Abstract
Key protein adapters couple translation to mRNA decay on specific classes of problematic mRNAs in eukaryotes. Slow decoding on non-optimal codons leads to codon-optimality-mediated decay (COMD) and prolonged arrest at stall sites leads to no-go decay (NGD). The identities of the decay factors underlying these processes and the mechanisms by which they respond to translational distress remain open areas of investigation. We use carefully-designed reporter mRNAs to perform genetic screens and functional assays in S. cerevisiae. We characterize the roles of Hel2 and Syh1 in coordinating translational repression and mRNA decay on NGD reporter mRNAs, finding that Syh1 acts as the primary link to mRNA decay in NGD. Importantly, we observe that these NGD factors are not involved in the degradation of mRNAs enriched in non-optimal codons. Further, we establish that a key factor previously implicated in COMD, Not5, contributes modestly to the degradation of an NGD-targeted mRNA. Finally, we use ribosome profiling to reveal distinct ribosomal states associated with each reporter mRNA that readily rationalize the contributions of NGD and COMD factors to degradation of these reporters. Taken together, these results provide new mechanistic insight into the role of Syh1 in NGD and define the molecular triggers that determine how distinct pathways target mRNAs for degradation in yeast.
Introduction
Translation of mRNAs to produce proteins is a fundamental cellular process that supports the cell’s ability to carry out the basic enzymatic reactions needed for life. To prevent errors that arise during this complex process from compromising cellular metabolism, specialized molecular pathways have evolved to recognize and regulate problematic translation events (D’Orazio and Green 2021; Inada 2017; Yan and Zaher 2019). These mechanisms are coupled to RNA decay pathways that target problematic mRNAs and prevent continuing diversion of ribosomes toward unproductive translation. The set of factors involved in this crucial recognition of problems that arise during translation elongation and the mechanisms by which they exert their downstream effects on mRNA stability remain only partially characterized.
General mRNA decay in yeast is catalyzed primarily by mRNA decapping and 5’ to 3’ exonucleolytic degradation by Xrn1, while the 3’ to 5’ exonuclease (the exosome) is thought to play a role only under certain circumstances (Muhlrad, Decker, and Parker 1994). Recent foundational work in yeast, and subsequently in zebrafish and mammals, discovered that mRNA stability is correlated with its codon usage (Presnyak et al. 2015; Mishima and Tomari 2016; Q. Wu et al. 2019): mRNAs enriched in non-optimal codons have short half-lives and are rapidly degraded by the cytoplasmic Ccr4-Not deadenylation complex and the decapping activator Dhh1 (Radhakrishnan et al. 2016; Webster et al. 2018; Sweet, Kovalak, and Coller 2012). Recent biochemical and structural evidence supports a model in which suboptimal codons in the ribosomal A site slow down translation elongation, allowing deacylated tRNA to diffuse away from the E site and enabling the critical adaptor protein Not5 to bind. Not5 binding in the vacant ribosomal E sites can recruit the Ccr4-Not complex to promote mRNA deadenylation, decapping, and decay (Buschauer et al. 2020). These observations provide molecular insight into codon-optimality-mediated decay (COMD) and support for the idea that this pathway represents an important determinant of general cellular mRNA half-lives.
In contrast to the normal slowing of translation that occurs transiently as ribosomes decode less optimal codons, the cell also possesses quality-control machinery to resolve more deleterious ribosomal stalls that can arise from chemical damage in the mRNA (truncation, depurination, nucleobase dimers, oxidative damage, etc.), difficult to unwind secondary structure, or incorrect nuclear mRNA processing events (D’Orazio and Green 2021). Adjacent pairs of specific rare codons can mimic these events and induce strong inhibition of translation elongation and associated mRNA decay in S. cerevisiae (Gamble et al. 2016). For example, consecutive CGA codons induce terminal stalls and have been routinely included in reporter mRNAs to trigger an mRNA surveillance pathway referred to as no-go decay (NGD) (Tsuboi et al. 2012; Letzring et al. 2013; Tesina et al. 2020). On these problematic mRNAs, ribosomes stall on the CGA codons, leading to ribosomal collisions that promote small-subunit protein ubiquitination by the E3 ligase Hel2 (mammalian ZNF598), ribosomal clearance by the helicase Slh1 and ribosome quality control trigger (RQT) complex (Ikeuchi, Izawa, and Inada 2019), and nascent peptide decay by the ribosome quality control (RQC) complex (Brandman et al. 2012). The accumulation of colliding ribosomes is thought to trigger decapping and Xrn1-mediated mRNA degradation, though the specific molecular players and interactions responsible for triggering this decay remain poorly defined (D’Orazio et al. 2019; Simms et al. 2019). Additionally, under conditions where the ribosome rescue machinery is compromised or overwhelmed, cleavage of reporter mRNAs by the endonuclease Cue2 and Dom34-mediated rescue provides an alternate route for mRNA degradation and ribosome rescue (Doma and Parker 2006; D’Orazio et al. 2019; Glover et al. 2020). NGD, as a “quality control” pathway, is thought to minimize the detrimental effects from damaged or problematic mRNAs in the cell and to reduce the overall impact of proteotoxic stress (Brandman et al. 2012; Ishimura et al. 2014; Martin et al. 2020).
Though NGD and COMD can be triggered by seemingly similar mRNA sequences and converge on Xrn1-mediated exonucleolytic decay of mRNAs (Pelechano, Wei, and Steinmetz 2015), the extent to which these processes overlap in specificity and activity remains unclear, as do their complete sets of accessory factors. Moreover, the molecular states of the ribosome which define and activate these pathways have not been systematically compared. In this study, we address these questions through genetic screening and functional assays in the yeast S. cerevisiae. We use reporter mRNAs designed to trigger NGD or COMD and perform synthetic genetic array (SGA) screens to identify factors critical to these separate mRNA decay pathways. Importantly, we identify a critical role for Syh1 as the primary effector for decay for mRNAs with terminal stalls. We use flow cytometry and northern blotting combined with genetic perturbations to reveal the contributions of other NGD factors and COMD factors to translational repression and decay of the reporter mRNAs. Finally, we use ribosome profiling of NGD and COMD reporters to isolate the activities of the major players in these pathways and connect these activities to the molecular states of elongating ribosomes. These data provide a basis for understanding the unique contributions of each of these pathways to translation-coupled mRNA decay and contextualizes their effects in the larger cellular process of translation surveillance.
