All C. elegans strains were cultured at 20°C (unless specified otherwise) on NGM media seeded with E. coli OP50 as previously described (Brenner, 1974). To synchronize growth of C. elegans animals, embryos were isolated by bleaching, washed in M9 buffer, and incubated overnight at room temperature. The resulting synchronized L1s were dropped on seeded plates and grown until adulthood. For auxin treatment, gravid adults were allowed to lay eggs on NGM plates containing 4 mM Naphthaleneacetic Acid (K-NAA, PhytoTech) and the progeny were grown to the L4 stage, picked to new plates containing auxin, and grown for an additional 24 h prior to imaging. The strains used in this study are listed in Supplementary Table S1.

The high-copy mgIs70[Pvit-3::GFP] and single-copy rhdSi42[Pvit-3::mCherry] transgenes have been previously described (Dowen et al., 2016; Torzone et al., 2023). The fbxl-5 rescue constructs were generated by fusing either the col-10 promoter (chromosome V: 9,166,416–9,165,291; WS284) or the vha-6 promoter (chromosome II: 11,439,355–11,438,422; WS284) to a mCherry::his-58::SL2::fbxl-5 cassette (fbxl-5 gene: 2,330 bp ORF and 176 bp of 3′UTR) within the pCFJ151 plasmid via Gibson assembly to generate plasmids pRD99 and pRD98, respectively (Frøkjær-Jensen et al., 2008; Gibson et al., 2009). The sequence-verified plasmids were microinjected into lin-4(e912); fbxl-5(rhd43); mgIs70[Pvit-3::GFP] animals at 20 ng/μL, along with 2.5 ng/μL pCFJ90(Pmyo-2::mCherry) and 77.5 ng/μL of 2-Log DNA ladder (New England BioLabs), to generate the DLS327 and DLS330 strains. To image these strains, L1 animals expressing Pmyo-2::mCherry were hand-picked on a SMZ-18 Stereo microscope with fluorescence, grown to adulthood, and were then randomly selected for fluorescence imaging under bright field using a standard Stereo microscope. To generate rhdIs2 and rhdIs4, wild-type animals carrying the extrachromosomal array Ex[Pvha-6::mCherry::his-58::SL2::fbxl-5 + Pmyo-2::mCherry] were gamma-irradiated using a¹³⁷Cs source (∼3000 rad), resulting in random integration of the array into the genome. The resulting two independent lines were backcrossed at least three times. The fbxl-5 promoter (chromosome V: 7,746,887–7,748,867; WS288) was amplified by PCR and fused to a mCherry::unc-54 3′UTR fragment by PCR fusion (Hobert, 2002) to yield a Pfbxl-5::mCherry::unc-54 3′UTR PCR fragment, which was microinjected into wild-type animals at 75 ng/μL, along with 5 ng/μL Pmyo-3::GFP and 20 ng/μL of 2-Log DNA ladder (New England BioLabs), to generate the DLS344 strain. All strains carrying the mgIs70 or rhdSi42 transgenes were imaged with a Nikon SMZ-18 Stereo microscope equipped with a DS-Qi2 monochrome camera, while the DLS344 strain was imaged on a Nikon Eclipse E800 microscope.

Strains carrying either the mgIs70 or rhdSi42 reporter were grown asynchronously, L4s were picked to new plates, and animals were imaged 24 h later. Alternatively, synchronized animals were grown for 72 h before imaging. Day 1 adults were mounted on a 2% agarose pad with 25 mM levamisole and imaged with a Nikon SMZ-18 Stereo microscope equipped with a DS-Qi2 monochrome camera. The worm bodies were outlined in the brightfield channel and the mean intensities (i.e., gray values) were calculated using the GFP (mgIs70) or mCherry (rhdSi42) channel using the Fiji 2.14.0 software (Schindelin et al., 2012). Body size measurements (pixels/worm) were simultaneously performed, and the data were converted to mm² based on the known imaging settings. The fluorescence and body size data were plotted in Prism 10 and a one-way ANOVA with a Bonferroni correction was performed to calculate p values. All tabular data and the associated statistical analyses can be found in Supplementary Table S5.

