We recently reported improved ribosome-footprint profiles (Ingolia et al., 2009) and mRNA-abundance measurements from exponentially growing S. cerevisiae (Weinberg et al., 2016). Under these growth conditions HAC1 mRNA is almost entirely unspliced (Figure 1—figure supplement 1A) and is distributed across a sucrose gradient with ~50% in the non-translating fractions and the remaining ~50% extending across all of the translating (i.e., 80S and larger) fractions without substantial enrichment in any particular fraction (Figure 1A), similar to previous observations (Arava et al., 2003; Chapman and Walter, 1997; Cox and Walter, 1996; Kuhn et al., 2001; Mori et al., 2010; Park et al., 2011; Payne et al., 2008; Rüegsegger et al., 2001; Sathe et al., 2015). In contrast, the well-translated actin (ACT1) mRNA is essentially absent from the non-translating fractions and is found mostly in large polysomes. Based on the sedimentation of HAC1 mRNPs, we predicted that the mRNA would generate a large number of ribosome-protected fragments, in a quantity only ~2-fold fewer than similarly abundant mRNAs (based on the fraction of HAC1 mRNA in the untranslated fractions). Strikingly, however, after normalizing for mRNA abundance HAC1 generates the fewest ribosome-protected fragments among all expressed yeast genes (Figure 1B)—and ~50-fold fewer than expected from the polysome profile. Rather than providing evidence for stalled ribosomes on HAC1^u mRNA, instead these observations suggest that either ribosomes are stalled on the mRNA in a closely packed configuration that prevents nuclease cleavage between ribosomes, which would eliminate the ~28-nucleotide fragments that are sequenced in the ribosome-profiling method (Figure 1C, middle); or that there are not stalled ribosomes on HAC1^u mRNA (Figure 1C, bottom).

Figure 1 with 1 supplement see all

Download asset Open asset

Although the polysome-like sedimentation of HAC1^u mRNA indicates ribosome association, it does not reveal if the ribosomes associated with the mRNA were ever engaged in its translation. Alternatively, the associated ribosomes might be bound in a conformation that is unrelated to translation of HAC1^u mRNA. We therefore devised an experiment to definitively determine whether the ribosomes bound to HAC1^u mRNA reflect ribosomes that were engaged in its translation. To do so, we took advantage of the observation that HAC1 mRNA is distributed across all of the translating fractions of a sucrose gradient (Figure 1A). If the deep sedimentation is due to multiple translating ribosomes being stalled on a single mRNA, then reducing the number of translating ribosomes that can fit on the mRNA should shift the sedimentation pattern toward lighter fractions. We designed a series of constructs containing point mutations in the first exon of HAC1 that created premature termination codons, which reduce the size of the ORF and thereby limit the number of translating ribosomes (Figure 1D, top). To ensure that the mutant alleles were expressed at near wild-type levels, we replaced the endogenous HAC1 allele without disrupting flanking regulatory regions. Remarkably, each of the mutant mRNAs had a sedimentation pattern that was indistinguishable from the wild-type mRNA (Figure 1D, bottom). In the most extreme case, the mRNA containing a 21-nucleotide ORF that can only accommodate a single translating ribosome still co-sedimented with polysomes containing upwards of 10 ribosomes (fraction 14 of the gradient). These data provide direct evidence that the polysome-like sedimentation of HAC1^u mRNA is not due to stalled ribosomes on the mRNA.

We hypothesized that the ribosome association of HAC1^u mRNA was instead due to non-specific interactions between the mRNA and bona fide polysomes formed on other mRNAs. To evaluate the extent of such non-specific interactions, we introduced an exogenous control RNA that should not be translated: an uncapped luciferase-encoding RNA purified from an in vitro transcription reaction. We analyzed the sedimentation behavior of this control RNA when it was added to lysis buffer compared to when it was added to the yeast lysate prior to centrifugation. Surprisingly, some of the exogenous RNA was found in the translating fractions of the lysate—a behavior not observed in lysis buffer alone (Figure 1E, middle). Through extensive optimization, we found that the addition of heparin to the lysis buffer (at a concentration of 0.2 mg/ml) was sufficient to largely prevent the deep sedimentation of exogenous RNA. The addition of heparin also had a major effect on the sedimentation of HAC1 mRNA, as most (83%) now sedimented in the non-translating fractions of the gradient (Figure 1E, bottom). In contrast, the sedimentation of ACT1 mRNA (Figure 1E, bottom) and the overall polysome profile (Figure 1E, top) were largely unchanged, suggesting that heparin competed away non-specific interactions without disrupting bona fide polysomes.

Based on these results, we re-analyzed the HAC1 mutants with shortened ORFs using heparin-containing lysis buffer. Under these conditions, the polysome co-sedimentation of each of the mRNAs was greatly reduced (from ~50% to < 20%) but there was still no difference among the constructs (Figure 1F), providing further evidence against stalled ribosomes on HAC1^u mRNA. Importantly, when we treated cells with the reducing agent dithiothreitol (DTT) to induce the UPR and concomitant splicing of HAC1 mRNA (Figure 1—figure supplement 1C) and repeated the experiment, the variant mRNAs now displayed the expected differential sedimentation based on ORF length (Figure 1G), validating our experimental design. Together, these results demonstrate that at steady state HAC1^u mRNA is not associated with either actively elongating or stalled ribosomes. This indicates that the primary block to production of Hac1^up is at the stage of translation initiation, not translation elongation as previously proposed (Chapman and Walter, 1997; Richter and Coller, 2015; Rüegsegger et al., 2001).

