improvement to add to the manuscript.
In response to reviewers’ suggestions, we have modified the model as follows: We propose that FBFs bind to the same mRNAs, and likely to the same sites within these transcripts (as documented by multiple CLIP studies). We note in the Discussion that it is possible that FBFs either compete for the same binding sites or simultaneously bind to the transcripts that contain multiple FBEs. Either mechanism would explain how change-of-function mutants such as FBF-2vrm and FBF-1(FBF-2vr4) are able to rescue the loss-of-function of the non-cognate fbf.
The model in Figure 8 now emphasizes that both FBFs repress the bound target mRNAs and both FBFs can maintain SPCs on their own (thus highlighting their common, redundant function). Furthermore, we propose that FBF effects on the extent of the mitotic zone result from their regulation of stem cell dynamics (how fast the stem cells are able to divide and the rate of meiotic entry).
Reviewer #1:
Wang et al. provide significant contributions to the interesting story of differing roles for two very similar PUF proteins FBF-1 and FBF-2 in the C. elegans germ line with implications for PUF proteins in general. First, based on cell cycle measurements, they define different and interesting roles for the FBFs in regulating rates of mitotic cell cycle progression and meiotic entry ("proliferation" and "differentiation"), and they identify a key cyclin. Second, they distinguish the role of FBF-1 as dependent on CCR4-NOT deadenylase, likely leading to degradation of target mRNAs. Third, they define sequences in FBF-1 that mediate CCR4-NOT dependence.
Comments and Suggestions:
1) Figure 5. A major conclusion is that FBF-1 is promoting deadenylation of its target mRNAs via CCR4-NOT. Demonstration of PolyA length differences with additional FBF-1 targets would bolster this conclusion.
As suggested, we now report poly(A) length differences for an additional FBF target, htp-1 (Figure 5B, E).
2) mRNA quantification after CCR4-NOT RNAi as in and compared with Figure 1G would also further bolster the degradation conclusion.
Regretfully, we did not have time to optimize worm culture/RNAi to run this experiment.
3) Does the FBF-2 VR mutant now associate with CCF-1 in the proximal ligation assay? If so, this would strengthen the conclusion that the function has been shifted to FBF-1-CCR4-NOT.
As suggested, we have tested association between FBF-2(VRM) and CCF-1 and found that proximity ligation assay detects significant increase of signal over background, supporting the conclusion that FBF-2(VRM) is associating with CCR4-NOT deadenylase. These data are reported in Figure 6F and Table 3.
4) Figure 8. Are you proposing a competition model? Can you clarify how your molecular model will play out in the control of the spatial extent of the PZ?
As noted in the Essential Revisions, we speculate that FBFs compete for mRNAs targets. FBF binding to the same targets is documented by multiple CLIP studies. We note in the Discussion that it is possible that FBFs either compete for the same binding sites or simultaneously bind to the transcripts that contain multiple FBEs. Simultaneous binding is less likely given distinct localization of FBFs in vivo. Either mechanism would explain how change-of-function mutants such as FBF-2vrm and FBF-1(FBF-2vr4) are able to rescue the loss-of-function of the non-cognate fbf.
Our model further suggests that FBF effects on the extent of the mitotic zone result from their regulation of stem cell dynamics (how fast the stem cells are able to divide and the rate of meiotic entry).
5) "Proliferation" and its control should be clarified where possible as control of "rate of mitotic cell cycle progression". In the past, there has been confusion in the field regarding "proliferation" as the alternative cell fate to "differentiation". For "meiotic entry", clarify where possible spatial position versus rate. Again, to help with the past confusion in the field, several places in the manuscript could clarify by substituting "differentiation" with position or rate of "meiotic entry".
As suggested, we have replaced “proliferation” with “cell division rate” or “cell cycle progression rate” where appropriate. We have also clarified where “differentiation” was used to mean the rate of meiotic entry.
6) Several comments regarding the term "SPC zone size", throughout the manuscript:
First, best to call the pool of REC-8-positive cells "progenitor cells", and the zone as the "progenitor zone", abbreviating as "PZ", rather than defining a new term and acronym.