Genomic edits were performed by microinjection of Cas9:crRNA:tracrRNA complexes (Integrated DNA Technologies) into the germline as previously described (Ghanta and Mello, 2020). The crRNA guide sequences are listed in Supplementary Table S2. To generate missense, nonsense, or deletion mutations, single-stranded oligodeoxynucleotides were used as donor molecules. The deletion edits employed two sgRNAs positioned at each end of the deletion site and an oligo that bridged the predicted repair junction. To insert the 3xFLAG::AID cassette into the 5′ end of the fbxl-5 gene, dsDNA donor molecules with ∼40 bp homology arms were prepared by PCR using Q5 DNA Polymerase (New England BioLabs) and purified using HighPrep PCR Clean-up beads (MagBio) according to the manufacturers’ instructions. The melted and reannealed DNA repair templates were prepared and microinjected into the germline as previously described (Ghanta and Mello, 2020).

E. coli HT115 (DE3) strains carrying plasmids for fbxl-5 or lin-29 RNAi knockdown (Ahringer RNAi library), or the control strain containing the empty L4440 plasmid, were grown for ∼19 h at 37°C in Luria–Bertani media containing ampicillin (100 μg/mL). The bacteria were concentrated by 20–30x via centrifugation, seeded on NGM plates containing 5 mM ​isopropyl-β-D-thiogalactoside (IPTG) and 100 μg/mL ampicillin, and maintained at room temperature overnight to induce dsRNA expression. Synchronized L1s were dropped on RNAi plates, grown at 20°C until they were day 1 adults (∼72 h), and imaged. For the skr/cul RNAi screen, we screened cul-1,2,3,4,5,6 and skr-1,2,3,5,7,11,12,13,14,16,17,19,20,21 using existing clones from the Ahringer RNAi library and assessed their ability to confer any detectable increase in Pvit-3::GFP reporter expression (relative to the L4440 control) in the DLS447 strain. The skr-4,6,8,9,10,15,18 genes were not screened due to a lack of a sequence-verified RNAi clone in our collection. RNAi conditions that prevented development to the adult stage (e.g., skr-1 RNAi) were not included in the analysis.

Animals were grown on standard NGM E. coli OP50 plates for at least two generations without starvation prior to lifespan and brood size assays. Longitudinal lifespan assays were performed at 25°C in the presence of 50 μM FUDR as previously described (Torzone et al., 2023). Approximately 150 animals were assayed for each genotype and a log-rank test was used to determine significance. Lifespan assays were repeated at least twice with similar results. For brood size assays (20°C), L4 animals (n = 11 per genotype) were picked to individual plates and transferred daily throughout the reproductive period. Progeny were counted as L3, L4s, or as adults. The lowest brood size for each genotype was dropped from the analysis due to potential injury and the data were reported as the mean +/- the standard deviation. The tabular data and statistical analyses can be found in Supplementary Table S5.

Animals were fixed in 60% isopropanol and stained for 7 h with 0.3% Oil Red O exactly as previously described (Torzone et al., 2023). Next, animals were mounted on 2% agarose pads and imaged with either a Nikon SMZ-18 Stereo microscope equipped with a DS-Qi2 monochrome camera (intensity quantification) or a Nikon Ti2 widefield microscope equipped with a DS-Fi3 color camera (representative color images). Quantification of Oil Red O staining was performed in Fiji 2.14.0. Animals were manually outlined, the mean gray value was measured in the worm area, and the value was subtracted from 65,536 (maximum gray value for 16-bit images). The data were plotted in Prism 10 and a one-way ANOVA with a Bonferroni correction was performed to calculate p values. The tabular data and statistical analyses are shown in Supplementary Table S5.