Results
A genetic screen identifies NGD factors in yeast
To identify protein factors that contribute to NGD, we developed reporter constructs with well-defined sequence features designed to trigger ribosome stalling and associated quality control. Reporter mRNAs were under the control of the inducible, bidirectional GAL1-10 promoter and encoded GFP followed by either a fully codon-optimized yeast HIS3 gene (termed OPT) or HIS3 interrupted by twelve repeats of the highly non-optimal CGA codon (termed CGA; Figure 1A). This repeat sequence has been shown to trigger NGD in S. cerevisiae by causing strong stalling of ribosomal elongation (due to overall low abundance of tRNAArg(ICG) compounded by inefficient decoding by the I:U wobble interaction) and ensuing ribosome collisions (Letzring, Dean, and Grayhack 2010; Tesina et al. 2020). A viral P2A sequence (Brown and Ryan 2010; Sharma et al. 2012) was inserted between the GFP and HIS3 open reading frames to decouple GFP levels from the protein decay induced by RQC factors in response to ribosome pausing on CGA codons. Importantly, knockout of LTN1, the major E3 ligase responsible for nascent peptide degradation by the RQC complex, did not increase the GFP/RFP ratio for our reporter (Figure S1A), demonstrating that nascent peptide decay does impact the levels of GFP protein. As a result, for this construct, GFP levels serve as a proxy for reporter mRNA levels and translation initiation rates, allowing us to follow these activities in individual cells by flow cytometry. An RFP mRNA is produced from the same GAL1-10 promoter in the reverse direction, allowing RFP fluorescence to be used to normalize for average transcription and metabolic changes within individual cells.
We performed a Reporter-Synthetic Genetic Array (R-SGA) screen (Fillingham et al. 2009) by introducing the OPT and CGA reporters into the 5377 yeast strains contained in the Yeast Knockout Collection (Giaever et al. 2002). A total of 4222 deletion strains were successfully grown and tested with fluorimetry. We obtained GFP and RFP data for each deletion strain with these two reporters and calculated Z-scores of the GFP/RFP ratio for every strain on a per-plate basis, allowing comparison between the CGA screen and the previously published OPT screen (Figures 1B, S1B) (D’Orazio et al. 2021). To identify genes contributing to NGD, we focused on knockout strains in which normalized CGA reporter levels were significantly increased or decreased relative to our normalized OPT reporter levels (Supplementary Table 1, Figures 1B, S1B). Among the strongest hits from the screen were known NGD factors including HEL2 and members of the RQT complex (SLH1, CUE3, and RQT4), all of which exhibited substantially decreased GFP reporter fluorescence compared to the wild-type control. This suggests that loss of these factors causes increased decay of the reporter mRNA. Among the strongest hits that increased CGA reporter levels, we identified the ribosomal protein gene ASC1 and the genes SYH1 and SMY2, homologs of the mammalian NGD factors GIGYF1/2, which we previously reported to impact CGA reporter levels in yeast (Hickey et al. 2020). We also performed a similar R-SGA screen using a reporter identical to the CGA reporter, except with the CGA12 repeat replaced by AAA12; results from this screen showed broad overlap with the CGA reporter screen (Figures S1C-D) as anticipated based on the similar stalling mechanisms of these sequences (Tesina et al. 2020; Koutmou et al. 2015).
Of the CGA reporter strains tested in the original screen, we selected 170 with the strongest increases or decreases in GFP levels (−2 > CGA Z-score > 2 and −2 < OPT Z-score < 2) and individually validated them by flow cytometry to determine which backgrounds affected CGA reporter levels. As in the initial screen, HEL2, SLH1 and RQT4 as well as SYH1 and SMY2 (Supplementary Table 2, Figure S1E) strongly impacted GFP levels, validating our screen results and providing confidence for further mechanistic analysis. A gene set enrichment analysis (GSEA) revealed additional significant hits including genes involved in tRNA modification and protein modification that were not explored further here (Supplementary Table 3, Figure S1F).
Functional assays recapitulate known effects of NGD Factors
To further explore the hits from our genome-wide screen, we deleted genes of interest to verify the observed effects in a clean knockout background and to establish the mechanism of repression for the CGA reporter. We integrated the OPT and CGA reporters at the ADE2 locus in yeast strains with deletions of SYH1 and other factors including HEL2, SLH1, SKI2 and XRN1. While our screen identified two homologs of mammalian GIGYF1/2, our earlier studies had shown that stronger effects were associated with deletion of SYH1 than with SMY2 (Hickey et al. 2020), therefore for simplicity, we focused here on the SYH1 deletion strain. In flow cytometry experiments, we found that the GFP/RFP fluorescence ratios for the OPT reporter were not strongly affected by deletion of HEL2, SLH1, SKI2 or SYH1 (Figure 1C). In the wild-type strain, CGA reporter levels were reduced ∼3-fold in comparison to OPT as expected for a reporter subject to active NGD. Consistent with the screen results, CGA reporter levels were further reduced by about 2-fold in the slh1Δ and hel2Δ strains, while the CGA reporter levels were rescued by ∼ 1.7-fold in the syh1Δ strain (Figure 1C); in contrast, knockout of the SKI2 gene had no effect on CGA reporter levels.
We next looked directly at mRNA levels in these strains using northern blotting to ask whether GFP levels reflect the mRNA levels. As seen in the flow cytometry assay, OPT mRNA reporter levels were unaltered across all the deletions strains and the CGA mRNA reporter level was reduced relative to the OPT mRNA reporter level in the wild-type strain (Figure 1D). In agreement with the flow cytometry data and previous literature, ski2Δ had no effect on CGA reporter levels, whereas xrn1Δ strongly rescued reporter levels (D’Orazio et al. 2019; Simms et al. 2019). These results are consistent with the long established importance of Xrn1 in general mRNA decay (Muhlrad, Decker, and Parker 1994) and its critical role in NGD (D’Orazio et al. 2019). Other patterns observed in the flow cytometry data were recapitulated here as well: reporter levels were decreased to a similar extent in the hel2Δ and slh1Δ strains as in flow cytometry (Figure 1E). And, as revealed by our screen, syh1Δ substantially rescued mRNA reporter levels. This focused analysis of protein and RNA reporter levels in these different strains validates results from the initial screen and indicates that the CGA reporter is being strongly regulated by canonical NGD machinery.