In our manuscript, we decided to follow the lead of multiple preceding publications (for example, Korta et al., Development 2012; Gerhold et al., Curr Biol 2015; Roy et al., Dev Bio, 2016) in referring to the proliferative cells as “stem and progenitor cells” to reflect that this population likely includes both self-renewing stem cells and progenitor cells primed for differentiation.
Second, the "zone size" should be (1) standardized to numbers of cells in the key genotypes to account for possible differences in gonad width and (2) defined more clearly as a distance measurement since the term "size" has been used in the literature to mean cell numbers, and (3) justified better as a readout.
1) and 3) we have determined that progenitor zone size robustly correlates with the numbers of cells in the wild type and fbf single mutant germlines and now report this correlation in the Materials and methods section and include the data in the supplement (R2=0.7794; Figure 1—figure supplement 2).
2) We have replaced SPC zone size with either “length” or “extent” to clarify our metric.
Third, since this distance measurement is a key phenotypic readout throughout the manuscript, define better how the position of the dotted lines indicating the edge of the PZ was determined: the distance between the end of the REC-8 positive cells and the dotted line is not uniform in the figures (compare, for example Figure 1C and 6B). It is especially unclear for Figure 7B.
Thank you for bringing this to our attention. It appears that the differences between panels in Figures 1C, 6B and 7B were due to inconsistent cropping of the images and post-acquisition adjustment of anti-REC-8 staining intensity. These have been now corrected. We define boundary of the mitotic zone as the proximal-most full row of nuclei strongly positive for REC-8; this definition is now included in Materials and methods section.
7) Introduction paragraph two. The position of cell divisions is not uniform (e.g., Maciejowski et al.). Please restate.
Restated to read “cell divisions are distributed throughout the progenitor zone”.
8) Subsection “FBF-1 and FBF-2 differentially regulate mRNA abundance of target genes controlling proliferation and differentiation”. Early adult glp-1(ar202) shifted to 25º as L1 does not contain "only mitotic cells" (Pepper et al., 2003). Please restate.
We restated to read “containing a large number of mitotic cells”. A highly penetrant phenotype of the early adult glp-1(ar202gf) is proximal proliferation rather than a fully-tumorous germline (Pepper et al., 2003). However, depending on the rearing conditions, late-onset fully-tumorous phenotype is often observed (Pepper et al., 2003). In our experiments, we often observe fully tumorous phenotype with only a minor contribution of gametogenesis at the restrictive temperature. We now document this approach in more detail in Materials and methods, and include the REC-8/pH3 staining images in Figure 5—figure supplement 1A.
9) I found the naming of the vr mutants and swaps made this section hard to follow. Perhaps you could name them based on the donor and target proteins?
We have changed the names of the VR mutants as suggested to indicate the target protein followed by the donor protein and variable region number in parentheses; for example, FBF-1(FBF-2vr4).
10) Discussion paragraph two. This point is subtle and a bit confusing. Perhaps clarify that you mean the proportions of cycling-competent versus non-cycling-competent pre-meiotic S that normally contribute to the PZ?
We have re-written this statement as suggested.
11) In the same paragraph add "specific", "no specific molecular markers…"
Changed as suggested.
Reviewer #2:
[…] However, the data provided by the Wang et al. suffers from improper statistical analysis, imprecise interpretation of result and needs additional data to support the claims in the manuscript.
1) FBF-1 and FBF-2 differentially modulate proliferation and meiotic entry of C. elegans germline SPCs.
The authors have found that fbf mutants have differential mitotic index and concluded that they have different rate of proliferation. For the fbf-2 mutant, a longer cell cycle is supported by slightly longer G2 phase. However, for the fbf-1 mutant G2 length is the same as wild type, which is inconsitent with the higher mitotic index (Lara-Gonzalez et al., 2019). Higher M-phase index could instead be due to cells spending more time in M-phase (which is possible due to overexpression of cell cycle regulators such as cyclin B proteins, which is downregulated to trigger M-phase exit). Additional data will need to be presented before making the conclusion that the fbf-1 mutant has a shorter cell cycle. Determining the S-phase index could provide support for a shorter cell cycle. The S-phase index data could also bolster the point that fbf-2 has a longer cell cycle, as the G2 lengthening is very modest and thus unlikely to explain the decreased meiotic output.