Synchronized day 1 adult animals were harvested in M9 buffer, washed, and flash frozen. Isolation of total RNA was performed using Trizol Reagent (Thermo Fisher) with chloroform extraction, followed by isopropanol precipitation. Synthesis of cDNA was performed by oligo (dT) priming using the SuperScript IV VILO Master Mix with ezDNase Kit per the manufacturer’s instructions (Thermo Fisher). Quantitative PCR was performed exactly as previously described (Dowen, 2019). The qPCR primer sequences are listed in Supplementary Table S3. Data from at least three independent experiments were plotted in Prism 10 as the mean fold change relative to wild-type with the standard error of the mean (SEM). The individual Ct values and the statistical analyses can be found in Supplementary Table S5.

Synchronized L1 animals expressing the 3xFLAG::AID::FBXL-5 protein (DLS874 or DLS889) were grown at 20°C for either 36 h (L3s), 48 h (L4s), 72 h (day 1 adults), or 96 h (day 2 adults) before harvesting them in M9 buffer. The animals were washed three times and pellets were frozen in liquid nitrogen. Whole cell lysates and protein concentrations were determined exactly as previously described (Torzone et al., 2023). The protein samples (50 μg) were resolved by SDS-PAGE, transferred to a PVDF membrane, blocked in 5% nonfat dry milk (BioRad), and probed with either anti-FLAG M2 (F1804, Sigma) or anti-Actin (ab3280, Abcam) antibodies. The developmental time course experiment was performed twice with similar results.

The GR2123 strain was mutagenized with ethyl methanesulfonate (M0880, Sigma-Aldrich) exactly as previously described (Dowen et al., 2016). Suppressor mutants displaying increased mgIs70[Pvit-3::GFP] reporter expression were selected from ten distinct mutagenesis pools and were singled to individual plates. Mutations that suppressed the egg-laying defects of lin-4(e912) were presumed to disrupt lin-14 function and were not selected for subsequent studies.

Suppressor strains were crossed to the GR2122 strain (mgIs70[Pvit-3::GFP]) and F2 recombinants that displayed the characteristic egl and lon phenotypes of the lin-4(e912) mutant, and that also showed elevated mgIs70 expression, were singled. Starved animals from the individual plates were then pooled and the genomic DNA was prepared with the Qiagen Gentra Puregene Tissue Kit (Doitsidou et al., 2010). The DNA-Seq libraries were prepared using the NEBNext DNA library preparation kit according to the manufacturer’s instructions (E6040, New England Biolabs) and the libraries were sequenced on an Illumina HiSeq 4000 instrument. Identification of candidate suppressor mutations was performed using in-house scripts according to previously described methods (Minevich et al., 2012).

Wild-type and mutant animals were reared on 10 cm NGM agarose plates (5000 animals/plate) until they reached the first day of adulthood. RNA was isolated using Trizol Reagent as described above (see quantitative PCR). The mRNA-Seq libraries were prepared with 1 μg of total RNA using the TruSeq RNA Library Prep Kit v2 per the manufacturer’s instructions (Illumina). Three independent biological replicates were prepared for each condition and libraries were sequenced on an Illumina HiSeq 4000 instrument (single-end, 50 bp) at the High Throughput Genomic Sequencing Facility at the University of North Carolina at Chapel Hill. The reads were aligned to the C. elegans genome (WS260), and the feature read counts were compiled exactly as previously described (Dowen, 2019). RPKM values and differentially expressed genes (1% FDR) were determined using the DESeq2 algorithm (Love et al., 2014). Lists of differentially expressed genes and RPKM values can be found in Supplementary Table S4. Tissue-specific enrichment for differential expression was calculated as the number of observed vs. expected differentially expressed genes using previously generated gene lists for each major tissue (Kaletsky et al., 2018). The gene ontology analysis was performed using WormCat (Holdorf et al., 2020). All scatter and violin plots displaying differential expression were generated using the DESeq2 RPKM values and the data were plotted in Prism 10. All tabular data and associated statistical analyses can be found in Supplementary Table S5. The raw and processed mRNA-Seq data have been deposited in GEO (GSE248602).

