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. 2018 Oct 4;210(4):1329–1337. doi: 10.1534/genetics.118.301450

Endoplasmic Reticulum Homeostasis Is Modulated by the Forkhead Transcription Factor FKH-9 During Infection of Caenorhabditis elegans

Erik J Tillman *, Claire E Richardson *,1, Douglas J Cattie *, Kirthi C Reddy *,2, Nicolas J Lehrbach †,‡, Rita Droste *,§, Gary Ruvkun †,‡, Dennis H Kim *,3
PMCID: PMC6283152  PMID: 30287474

Abstract

Animals have evolved critical mechanisms to maintain cellular and organismal proteostasis during development, disease, and exposure to environmental stressors. The Unfolded Protein Response (UPR) is a conserved pathway that senses and responds to the accumulation of misfolded proteins in the endoplasmic reticulum (ER) lumen. We have previously demonstrated that the IRE-1-XBP-1 branch of the UPR is required to maintain Caenorhabditis elegans ER homeostasis during larval development in the presence of pathogenic Pseudomonas aeruginosa. In this study, we identify loss-of-function mutations in four conserved transcriptional regulators that suppress the larval lethality of xbp-1 mutant animals caused by immune activation in response to infection by pathogenic bacteria: FKH-9, a forkhead family transcription factor; ARID-1, an ARID/Bright domain-containing transcription factor; HCF-1, a transcriptional regulator that associates with histone modifying enzymes; and SIN-3, a subunit of a histone deacetylase complex. Further characterization of FKH-9 suggests that loss of FKH-9 enhances resistance to the ER toxin tunicamycin and results in enhanced ER-associated degradation (ERAD). Increased ERAD activity of fkh-9 loss-of-function mutants is accompanied by a diminished capacity to degrade cytosolic proteasomal substrates and a corresponding increased sensitivity to the proteasomal inhibitor bortezomib. Our data underscore how the balance between ER and cytosolic proteostasis can be influenced by compensatory activation of ERAD during the physiological ER stress of infection and immune activation.

Keywords: endoplasmic reticulum stress, Unfolded Protein Response, proteasome, Caenorhabditis elegans, innate immunity, WormBase

RECENT studies in evolutionarily diverse organisms underscore the key role of endoplasmic reticulum (ER) stress responses during the activation of innate immunity (Martinon et al. 2010; Richardson et al. 2010). The IRE-1-XBP-1 branch of the Unfolded Protein Response (UPR) has an evolutionarily conserved role in activating the transcription of genes encoding ER chaperones, phospholipid biosynthetic enzymes, and ER-associated degradation (ERAD) machinery (Harding et al. 1999; Ng et al. 2000; Ye et al. 2000; Shen et al. 2005). In Caenorhabditis elegans, intestinal infection by pathogenic bacteria induces a secretory immune response, including putative antimicrobial effectors include lectins, lysozymes, and saponin-like peptides (Kato et al. 2002; Mallo et al. 2002; Schulenburg and Boehnisch 2008; Hoeckendorf et al. 2012), which represents a physiological source of ER stress (Richardson et al. 2010). C. elegans mutants lacking XBP-1 exhibit larval lethality in the presence of pathogenic Pseudomonas aeruginosa that can be alleviated by suppressing activation of the innate immune response (Richardson et al. 2010).

We have been interested in identifying the mechanisms activated in response to the physiological ER stress of infection and immune activation. To identify molecular pathways functioning in concert with the UPR to maintain ER homeostasis during infection, we have conducted a genetic screen for suppressors of xbp-1 mutant larval lethality on P. aeruginosa. We recently reported that mutations in the conserved but nonessential eukaryotic translation initiation factor subunits EIF-3.K and EIF-3.L suppress xbp-1 larval lethality on P. aeruginosa, enhance resistance to ER stress, and confer a marked extension in lifespan (Cattie et al. 2016). Here, we report the identification of loss-of-function mutations in four genes encoding conserved transcriptional regulators, FKH-9, ARID-1, HCF-1, and SIN-3, which suppress the larval lethality of xbp-1 mutant animals in the presence of P. aeruginosa. Characterization of FKH-9 suggests that mutations in fkh-9 result in increased activity of ERAD, which serves to promote ER homeostasis, while diminishing the capacity of the proteasome to respond to cytosolic stress. Our results reveal how compensatory changes in ERAD activity may alter the balance between mechanisms involved in the maintenance of proteostasis in the ER and cytosol of C. elegans during the physiological ER stress of pathogen infection.