Importantly, the result that meiotic output of fbf-1 and fbf-2 mutants is higher and lower, respectively, than wild type is sufficient to explain the differences in the SPC zone length and FBF proteins role in meiotic entry. However, the data suffered from low sample size.
We apologize for being unclear: our conclusion from cell cycle analysis in the wild type and fbf mutants was that wild type and fbf-1 have similar cell cycle lengths, while fbf-2 mutant has a longer cell cycle length. We now emphasize this conclusion in the Results section immediately after reporting the mitotic index increase in fbf-1 by telling that the hypothesis of faster cell cycle in fbf-1 mutant was later rejected to avoid confusion.
We added an alternative interpretation that fbf-1 cells spend longer time in M-phase in Discussion.
We now report new experiments to provide additional analysis of cell cycle in the three genotypes, including determination of the S-phase index in the adult and documenting accumulation of germline stem cells during larval development (Figure 1E, Figure 1—figure supplement 1D, E, F; see also our response to Essential Revisions).
Finally, we respectfully disagree with regard to the sample size used for meiotic entry rate determination. The rate of meiotic entry was estimated in five independent experiments, each using 5-7 germlines per strain per time point. This comes up to 89-94 total germlines scored per strain, which is larger than the sample size utilized in previous reputable publications (eg., Kocsisova, Kornfeld and Schedl, 2019 Development score 49-57 total germlines per strain). We now report the total number of germlines analyzed per strain in the legends of Figure 1 and Figure 1—figure supplement 1 to help communicate this point.
2) FBF-1 and FBF-2 differentially regulate mRNA abundance of target genes controlling proliferation and differentiation.
The authors hypothesized that "FBF-1 and FBF-2 might be acting differentially on the same set of mRNAs". They chose a set of mRNA targets of FBF (3 meiotic genes and 4 cell cycle regulators). Based on qPCR in fbf mutants and wild type (all in glp-1(gf) background, Figure 1G), authors argue that FBF-1 destabilize these target mRNAs, while FBF-2 promotes their accumulation. This data lays the foundation for their subsequent analysis. However, the data need improvement in several ways:
i) The authors have chosen linear trend analysis to show that, for all the mRNA tested, levels are higher in fbf-1(-) > wild type > fbf-2. While trend looks to be true, but given that control, tbb-2 mRNA levels are reduced by almost half in fbf-2 mutant compared to wild type, with a possible exception cyb-2.2, none of the mRNAs tested can be different in fbf-2 mutant compared to wild type. Based on the graph one can only argue that FBF-1 destabilize mRNA levels but FBF-2 does not have any appreciable effect.
As suggested, we now replaced linear trend analysis with comparisons between the wild type (glp-1gf) and fbf-2 mutant backgrounds. Additionally, we have replaced tbb-2 with a different housekeeping gene, unc-54 (myosin heavy chain). These new data are reported in Figure 5—figure supplement 1B.
ii) The statistical labels in Figure 1G is inconsistent with the statement made in the result. The authors claimed that other than cyb-1, all the other targets mRNA expression are increased in fbf-1(-), but reduced in fbf-2(-). While in Figure 1G, the statistical significance label for htp-2 and cyb-1 are EXACTLY the same. In addition, the authors are encouraged to explain why one-way ANOVA was chosen for their statistical analysis in the Materials and methods section. A pairwise comparison between wild type and each fbf single mutant should be performed for all the mRNAs tested; if there is a significant difference then one can make a conclusion.
ANOVA analysis is required to determine whether there is statistically significant difference among any of the data sets in groups of 3 or more sets, but does not indicate which data sets are different from each other. Therefore, it returns same significance for a group where fbf-1(-) is significantly different from both wt and fbf-2(lf) and a group where fbf-2(-) is different from fbf-1(-) and wt. Following detection of significant difference by ANOVA, a post-hoc test is used to determine which specific samples are different from each other. In response to the reviewer’s suggestion, we now follow ANOVA analysis by a Student’s t-test to compare the abundance of FBF target mRNAs in fbf-2; glp-1 mutants to those of glp-1 single mutants instead of linear trend analysis.
iii) The authors indicate that they are studying these targets' regulation in germline tumor, however the RNA is from whole worms. The authors are encouraged to explain their approach.