The heterochronic gene lin-4 encodes a miRNA that governs developmental timing by controlling larval cell divisions in the C. elegans hypodermis (Lee et al., 1993). Furthermore, lin-4 acts non-cell-anonymously to license expression of the vitellogenin genes (vit-1 through vit-6) in the intestine at the L4 larval to adult transition (Dowen et al., 2016). To identify genes that act in the lin-4 vitellogenesis pathway, we performed an EMS mutagenesis screen to identify mutations that suppress the vitellogenesis defects observed in the lin-4(e912) mutant using a Pvit-3::GFP vitellogenesis reporter as a readout (Supplementary Figure S1A). We selected lin-4 suppressor mutants that displayed elevated Pvit-3::GFP expression but that also maintained the egg-laying (egl) and long body length (lon) defects of the lin-4(e912) mutant, which selects against known lin-4 suppressors (e.g., lin-14 alleles). After backcrossing to remove unlinked background mutations, we performed whole genome sequencing of the lin-4 suppressor strains and identified candidate causative mutations using established bioinformatic approaches (Minevich et al., 2012). This resulted in the identification of two putative loss-of-function mutations in the uncharacterized T05B11.1 gene, which were recovered in two genetically distinct suppressor strains (Supplementary Figure S1B).

The T05B11.1 protein contains a F-box domain at the amino terminus and leucine-rich repeats (LRRs) that span most of the protein (Paysan-Lafosse et al., 2023). Thus, we will subsequently refer to the T05B11.1 gene as fbxl-5 (F-box leucine rich repeat containing protein 5). F-box proteins canonically function in protein ubiquitination, typically in concert with the Skp1 and Cullin proteins (the SCF complex), which together catalyze E3 ubiquitin ligase activity (Skaar et al., 2013). The F-box domain, and in some cases the LRRs as well, specifically recognizes target substrates and mediates poly-ubiquitination of proteins, resulting in 26S proteasomal degradation (K11/K48 linkages) or altered protein function/localization (K63 linkages). Although there is no clear mammalian orthologue of FBXL-5, highly similar proteins are found in other Caenorhabditis species, and it is possible that functional orthologues may exist in other species.

To characterize the role of FBXL-5 in vitellogenesis, we first compared expression levels of the Pvit-3::GFP reporter between the lin-4(e912) single mutant and the lin-4(e912); fbxl-5(rhd43) double mutant, finding that while reporter expression is increased in lin-4(e912) animals upon loss of fbxl-5 it does not reach that of wild-type (Figure 1A). Notably, the rhd43 lesion is a nonsense mutation and is thus predicted to be a strong loss-of-function or null allele. Consistently, knock-down of fbxl-5 by RNAi partially suppressed the Pvit-3::GFP expression defects observed in the lin-4 mutant (Supplementary Figure S1C). To rule out the possibility that the high-copy Pvit-3::GFP reporter lacks the sensitivity to robustly detect the fbxl-5 suppression phenotype, we employed a single-copy Pvit-3::mCherry reporter, which closely reflects endogenous vit-3 levels (Torzone et al., 2023), and generated large deletions within the fbxl-5 locus using CRISPR/Cas9 genomic editing (Supplementary Table S2; Supplementary Figure S1B). While each of the three fbxl-5 deletion alleles tested suppressed the lin-4(e912) mutant defects in Pvit-3::mCherry expression, they did not restore levels to those seen in wild-type animals (Figures 1B, C). Interestingly, all fbxl-5 deletion mutations also reduced the body size of the lin-4(e912) mutant (Figure 1D). Consistently, the fbxl-5(rhd43) mutation conferred smaller body sizes in otherwise wild-type animals (Supplementary Figure S2), suggesting that FBXL-5 may regulate additional developmental or metabolic processes outside of vitellogenesis.