Materials and Methods

C. elegans strains

C. elegans strains were maintained as previously described (Brenner 1974). The strains used in this study are listed in Supplemental Material, File S1.

Suppressor screen and mutant identification

Ethyl methanesulfonate was used to mutagenize the starting strain containing both the xbp-1(tm2482) allele and a transgenic reporter responsive to activation of the immune effector PMK-1, agIs219[pT24B8.5::gfp]. F2 animals were scored for their ability to survive through larval development on Slow Kill Assay (SKA) plates seeded with P. aeruginosa strain PA14. We screened ∼10,000 haploid genomes. Nonsterile mutants were retested for suppression of xbp-1 larval lethality on P. aeruginosa and were checked for fluorescence of the agIs219 reporter.

Isolated mutants were backcrossed at least two times. Genomic DNA from each backcrossed mutant was purified using the PureGene Kit (QIAGEN, Valencia, CA), and libraries were prepared using the NEBNext Ultra Library Prep Kit (New England Biolabs, Beverly, MA) and submitted for paired-end whole genome sequencing on an Illumina NextSeq. Reads were mapped using BWA and mutations were identified relative to the reference N2 genome (WS245; www.wormbase.org). After identifying mutations unique to each mutant, we used existing deletion or null alleles to confirm that mutations in candidate genes suppresses larval lethality of the xbp-1 mutant on P. aeruginosa as described below.

Generation of endogenously-tagged fkh-9::gfp allele by CRISPR/Cas9 genome editing

CRISPR/Cas9 genome editing was used to insert a 2xTY1::GFP tag at the 3′ end of the fkh-9 locus immediately upstream of the stop codon, as previously described (Cattie et al. 2016). The fkh-9::2xTY1::gfp homologous repair template was generated by inserting a 2xTY1::GFP tag (amplified from clone CBGtg9050D0789D; TransgeneOme Project) in-frame into a plasmid containing fkh-9 homology arms (1.2 kb upstream and 1 kb downstream of the stop codon) (Sarov et al. 2012). Two guide RNAs (gRNAs) were used to increase probability of target cleavage, targeting the sequences atcgacgcatagaaaaaggttgg (<50 bp from stop codon) and cagtcttttcttcttcaatagg (<5 bp from stop codon). gRNAs were constructed using pRB1017 as the backbone (Arribere et al. 2014). The homologous repair template was designed to disrupt both gRNA target sequences to prevent recleavage of successfully recombined DNA.

The injection mix consisted of Peft-3::cas9, injected at 50 ng/μl; gRNA plasmids, injected at 25 ng/μl each; homologous repair template, injected at 50 ng/μl; and pCFJ90, injected at 1.5 ng/μl.

Constructs and generation of transgenic animals

Tissue-specific expression constructs were generated as previously described (Cattie et al. 2016). fkh-9a complementary DNA (cDNA) was amplified from N2 cDNA by PCR. The wild-type Prab-3 and Pges-1 promoters were amplified from N2 genomic DNA by PCR and used to generate plasmids driving tissue-specific expression of fkh-9. Fosmid clone WRM0616dC12 covers the fkh-9 genomic locus including 15 kb upstream and 10 kb downstream of the coding sequence (Source BioScience). Prab-3::fkh-9a construct was injected at 10 ng/μl, Pges-1::fkh-9a was injected at 25 ng/μl, and fosmid was injected at 50 ng/μl. A plasmid containing Punc-122::gfp was injected as a co-injection marker at 50 ng/μl. At least three independent transgenic lines were evaluated for each construct.

P. aeruginosa development assay

Overnight LB cultures of either P. aeruginosa strain PA14 or Escherichia coli strain OP50 were seeded to Slow Kill Assay (SKA) plates and incubated at 37° overnight, and then at room temperature for 24 hr (Tan et al. 1999). Gravid C. elegans adults were transferred to the experimental plates and allowed to lay eggs for 2–4 hr. Fraction of embryos reaching at least the L4 stage was scored after 72 hr of growth at 25°. Plates contained 50–100 embryos and three plates were averaged within each experiment.