The absence of ribosomes on HAC1^u mRNA suggests that translation initiation is inhibited. To further understand the mechanism of this inhibition, we used a green fluorescent protein (GFP) reporter system that has been previously shown to recapitulate post-transcriptional silencing by the HAC1 intron (Chapman and Walter, 1997; Rüegsegger et al., 2001). We replaced the first exon of HAC1 with the GFP ORF lacking its own stop codon (Figure 2B), which allowed us to quantitatively analyze post-transcriptional silencing of GFP using a combination of flow cytometry (protein abundance), quantitative RT-PCR (mRNA abundance), and sucrose gradient fractionation (ribosome density). When GFP was embedded in an otherwise wild-type HAC1 context the mRNA sedimented almost entirely in the non-translating fractions (Figure 2A), and there was no detectable fluorescence above background (Figure 2C). In contrast, cells expressing a reporter construct missing the entire intron displayed a strong fluorescence signal (Figure 2C), and most of the mRNA was found in the translating fractions (Figure 2A). Thus, the reporter GFP mRNA behaves similarly to endogenous HAC1 mRNA.

Figure 2 with 1 supplement see all

Download asset Open asset

To determine whether 5′-UTR–intron base pairing is required to prevent ribosome loading on the reporter mRNA, we designed constructs in which the sequence implicated in base-pairing interactions in either the 5′-UTR or the intron was mutated to its complement to disrupt the interaction (Figure 2B, bottom), as was done previously (Rüegsegger et al., 2001). Both mutant mRNAs sedimented mostly in the translating fractions in a manner that was similar to the intronless construct (Figure 2A). When we combined the 5′-UTR and intron mutations and thereby restored base pairing, the mRNA was now found almost exclusively in the non-translating fractions and resembled the original reporter mRNA. These results demonstrate that base pairing between the 5′-UTR and intron is required to prevent ribosome loading by directly impeding the binding or progress of the scanning ribosome.

Given that the base-pairing mutant mRNAs were loaded with ribosomes similarly to the intronless mRNA (Figure 2A), we expected to observe GFP expression from the mutant mRNAs that was similar to that of the intronless construct. Remarkably, however, neither of the strains expressing a base-pairing mutant mRNA had any GFP detectable by either flow cytometry (Figure 2C) or immunoblotting (Figure 2—figure supplement 1). The lack of GFP signal despite polysome sedimentation was not due to low mRNA abundance, as the mutant mRNAs were present at similar levels compared to the intronless mRNA (Figure 2—figure supplement 1A, compare construct 3 with constructs 5–6). These results suggest that an additional silencing mechanism acting downstream of translation initiation prevents GFP accumulation when base pairing is disrupted.

In addition to removing the intron portion of the base-pairing interaction, splicing of HAC1 mRNA also alters the C-terminal tail of the encoded protein: The ORF in HAC1^u mRNA has a 10-amino-acid tail encoded by the intron, which in HAC1ⁱ mRNA is replaced by an 18-amino-acid tail encoded by the second exon (Figure 3A). The fact that the polypeptide tails encoded by the HAC1^u and HAC1ⁱ mRNAs are different suggests that the 10-amino-acid tail unique to Hac1^up may be functionally important. We therefore hypothesized that the intron-encoded C-terminal tail may be involved in the additional silencing mechanism revealed by our base-pairing mutant reporters (Figure 2).

Figure 3 with 2 supplements see all

Download asset Open asset

To investigate this possibility, we designed a reporter construct in which we removed the entire intron except for the 5′ end that codes for the 10-amino-acid tail of Hac1^up (Figure 3B, fourth construct). Remarkably, cells expressing this construct had no detectable GFP (Figure 3C), despite the corresponding mRNA being abundant (Figure 3—figure supplement 1A) and detected on polysomes due to the absence of the base-pairing interaction (Figure 3D). Thus, the coding region at the 5′ end of the intron is sufficient for complete silencing of the GFP reporter. Introducing a termination codon between GFP and the intron restored robust fluorescence comparable to that observed for the intronless construct, which implicated translation of the 5′ end of the intron as required for silencing. Furthermore, making 10 nucleotide changes that maintained the coding potential of the intron-encoded tail (Figure 3B, bottom) had no effect on silencing (Figure 3C), suggesting that the amino-acid sequence encoded by the 5′ end of the intron is more important than the nucleotide sequence itself.

To determine whether the 10-amino-acid element alone was sufficient for silencing in the absence of any other HAC1 sequences, we expressed GFP from a different locus in the yeast genome (TMA7) either with or without the 10-amino-acid tail. When the 10-amino-acid sequence was either absent or not translated due to a premature stop codon, we observed robust GFP signal (Figure 3C). In contrast, there was no fluorescence detected in cells expressing GFP containing the 10-amino-acid tail. Thus, the C-terminal tail of Hac1^up is sufficient for silencing independently of the rest of the HAC1 intron and any other HAC1 sequences.