Syh1 is the primary NGD factor in yeast
Since screens in this study and our previous work (D’Orazio, et al. 2019) implicated HEL2, CUE2, and SLH1 in altering CGA reporter levels, we asked how knockouts of these factors would alter the reporter mRNA levels in an syh1Δ strain to get some indication about epistasis. In order to compare more systematically the effects of these knockouts, we turned to a simpler set of reporters expressed from plasmids and containing problematic sequences within the HIS3 gene but lacking the upstream GFP ORF and the P2A sequence (Figure 2A). The P2A sequence, in particular, has been shown to induce ribosome collisions during translation thus confounding interpretations for certain reporters (C. C.-C. Wu et al. 2020). The minOPT reporter is a fully codon optimized, N-terminally FLAG-tagged HIS3 sequence expressed under a GAL promoter, and the minCGA reporter is identical to the minOPT reporter except it includes twelve CGA repeats in the same codon position within HIS3 as in the GFP-containing CGA reporter used in screening.
To recapitulate our findings with the new minCGA stalling reporter, we used northern blotting to assess the steady-state levels of the reporter mRNA (Figures 2B-C). First, hel2Δ does not decrease minCGA reporter levels as observed with the GFP-containing CGA reporter; this discrepancy may arise from effects of the P2A sequence or from differences in levels of expression and thus in ribosome loading. Importantly, although the loss of CUE2 alone has little effect on the minCGA reporter, the loss of SLH1 results in a decrease in the amount of full-length mRNA and a corresponding accumulation of a 3’ reporter fragment (Figure 2B, HIS3 probe bottom band; Figures 2C, S2A). This is consistent with our previous studies that established that NGD proceeds through Xrn1 under normal circumstances, whereas the endonuclease Cue2 plays a more important role in the absence of SLH1 (D’Orazio et al. 2019). In this paradigm, Cue2 acts a failsafe that only cuts mRNAs when stalled ribosomes accumulate when Slh1 is absent or overwhelmed. Finally, here again we observe a modest rescue of reporter levels in the syh1Δ strain, consistent with a model where Syh1 recruits Xrn1 and the machinery to mediate decay of the CGA reporter.
To further probe the role of Syh1, we created double knockout strains with syh1Δ and other relevant factors and then performed northern blot analysis to follow the levels of the minCGA reporter mRNA. Strikingly, we observe that the hel2Δsyh1Δ strain shows a complete rescue of the minCGA mRNA to the levels of the minOPT reporter (Figure 2C). A similar strong rescue was seen with the GFP-containing CGA reporter (Figure S2B). Because Hel2 functions upstream of other NGD factors in its role as an E3 ubiquitin ligase marking colliding ribosomes (Brandman et al. 2012; Saito, Horikawa, and Ito 2015; Matsuo et al. 2017; Sundaramoorthy et al. 2017; Juszkiewicz et al. 2018), the loss of Hel2 has many potential effects on mRNA decay. We found that the cue2Δsyh1Δ strain also showed much higher levels of minCGA reporter, arguing that Cue2 cleavage (which requires Hel2 activity) is the downstream step that degrades mRNA in the absence of Syh1. Further deletion of SLH1 had little or no additive effect in the cue2Δsyh1Δ strain. These results indicate that Syh1 is the major factor contributing to NGD in the absence of HEL2, but that Cue2 plays a major compensatory role in decay when Slh1 or Syh1 activity is impaired.
Several recent studies argued that mammalian GIGYF2 is recruited to collided ribosomes by the factor EDF1 (Sinha et al. 2020; Juszkiewicz et al. 2020). S. cerevisiae has a homolog of EDF1 known as Mbf1, a protein previously implicated in ribosome-mediated quality control pathways (Hendrick et al. 2001; Wang et al. 2018) that has been shown to interact with collided ribosomes in vitro (Sinha et al. 2020; Pochopien et al. 2021). Using tagged Syh1-TAP and Smy2-TAP, we performed affinity purification-mass spectrometry (AP-MS) to search for binding interactions that could help explain the mechanism of action of Syh1/Smy2. While our AP-MS data show strong enrichment of small and large subunit ribosomal proteins, indicating a connection to translation, Mbf1 was not identified as bound to Syh1-TAP or Smy2-TAP (Figure S2C; Supplementary Table 4). Moreover, we find that CGA reporter levels are unaffected in the mbf1Δ strain, suggesting that in S. cerevisiae this factor is not critical for RNA decay mediated by Syh1 (Figure S2D).
Similarly, we examined CGA reporter expression in yeast strains lacking Eap1, an Syh1-associated translation repressor thought to have a similar mechanism of action to the GIGYF2-associated mammalian protein 4EHP (Sezen, Seedorf, and Schiebel 2009; Cosentino et al. 2000) which is known to work together with GIGYF1/2 to bring about translational repression (Morita et al. 2012; Peter et al. 2019). Again, CGA reporter levels are unaffected in the eap1Δ strain relative to WT and in the eap1Δ strain relative to hel2Δeap1Δ (Figure S2E), meaning that the Syh1-mediated loss in GFP signal occurs independently of Eap1. Additionally, Eap1 was not among the proteins identified in AP-MS (Figure S2C). These data together suggest that Syh1 recruitment and function in S. cerevisiae differs substantially from that observed for GIGYF1/2 in mammalian cells.
COMD does not require canonical NGD factors
Given that the CGA reporter simply contains a stretch of highly non-optimal codons, we wondered whether the same set of factors might similarly regulate ORF sequences containing more widely distributed non-optimal codons. We tested this possibility first by developing a reporter similar to that used for the NGD screen with an N-terminal GFP, an internal P2A sequence, and a downstream HIS3 gene with an internal stretch of 129 codons synonymously re-coded as non-optimal (NONOPT; Figure 3A). As expected, the NONOPT reporter exhibited substantially diminished GFP levels compared to the OPT reporter and thus provided a starting point for subsequent analysis (Figure 3B).