Impairment of hypodermal lin-4 alters the expression of other developmental timing genes resulting in heterochrony. In particular, lin-4 mutants fail to properly express lin-29 at the larval to adult transition. Thus, we reasoned that loss of fbxl-5 may suppress the vit expression defects seen in the lin-29(n333) mutant. Using RT-qPCR to measure the endogenous vit transcripts, we found that mutation of fbxl-5 also partially suppressed the vitellogenesis defects in lin-29(n333) animals; however, loss of fbxl-5 on its own is not sufficient to increase vit transcripts in otherwise wild-type animals (Figure 1E). These data suggest that FBXL-5 may negatively regulate vitellogenesis only when developmental timing is impaired. Together, our results indicate that fbxl-5 acts genetically downstream or in parallel to lin-4/lin-29, possibly in the intestinal cells, to govern vitellogenin expression.

Although loss of fbxl-5 only partially suppressed the vitellogenesis defects in the lin-29 mutant, it is possible that FBXL-5 plays a broader role in regulating cellular metabolism or lipid allocation in the heterochronic mutants. Taking an unbiased approach, we employed mRNA sequencing (mRNA-Seq) to resolve transcriptome-level differences between the lin-29 single mutant and the lin-29; fbxl-5 double mutant (Supplementary Table S4). Remarkably, loss of fbxl-5 had no global impact on the expression levels of the 6,265 genes that are differentially expressed in the lin-29 mutant (R ² = 0.927, Figure 2A). Expression of the vit-3-5 genes, as well as a handful of other downregulated genes, did increase in the lin-29; fbxl-5 double mutant; however, their expression did not return to wild-type. Thus, FBXL-5 has very specific and limited effects on gene expression in the lin-29 heterochronic mutant.

To further define the role of FBXL-5 in wild-type animals, we performed mRNA-Seq on the fbxl-5(rhd43) single mutant. This analysis led to the identification of 828 genes that were differentially expressed upon loss of fbxl-5 (Supplementary Table S4). Mutation of fbxl-5 resulted in both increased and decreased mRNA levels for genes expressed in each of the major C. elegans tissues, which likely represents both the cell-autonomous and non-cell-autonomous effects of mutating fbxl-5 (Supplementary Figure S3). A gene ontology analysis revealed that the upregulated genes are enriched in ribosomal and mRNA function, while the downregulated genes are annotated to act in cellular metabolism, stress response pathways, or proteolysis (Figure 2B). Remarkably, expression of 196 of the 221 C. elegans ribosomal genes was increased in the fbxl-5 mutant (Figure 2C), suggesting that translation is globally upregulated upon loss of fbxl-5. Thus, FBXL-5 functions broadly in maintaining cellular energy and protein homeostasis, possibly by governing the activity of a core homeostatic regulator that senses metabolic or developmental signals to tune protein synthesis.

Since loss of fbxl-5 in an otherwise wild-type background fails to upregulate vit gene expression (Figure 1E), we reasoned that FBXL-5 levels may be specifically altered in the lin-4 and lin-29 heterochronic mutants. To test this hypothesis, we first measured fbxl-5 transcript levels at the L4 and adult stages in the lin-4(e912) mutant by RT-qPCR, finding that fbxl-5 expression is specifically upregulated in lin-4 days 1 adult animals (Figure 3A). Consistently, adult expression of fbxl-5 is also elevated in the lin-29 mutant relative to wild-type animals (Supplementary Figure S4A). To measure FBXL-5 protein levels, we inserted a 3xFLAG::AID cassette into the 5’ end of the fbxl-5 locus by CRISPR/Cas9 editing, which did not disrupt FBXL-5 function and enabled detection by Western blotting (Supplementary Figure S4B). This strain, however, did not respond to auxin treatment (Supplementary Figure S4B), possibly because the auxin-inducible degron (AID) tag on FBXL-5 is buried in a larger protein complex that was inaccessible to TIR1. We assessed FBXL-5 expression levels across development by Western blot, probing lysates from wild-type and lin-4(e912) animals isolated at different developmental stages with an anti-FLAG antibody. Consistent with our gene expression measurements, the lin-4 mutant ectopically expresses FBXL-5 at the first day of adulthood, which persists into the second day of adulthood (Figure 3B; Supplementary Figure S4C). We observed a similar FBXL-5 expression pattern for animals subjected to lin-29 RNAi (Supplementary Figure S4D). While wild-type animals had the highest amount of FBXL-5 expression at the L4 stage, FBXL-5 protein was reproducibly detected at the L3s stage (Figure 3B; Supplementary Figure S4C), suggesting that fbxl-5 may be upregulated at the L3-L4 transition and may serve a specific role during the L4 developmental stage. Together, these data indicate that heterochronic programs control fbxl-5 expression, suggesting that the heterochrony of the lin-4 and lin-29 mutants may be, at least in part, attributed to improper levels of FBXL-5 during development.