Having established the effects of the 10-amino-acid tail in isolation, we next examined the effects of the tail when translation initiation was inhibited by the base-pairing interaction. In an otherwise wild-type HAC1 context, preventing translation of the 10-amino-acid tail either by deleting the entire sequence or by introducing a stop codon caused GFP to accumulate to low but detectable levels (Figure 3C) without greatly affecting the sedimentation of the corresponding mRNAs (Figure 3D). Thus, even when base pairing is intact there is some low-level accumulation of GFP that is normally suppressed by the 10-amino-acid silencing element.

In the context of reporter constructs lacking the intron-encoded C-terminal tail, mutations in either the 5′-UTR or intron that disrupted base pairing greatly increased the amount of GFP, while the compensatory double-mutant construct with restored base pairing had only low levels of GFP (Figure 3C and Figure 3—figure supplement 1B). These results are in agreement with previous GFP reporter experiments, which used constructs that contained a stop codon between the GFP and intron sequences and therefore had inadvertently eliminated the effects of the 10-amino-acid tail (Chapman and Walter, 1997; Rüegsegger et al., 2001).

Together, our GFP reporter experiments indicate that the HAC1 intron mediates post-transcriptional silencing through a pair of independent but partially redundant mechanisms: base-pairing interactions with the 5′-UTR that inhibit translation initiation, and a novel silencing mechanism mediated by the intron-encoded C-terminal tail of Hac1^up. Robust expression requires that both silencing mechanisms be inactivated, as would happen simultaneously when the intron is removed by Ire1p-dependent splicing.

What is the mechanism by which the intron-encoded tail of Hac1^up silences gene expression? Because disrupting translation of the tail elevated protein levels (Figure 3C) without affecting polysome formation (Figure 3D), we inferred that the tail was exerting its effect downstream of translation initiation. We therefore reasoned that the 10-amino-acid sequence was acting either by halting translation across the entire mRNA (after polysome formation); or by promoting protein degradation. Our inability to detect GFP containing the 10-amino-acid tail (Figure 3C) prevented us from directly comparing the half lives of GFP with and without the tail. Instead, to distinguish between stalled elongation and protein degradation, we designed a reporter construct that generates two separate polypeptides from a single round of translation through co-translational ‘cleavage’ mediated by a viral 2A peptide (Sharma et al., 2012). After screening for a 2A peptide sequence that functions efficiently in S. cerevisiae (Figure 3—figure supplement 2), we designed a construct containing HA-mRuby and GFP sequences separated by the P2A peptide (derived from porcine teschovirus-1). The GFP sequence downstream of P2A was appended with the 10-amino-acid tail (or was not, as a control), which should lead to the absence of detectable GFP regardless of the mechanism of action. In contrast, the upstream HA-tagged mRuby should only be detected if translation itself is not affected by the 10-amino-acid sequence, since it reports on the number of rounds of translation but is not covalently linked to the inhibitory tail. Assaying for GFP by immunoblotting revealed that accumulation of the protein was suppressed by the 10-amino-acid tail, as expected (Figure 3E). In contrast, HA-mRuby was detected at similar levels whether or not the tail was included in the construct. These results indicate that the tail functions downstream of translation, likely by acting as a ‘degron’ (Varshavsky, 1991) that targets the protein for immediate degradation after synthesis (Cox and Walter, 1996).

If the C-terminal tail of Hac1^up functions as a degron, additional proteins may be involved in recognizing the degron and targeting the covalently linked protein for degradation. To identify such trans-acting factors, we used a genetic approach that took advantage of the strong silencing phenotype imparted by the 10-amino-acid tail alone (Figure 3C). We constructed strains in which the first exon of HAC1 was replaced by HIS3, which we reasoned might behave like the analogous GFP reporter genes and be completely silenced by the 10-amino-acid sequence. Similarly to GFP, replacing the first exon of HAC1 with HIS3 but keeping the HAC1 locus otherwise intact prevented expression of His3p as evidenced by histidine auxotrophy (Figure 4A). Removing the entire intron restored His3p expression and growth of the corresponding strain on medium lacking histidine, suggesting that silencing was mediated by the HAC1 intron. Strains containing a stop codon between HIS3 and the intron (to prevent translation of the 10-amino-acid degron) had a low but detectable level of growth on histidine-lacking medium (Figure 4A), consistent with the weak fluorescence signal observed from the corresponding GFP reporter construct (Figure 3C). On the other hand, strains containing His3p appended with the C-terminal tail of Hac1^up but no other elements of the HAC1 intron could not grow on medium lacking histidine. These results indicate that the degron is sufficient for functional silencing of HIS3 expression, providing a useful genetic tool to identify additional genes involved in the degron-dependent silencing mechanism.

Figure 4 with 2 supplements see all

Download asset Open asset

Although we had initially intended to use chemical mutagenesis to generate silencing-defective mutants, upon streaking out the selection strain on histidine-lacking medium we noticed that a small number of slow-growing colonies appeared even without mutagen treatment. We reasoned that such spontaneous suppressor strains would have very few mutations, making it possible to identify suppressor mutations by whole-genome sequencing without requiring backcrossing or forming complementation groups. Thus, to isolate mutants with defective degron-dependent silencing ('dds mutants') we simply plated the selection strain expressing HA-tagged His3p with the 10-amino-acid tail on histidine-lacking medium and isolated the rare single colonies that grew for further analysis (Figure 4B). From five independent platings we isolated a total of 123 mutant strains that, when re-streaked, could grow on medium lacking histidine. Sanger sequencing of the C-terminal region of the reporter gene in each mutant identified 15 strains harboring a cis mutation that either altered the sequence of the degron, introduced a premature stop codon before or within the degron, or removed the stop codon of the degron resulting in a six-amino-acid C-terminal extension (Figure 4—figure supplement 1A)—all of which provided confirmation that the genetic selection worked as desired.