Thank you for this suggestion. Materials and methods section now includes the explanation that FBF targets investigated in this paper are expressed in the germline and not in the somatic cells of the adult worms. Therefore, RNA abundance in whole worm lysates reports on germline-specific expression of these mRNAs.
3) Repression of cyclin B by FBF limits accumulation of germline SPCs.
The authors have mutated all three FBEs in cyb-2.1 and have successfully shown that CYB-2.1 protein levels have become higher in immunoblot. They also show that this mutation causes an increase of SPC size in fbf-2 mutant and this can be reverted by depleting cyb-2.1. Based on this the authors conclude that "levels of B-type cyclins limit SPC proliferation rate in fbf-2(lf) and disruption of FBF-1-mediated repression of a single cyclin B gene is sufficient to affect the size of germline SPC zone".
There are multiple issues with this result:
i) C. elegans has ~12 puf genes (Prasad et al., 2016) and most of them have not been fully characterized; thus it is possible that mutated FBEs might be recognized by one or more of these puf proteins rather than FBF. Immunostaining and quantification of CYB-2.1 in extruded germlines of the fbf-1 mutant and the fbf-2 mutant would be required to claim that there is reciprocal regulation by the FBF proteins, rather than other puf proteins.
We have restated our conclusions to reflect that FBEs in cyb-2.1 3’UTR might mediate repression through recruitment of other PUF proteins expressed in progenitor cells, although the documented differences between PUF proteins’ preferred binding sites (Bernstein et al., 2005; Opperman et al., 2005; Koh et al., 2009) would suggest that FBEs are likely recognized by FBFs rather than other PUFs expressed in the SPCs.
ii) Mutating FBEs in cyb-2.1, results in its higher expression. Rationale for checking the effect of its overexpression in fbf-2(-) background, instead of wild type, is not clear. Since there are no FBEs in cyb-2.1fbm, its effects are expected to be fbf-agnostic.
We now emphasize the rationale that the fbf-2(-) with its slow cell cycle rate provides a sensitized background to detect the effects of cell cycle regulator overexpression, since it is unclear whether cell cycle rate can be accelerated beyond that of the wild type.
iii) Subsection “Repression of cyclin B by FBF limits accumulation of germline SPCs” paragraph three authors have concluded that increase in SPC zone size in fbf-2; cyb-2.1fbm vs fbf-2 is due to increased proliferation and also the meiotic entry rate is unchanged between two mutants. They have not provided any evidence for either of these two possibilities, i.e. proliferation rate has increased in fbf-2; cyb-2.1fbm and meiotic entry rate is same in fbf-2; cyb-2.1fbm as compared to fbf-2.
Thank you for this suggestion. New experiments in response to these suggestions are summarized in Essential Revisions. As suggested, we now include the estimation of stem cell proliferation rate comparing fbf-2(lf) with the strains expressing wild type or mutant 3xflag::cyb-2.1 transgenes. These new experiments suggest that cell cycle rate is accelerated in fbf-2(lf); 3xflag::cyb-2.1fbm compared to fbf-2(lf) (Figure 2D). By contrast, meiotic entry rate of fbf-2(lf) is not affected by expression of either cyb-2.1 transgene (Figure 2E).
4) FBF-1 promotes deadenylation of its target mRNA.
The authors moved back to cyb-2.1 to show that poly A tails are affected in fbf mutants without giving any rationale. The assay should be performed on a few more genes if any generalized statement statement is made, otherwise it must be restricted to cyb-2.1. Also wild type (glp-1(gf)) is missing in the assay, which makes it hard to interpret the data. Is FBF-1 alone or both FBFs responsible for deadenylation of cyb-2.1 mRNA or if FBF-1 and FBF-2 have opposing effect on cyb-2.1 mRNA?