To identify potential contributing factors to COMD, we performed an R-SGA screen (as above) with this NONOPT reporter (Figure 3C) and again compared these results to our OPT screen results. Strikingly, none of the known NGD factors that we had identified in the previous NGD-CGA screen emerged. These data provide a first indication that NGD mRNAs are regulated very differently from non-optimal mRNAs at the molecular level. Interestingly, factors previously implicated in stabilizing non-optimally coded mRNAs (Webster et al. 2018; Radhakrishnan et al. 2016; Buschauer et al. 2020) were also not among the strains that revealed increases in NONOPT reporter levels (Figure 3C). While some of these strains are not present in the deletion strain collection due to their severe growth phenotype (dhh1Δ and not4/5Δ), knockouts of several other members of the Ccr4-Not deadenylase complex were present in the screen including caf40Δ, caf130Δ, and not3Δ; among these, NONOPT reporter levels were modestly increased only in the caf40Δ strain.
Consistent with the results from the SGA screen, knockouts of the major factors implicated in our analysis of NGD (hel2Δ or syh1Δ) had no effect on NONOPT reporter expression levels (Figure 3B). Together, these data suggest that there are fundamental differences in recognition by the decay machinery of overall non-optimal coding sequences and more problematic strong translational stalls.
Exploring differences between non-optimal and NGD-triggering mRNAs
In order to compare more systematically the effects of mRNA sequences that trigger NGD or COMD, we turned again to the minOPT and minCGA reporters, with the addition of a third reporter called minNONOPT. This minNONOPT reporter encodes the same HIS3 ORF recoded with highly non-optimal codons; importantly, there is a short stretch of sequence retained in the minNONOPT reporter that matches the minOPT reporter to allow a common oligonucleotide probe to be used in northern blots (Figure 4A). To evaluate these new reporters, we employed a northern-blot-based transcriptional shutoff assay to measure RNA half-lives as previously reported (Radhakrishnan et al. 2016) in various deletion strains including those implicated above in NGD as well as those previously implicated in COMD.
We first asked whether the half-lives of the minOPT reporter were affected by deletion of the various factors. As anticipated, neither of the factors implicated in regulation of NGD reporters (HEL2 or SYH1) had any discernible effect on the stability of the minOPT reporter mRNA (Figures 4B, S4A-B, Supplementary Table 5). We next determined the effects of the same factors on stability of the minNONOPT mRNA reporter and found that deletion of the NGD factor genes HEL2 and SYH1 had no effect while NOT5 deletion stabilized this mRNA by approximately 11-fold, consistent with earlier reports (Buschauer et al. 2020).
Lastly, we evaluated the effects of the same set of factors on minCGA mRNA stability. There we found that deletion of HEL2 has no discernible impact on minCGA reporter half-life and deletion of SYH1 increases half-life 2-fold, while deletion of both factors (hel2Δsyh1Δ strain) increases half-life 7-fold, consistent with results from our steady-state northern blots. These results additionally agree with steady-state measurements reported for the screening reporter constructs (Figure S2B). Finally, somewhat surprisingly, we find that deletion of NOT5 increases CGA reporter half-life (∼2-fold), suggesting features of general non-optimality for this reporter mRNA. Together, these data are broadly consistent with the existence of multiple modes for recognition of troubled elongating ribosomes, each recognized by a distinct set of factors, and leading to decapping and Xrn1-mediated exonucleolytic decay.
Ribosome conformations drive distinct mRNA decay pathways
With the goal of connecting ribosome states to downstream consequences, we next employed ribosome footprint profiling (Ribo-seq) to characterize the positions and conformational states of elongating ribosomes on the minimal reporter mRNAs (Ingolia et al. 2009). To increase the resolution of the approach, both cycloheximide and tigecycline, two specific elongation inhibitors, were added to lysates to capture distinct rotational states of the ribosome during translation represented by two populations of ribosome protected fragment (RPF) lengths centered at approximately 21 and 28 nucleotides (C. C.-C. Wu et al. 2019). The 21-mer population captures ribosomes without A-site tRNAs (waiting to decode) while the 28-mer population captures ribosomes with filled A sites (waiting to translocate). Finally, to further increase the resolution of the study, we separately sequenced single-ribosome (monosome) footprints and those from nuclease-resistant disomes, thought to represent the collided structure that triggers NGD (Guydosh and Green 2014).
We next looked at the distribution of RPFs along our reporter mRNAs. We first see that ribosomes are relatively evenly distributed across the entire ORF in the minOPT reporter for both the 21-mer and 28-mer tracks (Figure 5A). As previously observed for the GFP-containing CGA reporter (Sitron, Park, and Brandman 2017; D’Orazio et al. 2019), we observed that the CGA stall region profoundly disrupts translation of the minCGA ORF. The density of monosome ribosome footprints (both 28- and 21-mers) downstream of the CGA stall is greatly reduced compared to the upstream region on this reporter, indicating that the CGA repeat region comprises a significant translational block (Figure 5A, center; Figure S5B, top). Importantly, there is a high density of 21-mers near the beginning of the CGA repeat region, consistent with previous reports that decoding of CGA codons is slow. Additionally, 21 and 28-mer peaks appear approximately one footprint length upstream of the CGA stall, extending backwards in a repeating pattern approximately every ribosome length as confirmed by autocorrelation analysis (Figure S5A, S5C top row).
Disome profiling also reveals nuclease-resistant collided ribosome footprints accumulating in this region upstream of the CGA codons; and, autocorrelation analysis reveals that the pattern of “stacked” disomes is periodic with peaks at approximately 30 nucleotide intervals (Figure 5B, center; Figure 5C). In this analysis, disome RPM peaks on the minCGA reporter are dramatically increased (∼10-fold) relative to the minOPT reporter, indicative of an accumulation of collided ribosomes (compare scales of minOPT and minCGA panels in Figure 5B). These data together suggest that a key signal for the recruitment of Syh1 and the NGD machinery is the collided ribosome.
To further explore a potential ribosomal basis for the activity of NGD factors on stalling reporter levels, we performed ribosome profiling on the minCGA reporter in hel2Δ and syh1Δ strains. First, as previously observed (Letzring et al. 2013), deletion of HEL2 increased ribosome read-through past the stall region relative to the wild-type strain (Figure S5B). Additionally, we noted from the disome profiling that the ordered periodicity in footprint distribution that is present in the WT strain is diminished in the hel2Δ strain as revealed by an autocorrelation analysis (Figures 5C, S5D). The same diminished periodicity is observed in monosomes in the hel2Δ strain, whereas the ribosome periodicity is generally maintained in the syh1Δ strain (Figure S5C).