From the strains that displayed histidine prototrophy, we picked 35 (including four cis mutants) to analyze by immunoblotting and found that all had detectable HA-His3p (Figure 4—figure supplement 1B). We selected 20 of these strains with unknown mutations for whole-genome sequencing (as well as the parental selection strain as a reference, and three strains with known mutations in the degron as positive controls for our variant-calling procedure), taking care to select strains that varied widely in HA-His3p abundance or growth rate to minimize our chances of sequencing the same mutation in multiple strains. We sequenced the 24 genomes together in a single lane of a HiSeq sequencer using 50-nucleotide single-end reads, which provided 38–74X coverage (9.3–18.0 million reads) of each yeast genome. We then used standard mapping and variant-calling tools (BWA and FreeBayes, respectively) to identify variants that were absent from the parental genome, which successfully recovered the positive-control cis mutants. Remarkably, 17 out of the 20 strains that we sequenced contained a mutation in the same gene YJL149W (Figure 4C and Figure 4—figure supplement 1C), and in each case we confirmed the mutation by Sanger sequencing (Figure 4—figure supplement 1D). Because no other ORF was found mutated in more than one of the 20 mutant strains (Figure 4—figure supplement 1C), we focused our follow-up efforts on YJL149W.

YJL149W had previously been named DAS1 for 'Dst1-delta 6-Azauracil Sensitivity 1' when it was identified in a genetic screen unrelated to the UPR (Gómez-Herreros et al., 2012). Based on the protein’s domain structure (containing an F-box domain and leucine-rich repeats) and physical interactions with the SCF core components Cdc53p and Skp1p (Willems et al., 1999), YJL149W was annotated as a 'putative SCF ubiquitin ligase F-box protein' (Cherry et al., 2012). F-box proteins act as adapters to target substrates for ubiquitination and subsequent degradation by the proteasome (Skaar et al., 2013). We therefore propose to rename this gene DUH1 for 'Degrader of Unspliced HAC1 gene product 1' to reflect its role in the degradation of Hac1^up, as we demonstrate later.

The DUH1 variants that we identified in our 17 strains comprised 16 different mutations, of which 8 were nonsense, 7 were missense, and 1 was a frameshift (Figure 4C, right). The nonsense mutations tended to cluster in the first half of the ORF, suggesting that they likely functioned as null alleles. In addition, three of the 17 strains containing mutations in DUH1 did not contain any other mutations, implicating the DUH1 mutations as causative for the phenotype. We therefore tested whether knocking-out DUH1 (which is non-essential) in the original selection strain recapitulated the de-silencing phenotype. Disrupting DUH1 led to a dramatic increase in the steady-state abundance of HA-His3p containing the 10-amino-acid tail, while having no effect on HA-His3p lacking the tail or constructs repressed by long-range base pairing (Figure 4D). Analogously, deleting DUH1 in the GFP reporter strains completely eliminated the silencing effect of the Hac1^up tail but had no effect on base-pairing-mediated silencing (Figure 4E and Figure 4—figure supplement 2). Notably, in the absence of DUH1 the GFP reporter in a wild-type HAC1 context was now expressed at the same leaky level as previously seen for the reporter in which a stop codon was positioned between GFP and the intron, indicating that degron-dependent silencing is required to suppress leaky GFP expression. Together, these results provide genetic evidence that DUH1 is the adapter protein that recognizes the Hac1^up tail and targets the covalently attached protein for degradation.

Having established that DUH1 is required for degron-dependent silencing of two different reporter genes, we returned to HAC1 itself. To detect Hac1p we introduced a 3xHA tag at the extreme N terminus of the endogenous HAC1 allele (Figure 5A), which did not interfere with its post-transcriptional silencing in the absence of the UPR (Figure 5B and C). To determine which Hac1p isoforms are regulated by DUH1, we constructed a set of HA-tagged strains that constitutively produced either Hac1ⁱp containing the 18-amino-acid exon 2–encoded tail (construct 3), Hac1^up containing the 10-amino-acid intron-encoded tail (construct 5), or Hac1^Δtailp containing no tail at all (constructs 4 and 6). All of the mRNAs encoding HA-tagged Hac1p variants were expressed at similar levels in the presence versus the absence of DUH1 (Figure 5B). Strikingly, at the protein level only the abundance of Hac1^up was affected by disruption of DUH1, increasing by ~five fold in the knock-out strain (Figure 5C). The increased abundance of Hac1^up in the absence of DUH1 was not due to increased translation, as evidenced by deletion of DUH1 having no impact on ribosome density on any of the HA-tagged reporter mRNAs (Figure 5D). Instead, our results suggest that Duh1p specifically affects the turnover of Hac1^up due to its 10-amino-acid tail, as was suggested by the results of our reporter experiments.