Since loss of fbxl-5 resulted in altered transcription of genes expressed in a variety of tissues, we reasoned that FBXL-5 could act either cell-autonomously or non-cell-autonomously to control vitellogenin expression in the intestine. Importantly, LIN-29 functions in the hypodermis to regulate vitellogenesis in a non-cell-autonomous fashion (Dowen et al., 2016), and it is possible that ectopic expression of fbxl-5 in the hypodermis is partially responsible for the impaired vitellogenin production that is observed in the lin-29 mutant. To investigate the tissue-specificity of FBXL-5, we first generated a transgenic strain expressing a Pfbxl-5::mCherry transcriptional reporter, comprised of a ∼2 kb fragment of the fbxl-5 promoter driving the mCherry gene, and inspected animals for mCherry expression by fluorescence microscopy. This analysis revealed that fbxl-5 is indeed weakly expressed in the intestine of wild-type young adults; however, it is also expressed in a small set of neurons in the head and in the body wall muscle (Figure 4A; Supplementary Figure S5A), which is consistent with the widespread transcriptional changes that we observed. Surprisingly, we did not observe Pfbxl-5::mCherry reporter expression in the hypodermis despite finding that hypodermal genes were downregulated in the fbxl-5 mutant (Supplementary Figure S3). However, it is possible that fbxl-5 hypodermal expression is relatively low compared to other tissues, a hypothesis that is supported by existing tissue-specific mRNA-Seq data (Kaletsky et al., 2018). Intriguingly, in wild-type animals intestinal Pfbxl-5::mCherry reporter expression continued to increase throughout adulthood while FBXL-5 protein levels decreased (Figure 3B; Supplementary Figure S5B), suggesting that fbxl-5 may be post-transcriptionally or post-translationally regulated; however, the long-term stability of mCherry protein may at least partially contribute to this elevated signal in older animals.

Despite being expressed in several tissues, we reasoned that FBXL-5 likely acts in the intestine or in the hypodermis (the site of lin-29 action) to regulate vitellogenesis. Thus, we generated intestine-specific and hypodermis-specific fblx-5 rescue constructs to test whether the fblx-5(rhd43) mutation could be rescued by tissue-specific expression of fblx-5. Rescue of fblx-5 in the intestine, but not in the hypodermis, reversed the suppressive effects of fblx-5(rhd43) in the lin-4 mutant background back to baseline Pvit-3::GFP levels (Figures 4B, C). Surprisingly, hypodermal fblx-5 over-expression further enhanced the suppressive effects of the rhd43 allele, suggesting that FBXL-5 might promote vitellogenin expression when expressed outside of the intestine. While the vitellogenesis phenotype was rescued by intestinal expression of fbxl-5, the small body size phenotype that we observed in the lin-4(e912); fblx-5(rhd43) double mutant was not rescued by intestinal or hypodermal fbxl-5 expression (Figure 4D), suggesting that FBXL-5 may act non-cell-autonomously to control body size.