As suggested, we now include the wild type (glp-1(gf)) as a control for PAT-PCR assay (Figure 5), and find that FBF-1 and FBF-2 have opposing effects on cyb-2.1 poly(A) tail. Furthermore, we have included PAT-PCR assay for an additional FBF target, htp-1. Our choice of transcripts for analysis represents both cell cycle-related transcripts and differentiation-related transcripts. We focused on the targets for which we could obtain reliable and robust amplification of poly(A) tails.
5) While the authors present some solid lines of evidence that FBF-1 and FBF-2 are different (meiotic entry rate, FBF-1 acts with CCR-4/NOT to de-adenylated cyb-2.1 and there are specific sequences in FBF-2 that block this interaction), they are clearly redundant with each other for the stem cell fate and repression of htp-2. Thus, Figure 8 would be more useful to the community if their redundant function was also included.
The revised model (Figure 8) now emphasizes that both FBFs repress the bound target mRNAs and both FBFs can maintain SPCs on their own (thus highlighting their redundant function).
6) FBF-1 and FBF-2 do not explain the spatial control of expression in the SPC zone and thus balance between proliferation and differentation, which is through glp-1 targets lst-1 and sygl-1. It is surprising that there is no mention of lst-1 and sygl-1-1 in the Discussion, particularly since they are co-factors of FBF-1 and FBF-2 (Haupt et al., 2019, Shin et al., 2017).
We now include LST-1 and SYGL-1 in the Discussion. We have not investigated whether the changes in SPC zone size in fbf mutants are associated with altered expression of LST-1 or SYGL-1 (that are curiously themselves FBF targets), and it would be an interesting new direction to explore.
Reviewer #3:
[…] My critique of this manuscript can be broken into two categories: (1) it would benefit from some changes in writing to increase clarity and accessibility for non-experts and (2) there are a few questions that I had regarding experimental issues, some related to interpretation, which could be addressed in the text (points D, and F-H below) and some related to additional controls/experiments that would substantiate the authors' main conclusions (points A-C, E, and F). The suggested experiments are not substantially outside of those that the authors have already conducted and I think that most of them (enough to satisfy the majority of my major comments) could be performed within a reasonable time frame.
Major comments:
1) Writing/interpretation:
As someone who works in the germline, but not on this pathway in particular, I found that this paper was often hard to digest. It would be very helpful if the authors could make sure that their rationale for each experiment and its interpretation is accessible to a reader not intimately familiar with the field. For example, the difference in proliferative zone (PZ) size in fbf-1(lf) and fbf-2(lf), I think pointing out here how a change in PZ size shows a shift in the balance between proliferation and differentiation, as if both were slowed down or sped up in complete coordination, the size of PZ would likely be constant. In addition, explaining why experiments were designed in a certain way and why the (perhaps) easier experiment is not feasible would be very helpful.
We have revised to reflect that the changes in SPC zone size in fbf-1 and fbf-2 mutants suggest that each individual FBF protein does not perfectly balance the cell division and meiotic entry. Rather, antagonistic FBF activities combine to simultaneously modulate stem cell division with the rate of meiotic entry. Although our model might be oversimplified, it provides a framework for future exploration of factors controlling SPC dynamics.
Additionally, we have clarified the rationales for the reported experiments.
I also believe that the authors need to revise their language in several instances to avoid conflating two different questions/fields. For example, I found the use of "differentiation demands" confusing, because, for me, "differentiation demands" implies reproductive demand, i.e. the need for mature gametes. In both the C. elegans germline (e.g. Narbonne, et al., 2015) and in other stem cell populations (e.g. Hsu, Li and Fuchs, 2014), signaling from differentiated/differentiating progeny to their stem cell parents plays a significant role in regulating stem cell proliferation, such that proliferative output matches the demand for differentiated progeny. I do not think that the authors are addressing this type of "feedback" signaling – i.e. they do not show that the rate of meiotic entry regulates the rate of proliferation, but rather that these two things are coordinated. I think that it is important to clarify their questiion to avoid confusion and/or erroneous expectations from people, such as myself, who are more used to thinking about feedback mechanisms. There are several instances of this throughout, including: "Complementary activities of FBF-1 and FBF-2 combine to fine tune SPC proliferation and differentiation to respond to proliferative demands of the tissue."
Thank you for this suggestion. We have clarified our writing to convey that the focus here is on coordinate regulation of cell division rate and meiotic entry.