Figure 5

Download asset Open asset

Because we were able to detect Hac1^up by immunoblotting (using a high-sensitivity antibody) even when DUH1 was intact (unlike the corresponding GFP reporter), we could use cycloheximide (CHX) shut-off experiments to directly assay the impact of DUH1 on the turnover of Hac1^up. In both the presence and absence of DUH1, Hac1^up was degraded so rapidly that we could not accurately measure its half life even using a rapid harvesting procedure, due to the ~2 minutes required for CHX to accumulate in cells and halt translation (Gerashchenko and Gladyshev, 2014). However, the protein-degradation kinetics allowed us to calculate an upper bound for the half life of Hac1^up, which was 50 seconds when DUH1 was present (Figure 5E). Deletion of DUH1 stabilized Hac1^up and increased its half-life upper bound to 2 minutes. These results demonstrate that DUH1 is required for the extremely short half life of Hac1^up that normally limits its accumulation. The true half-life difference upon DUH1 deletion is likely to be greater than the ~2-fold difference in upper bounds that we measured, based on the ~5-fold difference in steady-state protein levels that could not be accounted for by differences in either mRNA abundance or ribosome density (Figure 5B and D).

It was previously observed that disrupting the base-pairing interaction between the 5′-UTR and intron of HAC1 mRNA was sufficient to allow accumulation of Hac1^up, which led to a model in which base-pairing alone was responsible for the post-transcriptional silencing phenomenon (Rüegsegger et al., 2001). Our results using constructs in which the base-pairing region was deleted (Figure 5) suggested that the previously observed accumulation of Hac1^up was unknowingly being buffered by Duh1p-dependent degradation. To directly address this possibility, we generated HA-tagged constructs in which the base-pairing region was disrupted by mutations in either the 5′-UTR or intron or was reconstituted by the compensatory mutations (Figure 6A) and determined the effect of DUH1 deletion on steady-state protein levels. Because the 5′ and 3′ splice sites remained intact in these constructs, we introduced them into an ire1Δ background to eliminate any potential confounding effects of background splicing (Figure 1—figure supplement 1B). As previously observed, mutating the base-pairing region in the presence of DUH1 resulted in detectable levels of Hac1^up (Figure 6C). However, the accumulation of Hac1^up was greatly stimulated by deletion of DUH1, which was not explained by a corresponding increase in mRNA abundance (Figure 6B and C). Thus, although disrupting the base pairing produces detectable amounts of Hac1^up as previously reported (Rüegsegger et al., 2001), Duh1p-dependent degradation restricts the steady-state level of the protein.

Figure 6

Download asset Open asset

Remarkably, even a single nucleotide change in the center of the base-pairing region (Sathe et al., 2015) was sufficient for some accumulation of Hac1^up, which again was enhanced by deletion of DUH1 (Figure 6D). This result suggests that the 5′-UTR–intron base-pairing interaction is only marginally stable, which may be required for efficient dissociation of the intron after splicing (see Discussion).

Collectively, we have shown that a pair of silencing mechanisms, one translational and the other post-translational, prevents spurious production of Hac1^up in the absence of the UPR. The existence of such a fail-safe silencing mechanism implies that ectopic production of Hac1^up has physiological consequences that negatively impact cellular fitness. However, a previous study suggested that Hac1^up is itself not an active transcription factor because it lacks the activating 18-amino-acid tail found in Hac1ⁱp (Mori et al., 2000), raising the question as to why Hac1^up accumulation would need to be tightly regulated. We hypothesized that Hac1^up was in fact an active transcription factor but that in the previous study its accumulation had been prevented due to the experiments being performed in a DUH1 background. To test this hypothesis, we evaluated the ability of strains that constitutively produced either Hac1ⁱp, Hac1^up, or Hac1^Δtailp to grow under conditions of chronic ER stress induced by the drug tunicamycin. Strains expressing Hac1ⁱp or Hac1^Δtailp grew on tunicamycin-containing medium regardless of whether DUH1 (or IRE1) was present (Figure 7A), consistent with both proteins being active transcription factors that are not targeted by Duh1p. In contrast, strains expressing Hac1^up only grew robustly on tunicamycin-containing medium when DUH1 was knocked out (but independently of IRE1). These results confirm our hypothesis that Duh1p-dependent degradation normally masks the activity of Hac1^up. Our findings also explain how HAC1 was able to be initially identified as a high-copy activator of the UPR in an Δire1 strain, since the UPR activity detected in this strain had to have resulted from Hac1^up produced from unspliced HAC1 mRNA (Chapman and Walter, 1997; Cox and Walter, 1996).

Figure 7 with 1 supplement see all

Download asset Open asset

Because Hac1^up has UPR-inducing activity (Figure 7A), we reasoned that the fail-safe mechanism we discovered is required to prevent leaky production of Hac1^up that would otherwise cause Ire1p-independent activation of the UPR. However, we initially failed to detect Hac1^up produced from unspliced HAC1 mRNA even when DUH1 was disrupted (Figure 5C). On the other hand, our results from the analogous GFP and HIS3 reporter gene studies demonstrated that translational repression mediated by 5′-UTR–intron base pairing was incomplete and allowed a low level of protein synthesis that was normally 'cleaned up' by Duh1p-dependent degradation (Figure 4D and E). This led us to hypothesize that Hac1^up itself was also being occasionally produced from HAC1^u mRNA and rapidly degraded in a Duh1p-dependent manner, but that the short half life of Hac1^up (Figure 5E)—relative to both GFP (Natarajan et al., 1998) and His3p (Belle et al., 2006)—further reduced the steady-state abundance of Hac1^up to an extremely low level that was initially undetectable.