Since fbxl-5 is ectopically expressed in the lin-4 and lin-29 heterochronic mutants, we reasoned that the consequences of FBXL-5 misexpression should be mirrored by constitutive over-expression of fbxl-5 in the intestine of otherwise wild-type animals. Thus, we integrated a Pvha-6::fbxl-5 extrachromosomal array into the genome via gamma irradiation and crossed the resulting strains to our vitellogenesis reporter. Consistent with our fbxl-5 rescue experiments, constitutive over-expression of fbxl-5 in the intestine is sufficient to abrogate Pvit-3::GFP expression (Figures 5A, B). Furthermore, fbxl-5 over-expression prevented the accumulation of intestinal lipids and restricted body size (Figures 5C, D), suggesting that FBXL-5 might impair nutrient absorption or stimulate lipid catabolism pathways. Lipid levels are reduced both at the L4 larval stage and during reproduction by fbxl-5 over-expression (Figure 5E), suggesting that FBXL-5 does not act exclusively on a larval-specific metabolic regulator.

Impaired vitellogenin production can indirectly extend organismal lifespan by stimulating intestinal autophagy and lipid catabolism (Murphy et al., 2003; Seah et al., 2016). Thus, we tested whether fbxl-5 over-expression, which severely reduces vitellogenin synthesis, increases longevity. Surprisingly, over-expression of fbxl-5 in the intestine reduced the lifespan of otherwise wild-type animals (Figure 5F), which is consistent with FBXL-5 playing a more widespread role in metabolic regulation. Moreover, the fbxl-5(rhd43) mutant also displayed a modest, but statistically significant, reduction in lifespan, which is consistent with the modest reduction in lipid levels and brood size that we observed in these mutant animals (Figures 5E, G). Predictably, intestinal over-expression of fbxl-5 reduced progeny production (Figure 5G), likely a consequence of the impaired vitellogenin production. Together, these data suggest that FBXL-5 can function broadly in the intestine to alter metabolic, reproduction, and longevity programs.

One canonical function of F-box proteins is to mediate poly-ubiquitination and proteasomal degradation of their substrates, which requires coordination with the Skp1 and Cullin proteins. While C. elegans expresses ∼520 F-box proteins, the genome only contains 21 Skp1-related (skr) genes and 6 Cullin (cul) genes, suggesting that these factors have less functional redundancy than the F-box family (Nayak et al., 2002; Thomas, 2006). To determine if FBXL-5 functions in concert with the SKR and CUL proteins, we performed a small-scale RNAi screen to identify the skr/cul genes that are required for reducing Pvit-3::GFP reporter expression in fbxl-5 over-expression animals. We screened all six cul genes, as well as a majority of the skr genes (14/21), for vitellogenesis phenotypes and excluded any clones that caused lethality or prevented larval development. Knock-down of skr-3 by RNAi modestly increased GFP levels in the reporter strain, which was recapitulated using the skr-3(ok365) mutant allele (Figure 6A; Supplementary Figure S6). Although we also identified cul-6 as a candidate in our RNAi screen, the cul-6(ok1614) mutation failed to replicate this finding; however, mutation of both skr-3 and cul-6 strongly increased Pvit-3::GFP expression in fbxl-5 over-expression animals relative to the control (Figures 6A, B; Supplementary Figure S6). Further mutation of skr-4 or skr-5, which both share significant similarity to skr-3 (Nayak et al., 2002), failed to dramatically enhance the skr-3(ok365); cul-6(ok365) double mutant. Together, these data suggest that FBXL-5 functions in concert with SKR-3 and CUL-6 to restrict vitellogenin production; however, it is likely that FBXL-5 acts with other redundantly-acting factors, as Pvit-3::GFP expression levels are only partially restored to wild-type levels in skr-3; cul-6 double mutant. Alternatively, high levels of FBXL-5 over-expression may facilitate interactions with other SKR/CUL proteins that are not normally preferred when FBXL-5 is expressed at endogenous levels, which may limit the suppressive effects of the skr-3 and cul-6 mutations in this experiment.