Tunicamycin development assay

Tunicamycin assay was performed as described (Cattie et al. 2016). Plates contained 50–150 embryos and three plates were averaged within each experiment. Representative experiments of at least three independent experiments are shown.

Transmission electron microscopy

L1 larvae synchronized by hypochlorite treatment were grown on E. coli strain OP50 or P. aeruginosa strain PA14 at 25° until the L3 stage. Animals were fixed in 0.7% glutaraldehyde, 0.7% OsO4 0.1 M cacodylate buffer for 1 hr on ice. Samples were cut in the anterior region, washed in 0.1 M cacodylate buffer, and postfixed in 2% OsO4 in 0.1 M cacodylate buffer overnight at 4°. After washing samples in 0.1 M cacodylate buffer on ice, three to five worms were mounted onto agar blocks and samples were then stained en bloc in 2% uranyl acetate in 50% ethanol for 1 hr, dehydrated in a series of alcohols, and embedded in Epon resin. Thin sections of 50–70 nm were obtained on an Ultracut E. Samples were observed on a JEOL 1200-EX electron microscope at 80 kV and imaged with a side-mounted AMT XR-41 CCD camera (Hall et al. 2012). Images were acquired at ×60,000 magnification.

Bortezomib development assay

Bortezomib assay was performed as described (Lehrbach and Ruvkun 2016). A minimum of 20 larvae were deposited in each well and allowed to develop in the presence of the indicated concentration of bortezomib and 5× concentrated E. coli OP50 in S-basal supplemented with carbenicillin (25 μg/ml). DMSO concentration was constant across all wells (0.05%). For each genotype, a total of six wells were scored at each concentration, and the average of the qualitative score is rendered in the heatmap.

RNA interference treatment

RNA interference (RNAi) clones were selected from the Ahringer library (Kamath et al. 2003) or the ORFeome library (Rual et al. 2004). For genes not targeted by clones in either library, RNAi constructs were made by cloning a 1–2 kb genomic amplicon into the plasmid L4440 and transforming into E. coli HT115. All RNAi clones were validated by Sanger sequencing. RNAi was grown overnight in LB broth supplemented with carbenicillin (25 μg/ml) and seeded to NGM plates supplemented with carbenicillin (25 μg/ml) and isopropyl β-D-1 thiogalactopyranoside (2 mM). L4 animals of the indicated strain were transferred to RNAi plates. After 3 days of incubation at 20°, L4 larvae (F1 progeny) were mounted and anesthetized in sodium azide (50 mM). Images were acquired with an Axioimager Z1 microscope. Quantification was performed using ImageJ software, averaging intensity across the intestine.

Quantification of ERAD function

The transgene vkEx1879 encodes a terminally misfolded procathepsin mutant fused to YFP that is exported from the ER and degraded under normal physiological conditions (Miedel et al. 2012). Upon ERAD disruption by sel-1 RNAi, the terminally misfolded procathepsin mutant accumulates and increases intestinal fluorescence.

Quantification of proteasome function

The transgene mgIs77 contains Prpl-28::Ub(G76V)::GFP, which encodes an uncleavable ubiquitin moiety fused to GFP, such that GFP is constitutively targeted for proteasomal degradation.

Data and reagent availability statement

Strains are available upon request. File S1 contains information about the strains used in this study. The authors affirm that all other data necessary for confirming the conclusions of the article are present within the article, figures, and tables. Supplemental material available at Figshare: https://doi.org/10.25386/genetics.6900653.