To enhance detection of HA-tagged Hac1^up, we made two modifications: We used a more sensitive anti-HA antibody for immunoblotting; and we knocked out IRE1 in our strains, which eliminates the background Ire1p-dependent splicing of HAC1 mRNA (Figure 1—figure supplement 1B) and concomitant production of Hac1ⁱp that can otherwise dominate the signal on immunoblots. With these modifications we could now detect Hac1^up being produced from unspliced HAC1 mRNA, but only when the protein was stabilized by deletion of DUH1 (Figure 7B) and at levels far below even those of Hac1^up being constitutively produced in the presence of DUH1 (Figure 7—figure supplement 1). Despite the relatively low level of leaky translation product we detected, these results provide molecular evidence that Hac1^up is being continuously produced from HAC1^u mRNA but rapidly degraded due to its C-terminal degron.

Does the leaky production of Hac1^up unmasked by deletion of DUH1 have any functional consequences? The ability of constitutively produced Hac1^up to promote survival under UPR-inducing conditions (Figure 7A) suggested that small amounts of Hac1^up might also induce the UPR to some extent. To test this possibility, we examined how strains expressing HA-tagged but otherwise unmodified HAC1 mRNA grew on media containing different concentrations of tunicamycin. At the highest tunicamycin concentration only strains expressing IRE1 could grow regardless of whether DUH1 was also present (Figure 7C), suggesting that growth was dependent on the abundant Hac1ⁱp produced from HAC1ⁱ mRNA. In contrast, at lower concentrations of tunicamycin the IRE1 deletion strain showed some growth, but this was reproducibly enhanced by simultaneous deletion of DUH1. These results indicate that the low level of Hac1^up detectable in strains lacking DUH1 (Figure 7B) is sufficient to activate the UPR enough to facilitate cell survival under stress, even in the absence of Ire1p. Thus, degron-dependent degradation of Hac1^up mediated by Duh1p is normally required to prevent ectopic Ire1p-independent activation of the UPR.

Saccharomyces cerevisiae strains used in this study were derived from BY4741 (MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0) and are listed in Supplementary file 1. Strains were cultured at 30°C with shaking in rich yeast-peptone-dextrose (YPD) medium, synthetic complete (SC) medium, or the appropriate drop out medium (e.g., SC–His) as indicated. Cultures were grown to mid-log phase (OD₆₀₀ ~0.5) before harvesting by either vacuum filtration (polysome analysis), methanol quenching (protein half-life), or brief centrifugation at 4°C (all other experiments) and then snap frozen in liquid nitrogen. For growth tests under UPR stress (Figure 7), 10-fold serial dilutions of cells were spotted onto YPD plates containing either the indicated concentration of tunicamycin (Sigma, St. Louis, MO) dissolved in DMSO (+Tm) or DMSO alone (–Tm).

To facilitate generation of HAC1 variant and reporter strains, the entire transcribed region of the HAC1 gene – as well as additional flanking sequence – was amplified from genomic DNA (extracted from BY4741) and cloned into the pCR4-TOPO vector backbone (Invitrogen, Carlsbad, CA) using Gibson Assembly Master Mix (New England Biolabs, Ipswich, MA) according to the Gibson assembly method (Gibson et al., 2009). GFP and HIS3 reporter constructs were generated from this plasmid by replacing the HAC1 ORF with the reporter gene using the Gibson assembly method. Variants of these constructs were generated using inverse PCR with 5′-phosphorylated primers (designed to introduce point mutations or deletions) and Phusion High-Fidelity DNA Polymerase (New England Biolabs) followed by ligation with Quick Ligase (New England Biolabs); or using the Gibson assembly method with either PCR products or synthetic gBlocks (Integrated DNA Technologies, Coralville, IA). All plasmid inserts were verified by Sanger sequencing before PCR amplification with Phusion DNA polymerase for integration into the yeast genome.

Yeast transformations were performed using the PEG–lithium acetate method (Gietz and Woods, 2002). HAC1 was deleted from the genome using the URA3 cassette of pRS416 to replace the entire transcribed region of HAC1 (from 366 nucleotides upstream of the start codon, to 750 nucleotides downstream of the stop codon in exon 2). IRE1 and/or DUH1 were deleted using the HIS3MX or URA3 cassettes of pFA6a-3HA-HIS3MX6 or pRS416, respectively, as indicated in Supplementary file 1. HAC1 variant and reporter strains were generated from hac1Δ::URA3 strains using 5-FOA counterselection. Transformants were screened by PCR to identify integrants, which were subsequently verified by PCR and Sanger sequencing of the entire integrated cassette.