2) Experimental issues:
A) Figure 1. The cell cycle phenotype of fbf-1(lf) is puzzling. Since there are no quiescent cells in the germline, an increase in mitotic index (MI) ought to reflect either a faster passage through the mitotic cell cycle or a longer duration of mitosis. The authors report no change in the length of G2, which leads me to wonder whether S phase is accelerated or mitosis is delayed (I believe there are assays to assess both (S phase, Fox and Schedl, 2011) and M phase (Gerhold et al., 2015)). This result is also odd in light of their observation that the transcript levels of most of the Cyclin Bs (except for CYB-1), are highly elevated – i.e. why is the duration of G2 unaffected despite higher levels? The authors explanation in the discussion that a higher MI could be due to a smaller proliferative population, due to accelerated meiotic entry, would predict (I think) that the stem cell pool should eventually be lost – i.e. in older fbf-1(lf) adults, do they see a progressive loss of proliferative cells? As this interpretation (i.e. FBF-2, rather than FBF-1, being primarily responsible for cell cycle regulation) is key to their model figure at the end, it bears a closer look.
Thank you for these suggestions. We addressed many of these in response to the Essential Revisions. Briefly, we determined the S-phase index of the adult germlines, and find that S phase is not accelerated in fbf-1(lf). We were not able to determine the duration of M-phase since we lack the required live imaging microfluidic set up. Additionally, an assay measuring germline stem cell accumulation during larval development suggested that overall division rate of fbf-1(lf) and wild type germ cells is the same.
Furthermore, we clarified the potential explanation for higher mitotic index: the total SPC cell count includes non-cycling cells in meiotic S-phase (they cannot be excluded because the field lacks specific molecular markers for this developmental stage). Accelerated meiotic entry in fbf-1(lf) might selectively reduce this non-cycling cell population due to their faster transit through the meiotic S-phase. This would lower overall SPC cell counts (without reducing the number of proliferative cells), and thus lead to a higher MI.
B) Also Figure 1. I am unsure as to how to interpret the RT-qPCR data. The authors state that their negative control (tbb-2) is unaffected, yet in fbf-2(lf) transcript levels appear to be at least a strongly depressed as those of the genes that they argue are maintained by FBF-2. It is possible that I am missing something here, but it would be good if the authors could address/explain this, as it is visually striking/distracting in their data/graph and if it is accurate, i.e. that in fbf-2(lf) transcript levels are generally lower, it is problematic for the interpretation that FBF-1 and FBF-2 have reciprocal effects on shared target transcripts. It is also not entirely clear to me what the linear trend test adds to the data interpretation, some clarification/discussion would be helpful.
We addressed these suggestions in the Essential Revisions. Briefly: First, the transcript levels in Figure 1G (now Figure 5—figure supplement 1B) are normalized to actin (act-1), so a decrease in FBF target transcripts in fbf-2(lf) does not reflect generally lower transcript levels. Additionally, we have tested other housekeeping mRNAs to provide an alternative negative control. We now replace tbb-2 with unc-54, which abundance changes in the opposite direction to that of FBF targets. Second, as suggested, we now replaced linear trend analysis with comparisons between the wild type (glp-1gf) and fbf-2 mutant backgrounds.
C) In Figure 2, the authors address the hypothesis that slow G2 and low MI in fbf-2(lf) is due to FBF-1-mediated translational repression of cyb-2.1. The authors state that protein levels for CYB-2.1 with mutated FBF binding elements (FBEs) are elevated, but they cannot detect protein levels for the WT version, which could imply that it is not expressed at all. It would be helpful if the authors could should that both transgenes are expressed by looking at transcript levels. I also wonder if this result predicts that in wildtype germlines, expression of cyb-2.1 with mutated FBEs, but not wild-type cyb-2.1 would affect cell cycle progression? Finally, this hypothesis also seems to predict that a partial RNAi of FBF-1 in a fbf-2(lf) background, should rescue, at least to some extent, the fbf-2(lf) phenotype. If this experiment is technically feasible, it would greatly strength the authors argument.
Thank you for these excellent suggestions!