Results and Discussion

We previously determined that xbp-1 is required for larval development of C. elegans in the presence of pathogenic P. aeruginosa (Richardson et al. 2010). Although wild-type animals eventually succumb to infection by P. aeruginosa as adults, they are able to complete larval development by utilizing P. aeruginosa as a food source and mounting a protective immune response. We have shown that xbp-1 mutant animals exhibit larval lethality because the activation of innate immunity leads to ER dysfunction and disruption of ER morphology (Richardson et al. 2010).To identify mechanisms that compensate for XBP-1 deficiency during physiological ER stress caused by immune activation, we conducted a genetic screen to identify mutations that can suppress xbp-1 larval lethality in the presence of pathogenic P. aeruginosa. The starting strain carried a GFP transgene under the control of the T24B8.5 promoter that serves as a reporter of PMK-1 function, enabling us to select for mutants that suppressed the larval lethality of xbp-1 without affecting levels of PMK-1–mediated innate immune activation (Figure 1A).

Figure 1.

Figure 1

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Identification of mutations that suppress larval lethality of xbp-1 mutant animals on P. aeruginosa. (A) Mutations in four genes were identified in a forward genetic screen in xbp-1(tm2482) animals carrying the integrated transgene agIs219, which responds to PMK-1–dependent immune activation. F2 animals that completed larval development and retained intestinal fluorescence on P. aeruginosa were retested and backcrossed more than two times before whole genome sequencing to identify mutations that suppress xbp-1 mutant larval lethality on P. aeruginosa. (B) Representative images of animals 72 hr after egg laying, on P. aeruginosa. Images were taken at ×12 magnification. (C) Development assay monitoring survival on nonpathogenic E. coli or pathogenic P. aeruginosa; 50–100 eggs were laid per plate and the fraction reaching at least the L4 larval stage was scored after 72 hr at 25°. Genetic identity of mutant alleles was confirmed with additional null mutants. (D) Intestinal ER imaged by transmission electron microscopy; ×60,000 magnification. Animals were grown on P. aeruginosa PA14 until reaching the L3 stage, then fixed and imaged. Loss of fkh-9 and arid-1 suppress ER morphological defects seen in the xbp-1 mutant, including loss of cisternal structure and depletion of rough ER. (E) Multiple loss-of-function alleles of arid-1 and fkh-9 suppress xbp-1 mutant larval sensitivity to elevated temperatures on nonpathogenic E. coli OP50. Each strain was assayed in triplicate, and data are representative of two independent experiments. Significance in C and E was assessed by one-way ANOVA followed by Dunnett’s multiple comparisons test. Error bars indicate SD of three plates. * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001, compared to xbp-1(tm2482).

We identified mutations in four genes not previously identified as regulators of ER homeostasis (Table 1): arid-1, an ARID/BRIGHT domain-containing transcription factor; fkh-9, a Forkhead/winged-helix domain-containing transcription factor (Hope et al. 2003); sin-3, a histone deacetylase complex (HDAC) subunit (Choy et al. 2007); and hcf-1, a transcriptional regulator previously shown to associate with histone deacetylases (Wysocka et al. 2003; Li et al. 2008; Rizki et al. 2012). Deletion mutant alleles of each of these genes suppresses the larval lethality of xbp-1 mutants exposed to P. aeruginosa, confirming that the mutations identified in the screen are loss-of-function alleles in these genes (Figure 1, B and C).

Table 1. Molecular identities of mutant alleles identified in genetic screen for suppressors of xbp-1 larval lethality on P. aeruginosa.

Gene Allele Mutation Function
fkh-9 qd197 G111E Forkhead family transcription factor
hcf-1 qd198 G110E HDAC-associated transcriptional corepressor
sin-3 qd199 Q212X HDAC subunit, transcriptional corepressor
arid-1 qd200 Exon 2_3 splice acceptor ARID/Bright domain-containing transcription factor
qd215 Exon 6_7 splice donor
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HDAC, histone deacetylase complex.

We have previously reported that larval exposure of wild-type C. elegans to P. aeruginosa does not perturb intestinal ER architecture, but transmission electron microscopy of xbp-1 mutant animals reveals loss of rough ER and a decrease in visible cisternae in the presence of P. aeruginosa (Richardson et al. 2010; Cattie et al. 2016). We observed a striking suppression of these xbp-1 mutant-specific ER structural changes upon loss of fkh-9 or arid-1 on P. aeruginosa (Figure 1D), consistent with a preservation of ER homeostasis during immune activation even in the absence of XBP-1. As we have previously reported that xbp-1 mutants are sensitive to larval development at 27° (Richardson et al. 2011), we asked whether mutations in fkh-9 and arid-1 also suppress this stress phenotype. Whereas xbp-1 single mutant animals die or arrest as early larvae when grown on E. coli at 27°, xbp-1;fkh-9 and xbp-1;arid-1 double mutant animals survive through larval development, reaching fertile adulthood after 72 hr (Figure 1E). These results demonstrate two additional ER stress resistance phenotypes, namely exposure to tunicamycin and xbp-1 mutant exposure to elevated temperature.