Polysome analyses by sucrose gradient fractionation are described in more detail at Bio-protocol (Aboulhouda et al., 2017). Yeast strains were grown to mid-log phase (OD₆₀₀ ~ 0.5) in YPD and harvested by vacuum filtration as previously described (Weinberg et al., 2016). The frozen cell pellet was transferred into a pre-chilled mortar that was surrounded by and filled with liquid nitrogen. The pellet was ground to a fine powder by hand using a pre-chilled pestle, transferred into a 50 ml conical tube filled with liquid nitrogen, and after boiling off the liquid stored at –80°C. Crude lysate was prepared by briefly thawing the cell powder on ice for 1 min and then resuspending in 1 ml polysome lysis buffer (20 mM HEPES-KOH [pH 7.4], 5 mM MgCl₂, 100 mM KCl, 1% Triton X-100, 2 mM DTT, 100 μg/mL cycloheximide, 20 U/mL SuperUPERase-In [Thermo Fisher Scientific, Waltham, MA], 1X cOmplete EDTA-free Protease Inhibitor Cocktail [Roche, Switzerland]). Where indicated, the lysis buffer also contained 200 μg/mL heparin (Sigma). The lysate was centrifuged at 1300 g for 10 min, and the supernatant was transferred to a new tube taking care to avoid the thick top lipid layer. The resulting clarified lysate was diluted to a concentration of 25 OD₂₆₀ units/ml, flash frozen in liquid nitrogen in single-use aliquots, and stored at –80°C.

A 10–50% continuous sucrose gradient was prepared in polysome gradient buffer (20 mM HEPES-KOH [pH 7.4], 5 mM MgCl₂, 100 mM KCl, 2 mM DTT, 100 μg/mL cycloheximide, 20 U/mL SUPERase-I) and allowed to cool to 4°C while lysate was thawed gently on ice. Where indicated, 100 ng luciferase RNA (Promega, Madison, WI) was then added to 100 μl of thawed cell lysate. 100 μl of cell lysate (at 25 OD₂₆₀ units/ml) was loaded on top of the gradient, and gradients were spun in an SW41-Ti rotor at 36000 rpm for 2.5 hr at 4°C. Gradients were fractionated into 15 fractions using a Biocomp Piston Gradient Fractionator, and fractions were flash frozen and stored at –80°C.

RT-qPCR was performed directly on gradient fractions using the TaqMan Gene Expression Cells-to-CT Kit (Life TechnologiesThermo Fisher Scientific). Briefly, 10 μl Lysis Buffer containing DNase was combined with 0.1 μl synthetic XenoRNA control (provided with the TaqMan Cells-to-CT Control Kit) and 1 μl of the fraction. Reactions were incubated at room temperature for 5 min, and then quenched on ice by addition of 1 μl STOP Solution followed by incubation at room temperature for an additional 2 min. 4.5 μl of this reaction was added to 5.5 μl RT Master Mix, and reverse transcription was carried out at 37°C for 1 hr followed by a 5 min incubation at 95°C before cooling to 4°C. RT reactions were diluted 6-fold in nuclease-free water prior to qPCR.

qPCR was carried out in duplicate for each fraction using the TaqMan Gene Expression Master Mix (Thermo Fisher Scientific) according to the manufacturer’s instructions, using 10 μl reactions containing 4.5 μl of six-fold diluted RT reaction per qPCR reaction. XenoRNA was analyzed using the primer-probe assay provided with the TaqMan Cells-to-CT Control Kit. ACT1 and luciferase RNAs were analyzed using predesigned TaqMan Gene Expression Assays from Life TechnologiesThermo Fisher Scientific (Sc04120488_s1 and Mr03987587_mr, respectively). HAC1 and GFP RNAs were analyzed using the following primer-probe qPCR assays from Integrated DNA Technologies (containing 6-FAM/ZEN/IBFQ quenchers with a primer-to-probe ratio of 1-to-2):

Relative mRNA abundances in each fraction (i.e., Cq values) were first normalized to XenoRNA to account for differences in qRT-PCR efficiency among fractions, and then calculated as a percentage of total mRNA detected across the gradient. All strains were analyzed in duplicate, beginning with separate cultures of the same strain, to confirm the reproducibility of each result. ACT1 mRNA was analyzed in every gradient and confirmed to co-sediment with polysomes in all cases.

Yeast strains were grown to mid-log phase (OD₆₀₀ ~ 0.5) in SC medium, then diluted 10 fold in fresh medium before quantifying GFP fluorescence using a LSR II flow cytometer (Becton Dickinson, Franklin Lakes, NJ) and 530/30 filter. Raw data from 10,000 cells per sample was analyzed using FlowJo and gated to remove debris before being exported to Microsoft Excel. Fluorescence in each cell was normalized to cell size using the side scatter measurement before calculating the median and quartiles of the population. All strains were analyzed at least twice, beginning with separate cultures of the same strain, to confirm reproducibility. The equivalent of 5 OD₆₀₀ units was harvested for both protein and RNA analyses from the same cultures used for flow cytometry.

Protein samples were prepared using the NaOH/TCA method (Riezman et al., 1983), separated by SDS-PAGE using 12% or 4–12% Bolt Bis-Tris gels (Thermo Fisher Scientific), and transferred in 1X CAPS Buffer onto 0.22 micron PVDF membrane (Bio-Rad, Hercules, CA). Blots were probed with the following antibodies diluted in 1X TBS-T containing 5% nonfat dry milk: mouse anti-GFP (RRID:AB_390913, Roche #11814460001, 1:1000), mouse anti-HA (RRID:AB_627809, Santa Cruz Biotechnology [Dallas, TX] sc-7392, 1:3000), rat anti-HA high sensitivity (RRID:AB_390918, Roche #11867423001, 1:5000), mouse anti–beta actin (RRID:AB_449644, AbCam [] ab8224, 1:10000), HRP-conjugated goat anti-mouse IgG (RRID:AB_631736, Santa Cruz Biotechnology sc-2005, 1:10000), and HRP-conjugated goat anti-rat IgG (RRID:AB_631755, Santa Cruz Biotechnology sc-2032, 1:10000). Blots were developed using Clarity ECL Western Blotting Substrate (Bio-Rad), and chemiluminescence was detected on a ChemiDoc Imaging System (Bio-Rad).