To further examine the role of poly-ubiquitination in the FBXL-5-dependent regulation of vitellogenesis, we introduced the uba-1(it129ts) temperature-sensitive mutation into the fbxl-5 over-expression strain. UBA-1 is the sole E1 ubiquitin-activating enzyme in C. elegans and its activity is required for the ubiquitin proteolytic pathway (Kulkarni and Smith, 2008). Consistent with our previous results, the uba-1(it129ts) mutation partially suppressed the effects of fbxl-5 over-expression at the restrictive, but not permissive, temperature (Figures 6C, D). Together, our results suggest that FBXL-5 acts in a SCF complex, likely with SKR-3 and CUL-6, to promote poly-ubiquitination-mediated proteasomal degradation. However, we are unable to eliminate the possibility that FBXL-5 associates with other SKR/CUL proteins in the intestine, or potentially a different set of SKR/CUL proteins in the muscle, to carry out some of its homeostatic functions.

We have previously shown that hypodermal lin-29 acts genetically upstream of intestinal mTORC2 signaling to regulate vitellogenesis (Dowen et al., 2016). We hypothesized that FBXL-5 might be responsible for modulating the signaling between LIN-29 and mTORC2, which we tested by performing genetic epistasis analyses. While knock-down of fbxl-5 by RNAi increased Pvit-3::GFP expression 3.3-fold in lin-4(e912) and 3.9-fold in lin-29(n333) mutant animals, the suppressive effects seen in rict-1(mg360) and sgk-1(ok538) animals were less striking (1.6- and 0.9-fold, respectively; Figure 7A; Supplementary Figure S7A). The mg360 allele is a missense mutation in rict-1, and thus, this mutant could retain some rict-1 function, which complicates the interpretation of our epistasis analysis. Therefore, we generated a complete rict-1 deletion mutant by CRISPR/Cas9 editing, removing the entire sequence between the start and stop codons, including the pqn-32 gene that is positioned between exons 10 and 11, and measured Pvit-3::GFP reporter expression. Consistent with the mg360 allele, complete deletion of rict-1 resulted in a robust decrease in Pvit-3::GFP levels, while further mutation of fbxl-5 resulted in a modest, but significant, 1.1-fold increase in reporter expression (Figure 7B). This weak suppression phenotype was not limited to vitellogenesis, as the fbxl-5(rhd43) mutation also partially suppressed the short lifespan of the rict-1(mg360) mutant (Figure 7C). To eliminate the possibility that FBXL-5 non-specifically restricts vitellogenin expression in the intestine, we performed fbxl-5 RNAi on daf-2(e1370) mutants, which also have reduced Pvit-3::GFP reporter expression (Dowen et al., 2016). Knock-down of fbxl-5 had no impact on vitellogenin expression in daf-2 mutant animals (Supplementary Figure S7B). Together, our results suggest that fbxl-5 acts genetically upstream of rict-1 and sgk-1 to regulate vitellogenesis; however, it is also likely that fbxl-5 acts simultaneously in a pathway parallel to rict-1 to regulate lifespan and vitellogenesis via a mTORC2-independent mechanism.

To further define the relationship between FBXL-5 and RICT-1 signaling, we performed mRNA-Seq on the rict-1(mg360) mutant and compared the differentially expressed genes (DEGs) between the rict-1 and fbxl-5 mutants (Supplementary Table S4). There was a striking overlap between these datasets, with 51% of the fbxl-5 DEGs also misexpressed in the rict-1 mutant (Figure 7D). The vast majority of these overlapping DEGs showed similar directionalities and degrees of mis-expression between the two mutants; however, a small cluster of genes was upregulated in the rict-1(mg360) mutant but downregulated in fbxl-5(rhd43) animals (Figure 7E). This could be a result of FBXL-5 having different roles within the intestine or having unique functions in different tissues. With many genes similarly altered in rict-1 and fbxl-5 mutants, we performed mRNA sequencing on the rict-1(mg360); fbxl-5(rhd43) double mutant and calculated the differential expression values for the 426 co-regulated genes. Simultaneous loss of both rict-1 and fbxl-5 did not further alter gene expression levels compared to the single mutants (Figure 7F), suggesting that these genes are regulated by FBXL-5 and RICT-1 in a linear pathway. Together, our results indicate that FBXL-5 regulates gene expression via mTORC2-dependent and independent mechanisms (Figure 7G).