We elected to focus on the further characterization of FKH-9, motivated in part by recent studies establishing that FKH-9 is a critical regulator of insulin signaling that promotes neuronal regeneration (Kaletsky et al. 2015). The restoration of pathogen-exposed xbp-1-mutant intestinal ER architecture observed upon mutations in fkh-9 led us to hypothesize that the main site of action for FKH-9 function is the intestine. To accurately examine fkh-9 expression patterns, we generated an endogenously GFP-tagged fkh-9 allele (allele designation fkh-9(qd327[fkh-9::2xTY1::gfp]) using CRISPR/Cas9-mediated genome editing (Tzur et al. 2013; Arribere et al. 2014; Ward 2015), taking advantage of all endogenous regulatory elements and without pleiotropies due to transgenic overexpression. We observed FKH-9 expression in both neurons and intestinal nuclei (Figure 2, A and B), consistent with published reports (Hope et al. 2003; Kaletsky et al. 2015). We also confirmed that our endogenously-tagged allele of fkh-9 encodes a functional protein, as this allele fails to suppress xbp-1 mutant larval lethality on P. aeruginosa (Figure 2C).

Figure 2.

Figure 2

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FKH-9 is expressed in both neuronal and intestinal tissues. (A) Confocal microscopy image of the head of a fkh-9(qd327) L2 larva, germline-edited with gfp inserted at the 3′ end of the endogenous fkh-9 locus. Open arrowheads point to unidentified neurons expressing FKH-9::GFP. Bar, 50 μm. (B) Confocal microscopy image of the intestine of a fkh-9(qd327) L2 larva. Solid arrowheads point to intestinal nuclei with FKH-9::GFP expression. Bar, 50 μm. (C) fkh-9(qd327) does not confer xbp-1-mutant larval resistance to P. aeruginosa, suggesting it encodes a functional protein. (D) Fosmid rescue, intestinal re-expression (using the Pges-1 promoter), and neuronal re-expression (using the Prab-3 promoter) of FKH-9 are all sufficient to sensitize xbp-1 mutant animals to P. aeruginosa PA14 exposure during larval development. Error bars indicate SD of three plates in C–E. (E) Overexpression of FKH-9 from an extrachromosomal array carrying either fosmid, plasmid encoding intestine-specific expression (Pges-1::fkh-9), or plasmid encoding neuron-specific expression (Prab-3::fkh-9) is not sufficient to perturb ER homeostasis of xbp-1 mutant animals on E. coli OP50. Significance in C and D was assessed by one-way ANOVA followed by Dunnett’s multiple comparisons test. *** P < 0.001, **** P < 0.0001, n.s., not significant (P > 0.05) compared to xbp-1(tm2482) (C) and xbp-1(tm2482);fkh-9(ok1709) (D).

To directly test the role of FKH-9 activity in different tissues in the suppression of xbp-1 loss of function, we generated independent transgenic lines expressing either a fosmid covering the fkh-9 genomic locus (WRM0616dC12), or fkh-9 cDNA expressed under the Pges-1 promoter for intestinal specificity or the Prab-3 promoter for neuronal specificity (Nonet et al. 1997; Marshall and McGhee 2001). Extrachromosomal fosmid rescue and intestinal re-expression, as well as neuronal re-expression of fkh-9 in the xbp-1;fkh-9 double mutant background were all sufficient to suppress organism-wide loss of fkh-9 (Figure 2D), consistent with some basal function of FKH-9 in both neuronal and intestinal tissue leading to alterations in ER homeostasis and organismal physiology specifically upon immune activation. We had some concern that the results of our transgenic rescue experiments might be influenced by transgene toxicity arising from overexpression of FKH-9, but we found that none of these constructs affected the development of xbp-1;fkh-9 double mutant animals on nonpathogenic E. coli (Figure 2E).