Total RNA was isolated using the hot acid phenol method (Ares, 2012) and resuspended in nuclease-free water. For each sample, 2.5 μg of RNA was treated with RQ1 DNase (Promega) according to the manufacturer’s instructions. cDNA synthesis and qPCR were performed using the Cells-to-CT Kit (Thermo Fisher Scientific) as described above.

For RT-PCR analysis of HAC1 splicing, cDNA generated from input samples (as described in 'Sucrose gradient analysis') was used as a template for PCR amplification using the following intron-flanking primers: forward, ACGACGCTTTTGTTGCTTCT; reverse, TCTTCGGTTGAAGTAGCACAC. PCR products were analyzed by agarose gel electrophoresis.

Yeast strains YRDS221 and YRDS241 (Supplementary file 1) were grown to mid-log phase (OD₆₀₀ ~ 0.5) in SC medium. Cycloheximide (Sigma) was added to a final concentration of 100 μg/ml and samples collected at the indicated time points. To rapidly harvest samples, 5 ml of culture was directly added to a prechilled 50 ml tube (in a dry ice–ethanol bath) containing 25 ml Quench Solution (60% methanol, 10 mM HEPES-KOH pH 7.4) and mixed well. Cells were collected by centrifugation at 4°C, snap frozen, and processed for immunoblotting as above. Quantification of immunoblot signal was performed by densitometry with ImageJ. An upper bound on protein half-life was calculated from 3 independent experiments as the time point at which ~50% of the protein remained.

Yeast strain YRDS57 (Supplementary file 1) was grown overnight in liquid YPD, plated on SC–His solid medium, and allowed to grow for 5–7 days at 30°C. Spontaneous mutant colonies were isolated from five independent platings and verified for growth on SC–His by restreaking and for expression of the HA-His3p by immunoblotting. Clones were then examined for cis mutations by Sanger sequencing the 3′ end of the HIS3 reporter gene. Of the remaining clones that had a ‘trans’ mutation, 20 were chosen for analysis by genomic DNA sequencing along with 4 control strains (the parent strain YRDS57 and 3 strains with cis mutations).

Genomic DNA was isolated from individual saturated overnight cultures using the MasterPure Yeast DNA Purification Kit (Epicentre) according to the manufacturer’s instructions. Purified genomic DNA was treated with RNase A/T1 Cocktail (Thermo Fisher Scientific), phenol extracted, precipitated with ethanol, and quantified with the Qubit dsDNA HS Assay Kit (Thermo Fisher Scientific). For each strain 1 ng of genomic DNA was used as input into the Nextera XT DNA Library Preparation Kit (Illumina, San Diego, CA), and genomic DNA libraries were prepared according to the manufacturer’s instructions using the Nextera XT Index Kit for barcoding during PCR. Genomic DNA libraries from each strain were quantified with the Qubit dsDNA HS Assay Kit (Invitrogen), and pooled into a single tube in equal proportions. Pooled DNA was purified using the DNA Clean and Concentrator-5 Kit (Zymo Research, Irvine, CA), and 200–350 bp fragments were isolated using a BluePippin Gel Cassette (2% agarose dye-free with internal standards; ). The size-selected library was sequenced on a single lane of a HiSeq 4000 sequencer using 50-nucleotide single-end reads.

Data analysis was performed using the Galaxy web server (Afgan et al., 2016 ). Reads in FASTQ format were aligned to the sacCer3 reference genome using BWA (version 0.1) with default 'Commonly Used' settings. Sequence differences between the reference genome and aligned reads were identified using FreeBayes (version 0.4) to generate a variant call format (VCF) file for each strain. The VCF-VCFintersect tool (with options '-i --invert') was then used to compare VCF files for mutants against the VCF file for the parental selection strain, in order to identify variants that were unique to the mutants. Finally, variants were manually annotated using the UCSC Genome Browser. For proof-of-principle analysis of cis mutants, the same mapping and variant-calling procedures were used with a custom reference genome containing only the reporter gene.

Genome-sequencing data from the genetic selection have been deposited in the NCBI Sequence Read Archive (SRA) under accession SRP081128.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Peter Walter, David Toczyski, Graeme Gowans, and members of the Weinberg lab for insightful discussions and comments on the manuscript, Manuel Leonetti and Meghan Zubradt for assistance with flow cytometry, and Eric Chow for advice about high-throughput sequencing. We also acknowledge Jonathan Weissman, Raul Andino, Keith Yamamoto, and Alan Frankel for generously sharing equipment. High-throughput sequencing was performed at the UCSF Center for Advanced Technology. This work was supported by the UCSF Program for Breakthrough Biomedical Research (funded in part by the Sandler Foundation) and by an NIH Director’s Early Independence Award (DP5OD017895).

© 2016, Di Santo 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.