In contrast to the minCGA reporter, the minNONOPT reporter contains non-optimal codons distributed throughout the ORF; accordingly, monosome footprints are relatively evenly distributed across the ORF (Figure 5A, right panel). Disome footprints are distributed across the ORF as well, suggestive of stochastic short-lived collisions where the ribosome density does not accumulate. To allow for accurate comparison, we performed a more detailed analysis of 21-mer/28-mer ratios for the reporter after excluding the identical oligonucleotide probe region for the two reporters as well as regions around the start and stop codons that can be sensitive to library preparation and variable within ribosome profiling data sets (O’Connor, Andreev, and Baranov 2016). Importantly, we normalized these data from the reporter ORF to global ORF 21-mer/28-mer ratios within each dataset to account for differing RNase digestion efficiencies in the library preparations. Our data reveal an increased 21-mer/28-mer ratio for the minNONOPT relative to the minOPT reporter; as 21-mer RPFs report on empty A sites on the elongating ribosomes, these data are consistent with an enrichment of ribosomes collectively undergoing slow decoding of tRNAs during elongation on non-optimal codons (Figure 5D). These data suggest that a key signal for the recruitment of the COMD machinery (including Not5) is the accumulation of slowly decoding ribosomes.
Given the abundance of 21-mer RPFs on the minNONOPT reporter and the strong decay activity of Not5 on this reporter, we wondered whether the activity of Not5 to reduce minCGA reporter half-life (Figure 4B) could be similarly explained by accumulation of 21-mer RPFs. Interestingly, a deeper analysis of the distribution of multiply aligned monosome footprint reads within the CGA repeat region of the minCGA reporter revealed that the 21mer/28mer read ratio is greatly enriched compared to the ratio on all genes (Figure 5E). We suggest that this preponderance of slow (21mer RPFs) elongating monosomes in the CGA repeat region might explain the partial sensitivity of the NGD reporter to Not5-mediated destabilization. Since Not5 recognizes ribosomes with open A and E sites, it may be able to bind those ribosomes that continue translating within the CGA12 region to elicit decay of the reporter.
Taken together, these experiments provide compelling data to rationalize the differing responses of our reporters to various gene knockouts. The accumulation of distinct ribosome signals—either nuclease-resistant collided disomes or slowly decoding ribosomes with empty A sites—acts as a strong determinant for which factors will lead to mRNA destabilization; Syh1 and Hel2 respond to the accumulation of terminally stalled, collided ribosomes, while Not5 responds to slowly decoding ribosomes with open A and E sites.
Discussion
In this study, we use carefully designed reporter mRNAs to study translation-coupled mRNA decay pathways in S. cerevisiae. Using R-SGA screening with a reporter mRNA containing iterated CGA codons, we identified and validated a set of genes that contribute to no-go decay (NGD). Subsequent analysis allowed us to compare the mechanisms of the pathways that regulate decay of mRNAs with either highly problematic (NGD) or slowly decoded sequences (COMD). We find that NGD is driven primarily by the actions of the GIGYF1/2-homologous protein Syh1; in contrast, Syh1 has no discernible impact on the stability of non-optimal mRNA sequences. We show that the previously defined COMD factor Not5 contributes modestly to decay of the NGD reporter and very strongly to decay of non-optimally coded mRNAs. Finally, we connect these distinct molecular decay profiles with ribosome states using ribosome profiling, showing that colliding ribosomes (disomes) are the key trigger for Syh1/Hel2 function while slow ribosomes (monosomes with empty A and E sites) are the key trigger for Not5 function.
Our assays using the NGD reporters reveal the interplay between Hel2, its dependent NGD factors, and Syh1 in responding to ribosome collisions. First, we found that deletion of SYH1 led to modest stabilization of NGD reporter mRNAs and deletion of CUE2 and SYH1 together led to very potent stabilization (Figure 2C). In light of previous work, we interpret the mRNA stability data as follows: 1) in the wild-type strain, the NGD reporters are stabilized because colliding ribosomes lead to the recruitment of Syh1 and elicit mRNA destabilization through Xrn1 while Hel2-mediated ubiquitination triggers ribosomal clearance by Slh1; 2) in the syh1Δ strain, NGD reporter levels are somewhat rescued because recruitment of Xrn1 is impaired, but Cue2-mediated endonucleolytic decay plays a larger role as Slh1 becomes overwhelmed; and 3) in the hel2Δsyh1Δ and cue2Δsyh1Δ strains, NGD reporters are strongly stabilized because both exonucleolytic NGD through Xrn1 and endonucleolytic NGD through Cue2 are inactivated. This model gives new context to the pathways that respond to elongation stalls; it emphasizes the essentiality of Syh1 for Xrn1-mediated NGD and supports our previously discovered mechanism in which the activity of Hel2 primarily triggers RQT-mediated ribosome rescue through Slh1, resorting to Cue2-dependent endonucleolytic cleavage only when other mechanisms to resolve collisions are overwhelmed. The synergistic activities of Syh1-assisted, Xrn1-mediated decay and Hel2-assisted, Cue2-mediated decay form the basis of a robust cellular system for targeting problematic mRNAs for destruction (Figure 6).
This interplay was further explored in our Ribo-seq experiments. For the minCGA reporter, we observed an ordered, periodic pattern of monosome and disome footprints upstream of the CGA region in the WT strain, and this pattern was substantially disrupted in the hel2Δ strain. Furthermore, we observe a larger proportion of monosome footprints downstream of the CGA region in the hel2Δ strains. Given the role of Hel2 in recognizing collided ribosome and promoting clearance by RQT, we interpret these data to mean that the activity of Hel2 on collided ribosomes stabilizes the structure and triggers clearance by Slh1 (Meydan and Guydosh 2020), thus preventing ribosome build-up in the downstream region. This stands in contrast to the maintenance of wild-type-like periodicity when SYH1 is knocked out (Figure S5C).