1) We have generated a new cyb-2.1wt transgene that we can now detect both at the transcript and at the protein level, and report these new data in Figure 2 (Figure 2B, C). Crossing the new cyb-2.1wt into fbf-2(lf) mutant did not affect the stem cell dynamics (Figure 2D-G).
2) As mentioned in the response to reviewer 2 (3ii), we now emphasize the rationale that the fbf-2(-) with its slow cell cycle rate provides a sensitized background to detect the effects of cyclin overexpression.
3) We have not been able to partially knock down fbf-1 to address whether this would be able to rescue fbf-2(lf).
D) In Figure 3, the authors use a translational reporter for the htp-2 3'UTR, which their RT-qPCR results suggest should be derepressed in fbf-1(lf), but they do not seem to see this and I am unsure as to why not? In addition, their results strongly suggest that the change in PZ size in fbf-2(lf) requires CCR4-NOT complex members, which supports the hypothesis that FBF-1 activity with CCR4-NOT is leading to transcript loss. However, if FBF-1 acts with CCR4-NOT to regulate transcript levels, shouldn't RNAi of CCR4-NOT complex members phenocopy fbf-1(lf)? It would be helpful if the authors could address these questions. This may be another instance of where the data/results are not fully accessible to a non-expert.
We now explain that FBF targets are not derepressed in fbf-1 mutant even though their mRNAs are accumulating in cytoplasmic foci (Merritt and Seydoux, 2010; Voronina et al., 2012) since FBF-2 on its own is capable of translational repression. However, FBF-2-mediated repression might be less effective since partial deprepression of several targets including gld-1 has been previously observed in fbf-1(lf) mutants (Crittenden et al., 2002, Brenner and Schedl, 2016). To test whether fbf-1 requires CCR4-NOT for function, we assess whether fbf-2(lf) after CCR4-NOT subunit RNAi show the same range of phenotypes as fbf-1(lf) fbf-2(lf). These phenotypes include derepression of the htp-2 3’UTR reporter, sterility, and failure to initiate oogenesis. This reasoning is now included in the Results section.
E) In Figure 5, the authors quantify the polyA tail length of cyb-2.1 and show that it is shorter in fbf-2(lf), suggesting that it is being deadenylated, presumably by FBF-1/CCR4-NOT. It would strengthen the authors conclusion substantially, if they could show that the change in polyA tail length was rescued by depletion of CCR4-NOT complex members. It is also unclear to me why the authors compared fbf-1(lf) to fbf-2(lf) rather than to wildtype? I also wonder whether tbb-2 is the best control as the detection levels seem very low which could make slight changes in peak detection challenging.
As suggested, we have added the wildtype control (glp-1(gf)) to the polyA tail length assay (Essential Revisions), and now use unc-54 myosin instead of tbb-2 as the control. These changes resulted in revised Figure 5. While testing whether changes in polyA tail length in fbf-2(lf) are dependent on CCR4-NOT would provide additional evidence supporting FBF-1/CCR4-NOT regulatory association, we did not have the time to perform the assay as large-scale culture of fbf-2(lf); glp-1(gf) nematodes on RNAi is challenging. Nevertheless, multiple lines of evidence presented in the paper strongly support this conclusion.
F) Figure 6: I think that this a key place where the authors interpretation could be clarified somewhat. I believe that in Figure 6, the fact that FBF-2 without variable regions (VRs) 1, 2 and 4 cannot fully rescue the fbf-2(lf) phenotype in either a fbf-2(lf) or a fbf-1(lf) fbf-2(lf) background suggests that either this version of FBF-2 is only partially functional or it is acting like FBF-1. The authors then provide support for the latter interpretation by showing that the activity of FBF-2(vrm) requires CCR4-NOT and can also partially rescue fbf-1(lf). This last piece of data is in Figure 6—figure supplement 1, but, in my opinion, should be moved into the main figure panel. This also suggests that FBF-1 may be able to act with CCR4-NOT without any of its own variable regions, which (I think?) is somewhat surprising. The authors could test this by using a similar approach.