We hypothesized that the identified mutations may directly enhance ER homeostasis, thereby promoting survival upon innate immune activation in the xbp-1 mutant background. To test this hypothesis, we exposed the mutants to the compound tunicamycin, an inhibitor of N-linked glycosylation that causes the accumulation of misfolded proteins in the ER lumen and activates the UPR. Mutations in arid-1 and fkh-9 strongly enhanced tunicamycin resistance (Figure 3A), consistent with generally enhanced ER stress resistance.

Figure 3.

Figure 3

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Mutations in fkh-9 promote ER homeostasis and enhance ERAD function. (A) Larval development assay on NGM plates containing 2.5 μg/ml tunicamycin, seeded with E. coli OP50. At 72 hr after egg-lay, animals were scored for development to at least the L4 larval stage. Loss of arid-1 or fkh-9 enhances ER stress resistance in a UPR-independent manner. Each strain was assayed in triplicate and data are representative of three independent experiments, with significance assessed by one-way ANOVA corrected by Dunnett’s multiple comparison test. ** P < 0.01, *** P < 0.001. (B) vkEx1879[Pnhx-1::cpl-1W32A Y35A::YFP + Pmyo-3::mCherry] was used to visualize intestinal ERAD activity. (C) Quantification of fluorescence micrographs in B, with significance assessed by one-way ANOVA followed by Sidak post hoc correction. **** P < 0.0001, n.s., not significant (P > 0.05). Error bars indicate SD of three plates in A and C.

To further investigate mechanisms of enhanced ER homeostasis conferred by fkh-9 mutations, we conducted an assay for ERAD activity. ERAD is a regulated program responsible for recognizing, retrotranslocating, and targeting to the proteasome misfolded ER luminal and membrane proteins (Friedlander et al. 2000). Using a reporter encoding a terminally misfolded procathepsin L mutant fused to YFP, we monitored ERAD function upon knockdown of sel-1, which encodes a component of the HRD1/HRD3 ERAD complex (Plemper et al. 1999; Miedel et al. 2012). Whereas we observed marked accumulation of the ERAD substrate in wild-type animals upon sel-1 RNAi, we observed attenuation in fluorescence accumulation upon sel-1 RNAi in the fkh-9 mutant, consistent with improved retrotranslocation and cytoplasmic degradation of the misfolded ER cargo (Figure 3, B and C).

We considered that enhanced ER homeostasis could represent a generalized increased stress resistance and proteostasis across cellular compartments in particular, conferred by mutations in fkh-9, and thus we sought to determine whether mutations in fkh-9 might also confer enhanced proteostasis in other subcellular compartments. We exposed fkh-9 and arid-1 mutant animals to increasing concentrations of the proteasome inhibitor bortezomib (Lehrbach and Ruvkun 2016). Interestingly, we found that mutations in fkh-9 and arid-1 sensitized animals to proteasome inhibition, with larval lethality and developmental delay seen at lower concentrations of bortezomib in the mutants than in wild-type animals (Figure 4A). Next, we asked whether proteasome function in the cytosol was impaired in fkh-9 mutant animals. We generated a reporter strain carrying a transgene consisting of GFP fused to a well-characterized ubiquitin mutant that cannot be cleaved from GFP (UbG76V::GFP, hereafter referred to as UbV::GFP). Under normal proteasomal function, this ubiquitinated GFP is degraded by the proteasome. However, RNAi depletion of hecd-1, which encodes an E3 ubiquitin ligase required for degradation of the ubiquitin fusion protein causes accumulation of UbV::GFP (Segref et al. 2011) (Figure 4B, top). While fkh-9 loss itself does not affect UbV::GFP degradation, fkh-9 loss caused increased UbV::GFP accumulation upon hecd-1 RNAi, consistent with mild proteasome dysfunction (Figure 4, B and C).

Figure 4.