The next set of questions focused on how Syh1 is recruited to problematic mRNAs and how it triggers translational repression or mRNA decay. Recent work in mammalian cells investigating the mechanism of recruitment of GIGYF2 to NGD-targeted mRNAs yielded two competing models: one in which ZNF598 (a mammalian HEL2 homolog) acts to recruit GIGYF2 (Hickey et al. 2020) and one in which EDF1 acts to recruit GIGYF2 (Sinha et al. 2020; Juszkiewicz et al. 2020). We tested both models. First, we observed strong stabilization of the CGA reporter mRNA levels when SYH1 is deleted in a hel2Δ background (comparing hel2Δ to hel2Δsyh1Δ) thus establishing that Hel2 function is not necessary for Syh1 function in yeast. Second, we observed the same strong repression of CGA reporter mRNA levels in a wild-type strain and a knockout of the yeast EDF1 homolog MBF1 (a result that differs from the rescuing effect of SYH1 knockout), suggesting that Syh1 function is not dependent on Mbf1. Although Mbf1 binds collided disomes in yeast (Sinha et al. 2020; Pochopien et al. 2021), these data suggest that Mbf1 is not necessary for NGD in yeast. One possibility raised by bioID mass spectrometry (Opitz et al. 2017) is that Syh1 interacts directly with Asc1, a ribosomal protein known to be important for NGD in yeast (Kuroha et al. 2010; Letzring et al. 2013; Brandman et al. 2012), and a top effector of NGD from our genetic screen (Figure 1A). It is possible that Syh1 detects collisions by direct association with collided disomes.
Regarding the downstream effect of Syh1 to activate mRNA decay, we examined a role for the protein Eap1 (a potential functional homolog of the known GIGYF1/2-interacting factor eIF4E2 in mammals), since a direct homolog of eIF4E2 is lacking in yeast. However, deletion of EAP1 had no discernible effect on our CGA reporter (Figure S2E) suggesting that Eap1 does not function as a bridging factor between Syh1 and Xrn1-mediated mRNA decay. Another candidate effector protein for Syh1 function is Dhh1, whose mammalian homolog DDX6 interacts with GIGYF1/2 to facilitate translational repression (Peter et al. 2019; Weber et al. 2020). In yeast, however, there is only scarce evidence for a Dhh1-Syh1/Smy2 complex (Ergüden 2019) and the conserved DDX6 binding motif of GIGYF1/2 is absent in Syh1/Smy2 (Figure S2F). These data together raise the interesting possibility that Syh1 has a distinct mechanism of regulation in yeast that involves direct signaling of mRNA decay independent of translational repression. We note that the strong mRNA decay phenotype associated with Syh1 function and the NGD reporters in yeast (here and in Hickey, et al. 2020) is distinct from the translational repression phenotype associated with GIGYF2:4E2 function in mammalian systems (Morita et al. 2012; Peter et al. 2019).
Though NGD and COMD both converge on Xrn1-mediated decay of mRNAs, these processes are thought to be triggered distinctly (Radhakrishnan et al. 2016). What has not been clear is whether ribosomes translating highly non-optimal CGA repeats (that efficiently trigger NGD) may also be recognized by Not5. Conversely, the possibility remained open that highly non-optimal reporter mRNAs cause ribosome collisions that activate NGD in addition to COMD.
We observe that the loss of NOT5 modestly stabilizes the minCGA reporter, suggesting that the non-optimal decay machinery can respond to slow ribosomes even at sites undergoing NGD. For this reporter, accumulated ribosome footprints from within the CGA-repeat region are short (21-mers), reflect ribosomes with unoccupied A sites as they struggle to decode this problematic sequence (D’Orazio et al. 2019). 21-mer RPFs are also enriched in the minNONOPT reporter. These short footprint reads on both reporters represent very slow ribosomes with open A and E sites that are recognized by Not5, leading to recruitment of the Ccr4-Not complex. This observation rationalizes the modest stabilization of the minCGA reporter and the strong stabilization of the minNONOPT reporter in the not5Δ strain. In contrast, we saw no half-life increase of the minNONOPT reporter in the syh1Δ, hel2Δ, or hel2Δsyh1Δ strains, arguing that these abundant non-optimal codons do not induce ribosomal collisions that trigger NGD, consistent with a relatively even distribution and low abundance of disome peaks in the ribosome profiling data. These data provide strong evidence that non-optimal sequences are recognized distinctly from NGD promoting sequences.
Our study provides strong evidence for generally non-overlapping targets and mechanisms of NGD and COMD. While the NGD machinery, under the control of Syh1 and Hel2, responds to specific defects in elongation due to stalled and collided ribosomes, the COMD machinery, under the control of Not5, surveys the pool of translating ribosomes for mRNAs on which there is overall inefficient translation. We speculate that the COMD pathway is a general one that regulates overall mRNA stability, independent of ribosome dysfunction, while the NGD pathway evolved to deal with more acute environmental disturbances such as UV or oxidative damage (Yan et al. 2019; C. C.-C. Wu et al. 2020). Future studies will better characterize the molecular mechanisms of these pathways and will provide new foundations for an understanding of the homeostasis of cellular translation and mRNA decay.
Materials and Methods
Reporter cloning & reporter strain generation
Plasmids for OPT (pKD065), NONOPT (pKD064), and CGA (pKD080) reporters were cloned as described in D’Orazio, et al. 2019. To generate stable, genomically integrated strains containing these reporters, 0.5-2 µg of plasmid was digested using StuI to produce an insertion cassette containing the RFP and GFP reporters plus a MET17 gene for selection, all flanked by homology arms to the endogenous ADE2 locus. Strains were then transformed by lithium acetate transformation to replace the ADE2 gene as described below, with the difference of being plated directly onto selective media after transformation rather than a nonselective recovery plate. The minimal reporters minOPT and minNONOPT (plasmids pJC867 and pJC857, respectively) were a generous gift from Jeff Coller and cloned as described in Rhadakrishnan, et al. 2016. The CGA repeat stretch was introduced into the pJC867 plasmid by first isolating the plasmid backbone via digestion with PacI and AscI. Two PCR fragments making up the HIS3 ORF were generated, one containing the first portion of the HIS3 ORF and a the CGA repeat region in a primer overhang (primers AV_his3CGAupstr_fwd and AV_his3CGAupstr_rev, see table of oligos), the other containing the downstream HIS3 sequence and stop codon (primers AV_his3CGAdwnstrm_fw and AV_his3CGAdwnstrm_rv, see table of oligos). Both fragments were amplified off of pJC867 and inserted into the linearized pJC867 backbone using NEB Gibson Assembly Master Mix resulting in pAV_minCGA plasmid. Since these plasmids contained a URA3 selectable marker, reporters were introduced into various backgrounds by transformation of 0.5-1 µg plasmid as described below and all subsequent culture was performed in SC-URA media (plus additives appropriate to experiment) to retain the plasmid.