As suggested, we have moved the fbf-1(lf) rescue from Figure 6—figure supplement 1B to the main Figure 6. We agree that our data suggest that FBF-1’s variable regions flanking the RNA-binding domain are permissive for acting with CCR4-NOT. One simple possibility is that VR4 of FBF-2 includes a 16-amino acid C-terminal extension that might be sterically interfering with the recruitment of CCR4-NOT. However, rigorously testing this and other possible determinants allowing FBF-1/CCR4-NOT cooperation is beyond the scope of the present work.
G) Figure 7: For the domain swapping experiments here, I believe the main conclusions are that VR3 (in the RNA-binding domain) is not required for FBF-1 activity, and that VR4 from FBF-2 is sufficient to prevent FBF-1 from acting with CCR4-NOT to regulate polyadenylation, as FBF-1 with FBF-2's VR4 cannot rescue fbf-1(lf) in a fbf-1(lf) or a fbf-1(lf), fbf-2(lf) background, while FBF-1 with FBF-2's R3 can. However, what I do not understand is why FBF-1 with FBF-2's VR4 rescues fbf-2(lf) (Figure 7—figure supplement 1)? Does this mean that it is acting as a dominant negative, since presumably wild-type FBF-1 is still present in this background?
We propose that FBFs bind to the same mRNAs, and either compete for the same binding sites or simultaneously bind to the transcripts that contain multiple FBEs. Either mechanism would explain how change-of-function mutants such as FBF-2vrm and FBF-1(FBF-2vr4) are able to rescue the loss-of-function of the non-cognate fbf. We do not think that change-of-function mutants are actually dominant-negative (i.e., poison the function of the original FBF), because in that case a combination of FBF-1(FBF-2vr4) with the wild type fbf-1 in the fbf-2(lf) background would disrupt the activity of the sole remaining wt FBF-1 and produce a phenotype similar to a loss of function of both fbfs (loss of SPCs in the adult).
H) Figure 8: The model figure as currently drawn doesn't really help me understand the authors' mains conclusions/findings. It would be great if this could be re-worked to reflect the effects of FBF-1 and FBF-2 on the stem cell population (i.e. perhaps in relation to PZ size which is their phenotypic readout for most experiments) and how these activities lead to a balance between stem cell maintenance and differentiation.
The model in Figure 8 now depicts that FBF effects on the extent of the mitotic zone result from their regulation of stem cell dynamics (how fast the stem cells are able to divide and the rate of meiotic entry).
https://doi.org/10.7554/eLife.52788.sa2Article and author information
Author details
Ekaterina Voronina
Funding
National Institute of General Medical Sciences (GM109053)
- Ekaterina Voronina
National Institute of General Medical Sciences (P20GM103546)
- Ekaterina Voronina
American Heart Association (18PRE34070028)
- Xiaobo Wang
Montana Academy of Sciences
- Xiaobo Wang
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
Acknowledgements
We thank the members of Voronina laboratory for insightful discussions and Geraldine Seydoux for comments on our manuscript. We are grateful to Ella Baumgarten and Jessica Bailey for help with cloning and crosses. We appreciate Ariz Mohammad for sharing the modified R script (originally from the Kimble lab) and instructions on using R for cell counts. Some nematode strains used in this study were provided by the Caenorhabditis Genetics Center funded by the NIH (P40OD010440). Confocal microscopy was performed in the University of Montana BioSpectroscopy Core Research Laboratory operated with support from NIH awards P20GM103546 and S10OD021806. This work was supported by the NIH grants GM109053 to EV and P20GM103546 (S Sprang, PI; EV Pilot Project PI), American Heart Association Fellowship 18PRE34070028 to XW, and Montana Academy of Sciences award to XW.
Senior Editor
- Marianne E Bronner, California Institute of Technology, United States
Reviewing Editor
- Yukiko M Yamashita, University of Michigan, United States
Version history
- Received: October 16, 2019
- Accepted: August 14, 2020
- Accepted Manuscript published: August 17, 2020 (version 1)
- Version of Record published: September 2, 2020 (version 2)
Copyright
© 2020, Wang et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.
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Antagonistic control of Caenorhabditis elegans germline stem cell proliferation and differentiation by PUF proteins FBF-1 and FBF-2
eLife 9:e52788.
https://doi.org/10.7554/eLife.52788
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