Figure 4

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FKH-9 loss perturbs proteasome function and sensitizes animals to proteasomal inhibition. (A) Loss of fkh-9 or arid-1 sensitizes C. elegans to proteasome inhibition during larval development. A minimum of 20 synchronized L1 larvae were dropped into indicated concentrations of bortezomib in concentrated E. coli OP50, and each well was scored qualitatively after 4 days at 20°. Each cell of the heatmap is an average of six wells representative of two to three independent experiments. Images at right are representative wells corresponding to each qualitative score. (B) L4 larvae carrying a transgene expressing UbV::GFP were grown for a full generation on indicated RNAi bacteria and imaged to assay proteasome function. GFP with an uncleavable Ub tag is constitutively degraded by the proteasome, and accumulates upon knockdown of hecd-1. Representative images are shown in B, and ≥12 animals per condition are quantified in C, with significance assessed by one-way ANOVA followed by Sidak post hoc correction. **** P < 0.0001, n.s., not significant (P > 0.05).

Taken together, our data suggest that fkh-9 loss enhances the removal of misfolded proteins from the ER lumen at the expense of proteasomal degradation in the cytosol. We speculate that the role of FKH-9 in balancing cellular proteostasis may also contribute to its role enhancing axonal regeneration in adult neurons (Kaletsky et al. 2015). Ongoing characterization aims to determine how FKH-9 may function in relation to other genes identified in our study, as well as previously described roles for EIF-3.K and EIF-3.L (Cattie et al. 2016).

SIN-3 and HCF-1 are broadly conserved subunits of histone deacetylate complexes, and have been implicated in the modulation of cellular stress responses and longevity (Wysocka et al. 2003; Schröder et al. 2004; Li et al. 2008; Rizki et al. 2012). arid-1 is an ortholog of the human genes ARID4A/B, which encode transcription factors that have been shown to interact with pRB (Defeo-Jones et al. 1991; Fattaey et al. 1993; Patsialou et al. 2005), leading to gene silencing through histone deacetylase complex recruitment and activity (Lai et al. 1999). In mice, Rb and Rbp1 have been shown to interact with mSin3A, suggesting a link between ARID4A/B and SIN3-dependent corepression of target transcription (Fleischer et al. 2003). Further investigation will uncover whether ARID-1 functions in concert with SIN-3 and HCF-1 in regulating transcription in C. elegans stress responses.

Our demonstration of the balance between ER and cytosolic proteostasis mediated by FKH-9 activity raises interesting possibilities for the reported phenotypes of fkh-9 loss both here and elsewhere (Kaletsky et al. 2015). For example, FKH-9 may modulate the balance of protein degradation across the cytosol and ER through its direct modulation of ERAD or proteasome function. fkh-9 loss could then have opposing effects on cytosolic and ER proteostasis, as we have observed, and this model is consistent with the importance of cytosolic stress-response pathways in neuronal homeostasis and axon regeneration (Silva et al. 2013; Alam et al. 2016). Although we observe an organismal benefit upon loss of fkh-9 function, Kaletsky et al. (2015) observe that FKH-9 is a critical regulator of insulin signaling that promotes neuronal regeneration. These apparently contrasting effects of fkh-9 loss may be explained via opposing roles of FKH-9 across cellular compartments. Further characterization of protein folding homeostasis in the ER and the cytosol is necessary to completely understand FKH-9 activity during pathogen infection of C. elegans. Importantly, our data corroborate prior studies pointing to a homeostatic interrelationship among proteostasis networks across subcellular compartments (Liu and Chang 2008; Pereira et al. 2014; Hourihan et al. 2016).

Acknowledgments

We thank H.R. Horvitz and the Caenorhabditis Genome Center (which is funded by National Institutes of Health Office of Research Infrastructure Programs grant P40OD010440) for strains and reagents. We thank members of the Kim laboratory for helpful conversation and feedback on the manuscript and figures. This work was supported by National Institutes of Health grant R01GM084477 (to D.H.K.), the National Institutes of Health predoctoral training grant T32GM007287 (to E.J.T.), and the Massachusetts Institute of Technology Office of Graduate Education Hugh Hampton Young Memorial Fellowship (to E.J.T.).

Footnotes

Supplemental material available at Figshare: https://doi.org/10.25386/genetics.6900653.

Communicating editor: B. Goldstein

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