R-SGA screening
Screening procedure
Screening was performed as described in D’Orazio et al, 2019. Briefly, OPT, CGA, NONOPT, and AAA reporters were inserted into the Yeast Knockout Collection (Giaever et al. 2002) by mating and four replicate colonies were grown for each strain. Incubation times were increased during this process by 50-75% to account for decreased mating efficiency in our query strains. Cells were selected on appropriate media, then plated on 2% GAL/RAF media for expression and analyzed by imaging on a Typhoon FLA9500 imager.
Screen data analysis
Data was analyzed as previously described in D’Orazio et al. 2019 and D’Orazio et al. 2021. OPT screen data used for normalization is the same data previously published in D’Orazio et al. 2021. Briefly, median GFP and RFP values were extracted from colony images using specialized software (Saeed et al. 2003; Wagih et al. 2013) and outliers were excluded (border colonies and those <1500 or >6000 pixels). Average GFP and RFP values from all colonies were then converted to log2(GFP/RFP) ratios and LOESS normalized on each plate. Z-scores were calculated on a per-plate basis. Z-scores for volcano plots were calculated without prior LOESS normalization.
Screen validation
The 170 Yeast Knockout Collection strains with integrated reporters that showed the greatest change in GFP/RFP for the CGA reporter (−2 > CGA Z-score > 2) and lowest change for the OPT reporter (−2 < OPT Z-score < 2) relative to a his3Δ control strain were selected and grown to saturation overnight in YP + 2% galactose + 2% raffinose media in deep-well 96-well plates. Cultures were diluted in triplicate to approximately OD600 0.1 in deep-well 96-well plates and grown to approximately OD600 0.4-0.6. A 10 µL aliquot of culture was then added to 190 µL PBS and flow cytometry was performed as described below in a Guava EasyCyte HT flow cytometer. P-values were calculated by Fishers T-test.
Yeast strain generation, culture & harvesting
Knockout strain generation
Knockout strains were created using the BY4741 (MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0) background as wild-type. DNA fragments containing 40-70 nt homology arms to the gene of interest were amplified by PCR using MX cassette plasmids as template (McCusker 2017) and purified using a Zymo DNA Clean & Concentrator-5 kit. Yeast were then transformed using high-efficiency lithium acetate transformation (Gietz and Schiestl 2007). Briefly, strains to be transformed were grown to saturation at 30 °C overnight in an appropriate medium (typically YPD, YPAD, or SC-Ura), then diluted to OD600 0.2 in 5 mL media. Meanwhile, the transformation mixture was prepared, consisting of 33% PEG 3350, 100 mM LiAc, 0.28 mg/mL boiled salmon sperm DNA, and 1-5 µg PCR product. When cultures reached OD600 0.4-0.6, they were harvested by centrifugation (3000xg, 5 min) and resuspended in the transformation mixture. Transformation mixtures were incubated at 42 °C (or 30 °C for not5Δ strains) on a thermomixer for 40-60 minutes, then centrifuged briefly to collect a yeast pellet, discarding the supernatant. Finally, yeast were resuspended in 200 µL water (or media for not5Δ strains), plated on an appropriate nonselective agar medium and incubated at 30 °C. Transformants were then streaked to single colonies on a fresh plate, and these colonies were tested for MX cassette insertion by PCR using Phire Plant Direct PCR Master Mix. Confirmed strains were later maintained as patches on selective agar medium.
Growth conditions
Unless noted otherwise, yeast for steady-state reporter expression measurements (by flow cytometry or northern blot) and ribosome profiling were grown to saturation in an appropriate medium lacking glucose and containing 2% galactose and 2% raffinose. Cells were then diluted to OD600 0.1 and grown to OD600 0.4-0.65 before being harvested according to the requirements of the particular assay to be performed.
Flow cytometry
Cell lines to be analyzed with biological replicates were streaked to single colonies and three individual colonies were selected for outgrowth and analysis. Cells were grown in liquid culture as described above, then 500 µL of cell culture was transferred to a microcentrifuge tube and pelleted by centrifugation. Cells were washed once with PBS and then resuspended in 500 µL PBS. Flow cytometry was carried out using either a Guava EasyCyte or EasyCyte HT instrument, collecting >5000 events. Cellular debris and dead cells were excluded on the basis of forward and side scatter, and geometric means of per-cell GFP/RFP fluorescence distributions were used to calculate GFP/RFP for each replicate (Fig 1, 2A-B, 3) or GFP/RFP ratios were calculated on an individual cell basis for plotting of distributions (Fig S2D-E). For steady state measurements of OPT and CGA reporters in Figs 1C and 2A GFP/RFP ratios were further normalized to the mean of WT OPT or WT CGA replicates, respectively, to place them on a similar scale to northern blotting measurements.
Galactose shutoff RNA half-life assay
Biological replicates of individual cell lines were grown and diluted into 200 mL cultures in SC-URA + 2% Gal + 2% Raf media as described above. The dhh1Δ and not5Δ strains were typically slow growing and required longer incubations at 30 °C to reach saturation before dilution. When cultures reached OD600 0.4-0.6, they were split into four 50 mL conical tube and pelleted by centrifugation (3000xg, 5 min). Cell pellets were resuspended in 15 mL total prewarmed SC-URA media without added sugar to wash out residual galactose and raffinose and pelleted again by centrifugation in a single 50 mL conical tube. Pellets were resuspended in 10 mL prewarmed SC-URA without added sugar and transferred to a 125 mL beveled flask in a shaking 30 °C incubator. Zero timepoints were taken by removing a 1 mL aliquot of culture, quickly transferring to a microcentrifuge tube and pelleting cells by a snap spin to 4000xg. Supernatant was decanted and tubes were dropped into liquid nitrogen. To initiate GAL promotor shutoff, 1 mL 40% glucose was added to the 9 mL remaining culture to a final concentration of 4% and a timer was started. Subsequent timepoint samples were taken in a similar manner to the zero timepoint, with the time for each sample recorded at the moment it was dropped into liquid nitrogen. All samples were stored at −80 °C. Downstream RNA extraction and northern blotting proceeded as described below.
Steady state reporter cell harvesting for northern blot
Cells were grown as described above in 10-15 mL. At OD600 0.4-0.6 cultures were pelleted at 4 °C by centrifugation in a 14 mL culture tube, resuspended in 1 mL PBS (or residual growth media), and transferred to a microcentrifuge tube. Cells were pelleted again by centrifugation at 4 °C and supernatant was decanted. Tubes were dropped into liquid nitrogen and stored at −80 °C until RNA extraction.
Northern blotting
RNA extraction
RNA was extracted from frozen cell pellets by hot acid phenol/chloroform extraction. Aliquots of 325 µL acid phenol, pH 4.5 were heated to 65 °C in microcentrifuge tubes on a thermomixer. Cell pellets were retrieved from −80 °C storage and placed on dry ice. Working quickly, individual cell pellets were resuspended in 300-320 µL lysis buffer (8.4 mM EDTA, 60 mM NaOAc pH 5.5, 1.2% SDS) by vortexing just until pellet was fully resuspended. One aliquot of preheated phenol was immediately added to the resuspended pellet and sample was placed onto a thermomixer to minimize time between pellet resuspension and cell lysis. This procedure was repeated for samples being processed in parallel, with each sample shaking at the highest setting on the thermomixer for at least 15 minutes. Tubes were then placed in a dry ice-ethanol bath for ∼30 s to help precipitate residual SDS and centrifuged at top speed for 3 minutes. The top aqueous layer was placed in a new tube containing 300 µL room temperature acid phenol. Samples were vortexed several times for a total of 5 minutes, then centrifuged again at max speed for 30 s. The top aqueous layer was transferred to a tube containing 300 mL room temperature chloroform, vortexed several times for a total of 5 minutes, and centrifuged at max speed for 3 minutes. The aqueous phase was then transferred to a tube containing 30 µL 3.5 M NaOAc, pH 5.5. During each step of this process, particular care was taken to avoid transferring any of the organic phase or precipitate at the interface. To each RNA-NaOAc solution, 350 µL of isopropanol was added and mixed well. Tubes were placed on dry ice for at least 30 minutes, or stored at −80 °C overnight. Samples were spun at top speed in a microcentrifuge for 30 minutes and the supernatant was aspirated, taking care not to disturb the RNA pellet. Samples were centrifuged again at max speed for 5 minutes and any remaining supernatant was carefully removed with a 10 µL micropipette. To each RNA pellet, 30 µL of nuclease-free water was added and samples were incubated at 37 °C for 5 minutes on a thermomixer with gentle shaking to facilitate pellet dissolution. Tubes were then moved to ice and pipetted by hand to ensure full pellet resuspension. Finally, RNA concentrations were measured by a nanodrop spectrophotometer and samples were either used immediately for northern blotting or stored at −80 °C for subsequent use.
Gel and RNA preparation
A 1.2% agarose formaldehyde gel was prepared by mixing a final concentration of 1x MOPS electrophoresis buffer, 2.4 g electrophoresis-grade agarose and water to a final volume of 192 mL in a glass 500 mL beaker. This solution was heated in a microwave to boiling and agarose dissolution, mixed, then cooled to approximately 65 °C, placing an insulating material like paper towel beneath to promote even cooling of the solution. Particular care was taken not to allow the agarose to cool further than this before formaldehyde addition, as pieces of unevenly cooled agarose can alter RNA mobility across the gel. When initial cooling was complete, 8 mL 37% formaldehyde and 8 µL ethidium bromide were added and mixed well by swirling. Gel was poured into a mold and allowed to cool fully, then submerged in formaldehyde gel running buffer (1x MOPS buffer, 1.67% formaldehyde). Meanwhile, RNA samples were prepared by aliquoting an equal mass of total of RNA (typically 10 µg) into microcentrifuge tubes on ice containing an appropriate amount of 5x RNA loading buffer (bromophenol blue, 4 mM EDTA, 2.66% formaldehyde, 20% glycerol, 30% formamide, 4x MOPS buffer).
Gel running and transfer to membrane
RNA samples were boiled at 95 °C for 8 minutes, then cooled to room temp, spun briefly and loaded onto the gel. Gel was run at 100 V for ∼2.5 hrs. Gels were imaged on a Typhoon imager to assess RNA quality, then transferred to a Amersham Hybond N+ charged nitrocellulose membrane by a BioRad Model 785 Vacuum Blotter following the manufacturer’s instructions for transferring RNA, with the alterations of prewetting the membrane with 10x SSC only and maintaining vacuum between 10-15 inHg. Transfer proceeded for 2 hours. Following transfer, the membrane was carefully removed from the vacuum blotter and placed face up on paper towel for UV crosslinking in a Stratagene UV Stratalinker 2400 on the automatic setting (120 mJ) three times.
Oligonucleotide probe radiolabeling and hybridization
After crosslinking, the membrane was placed in a glass hybridization bottle with the RNA-side facing away from the glass. Approximately 15 mL Sigma Perfecthyb Plus Hybridization Buffer was added to the bottle and it was placed in a hybridization oven to prewarm for 30 min at 42 °C. Meanwhile, the appropriate oligonucleotide probe was enzymatically radiolabeled with the final reaction concentrations 1 µM oligonucleotide probe, 1x NEB T4 PNK buffer, 3-6 µL Perkin Elmer gamma-32P-ATP, 25 units NEB T4 PNK in a 50 µL reaction volume. This reaction was incubated at 37 °C for 1 hr, then the probe was purified using Cytiva Microspin G-50 columns according to the manufacturer’s instructions. The entire volume of probe was then added directly to the prewarmed hybridization solution in the hybridization bottle. Membrane and radiolabeled probe were incubated at 42 °C with rotation overnight. The radioactive hybridization solution was discarded and the membrane was washed three times for 20 minutes each with ∼15 mL low-stringency wash buffer (0.1% SDS, 2x SSC) at 30 °C. The membrane was placed between transparency film or sheets of plastic wrap and secured into a phosphor storage screen cassette. A blanked phosphor storage screen was exposed to the radioactive membrane long enough to produce adequate exposure (typically overnight) and imaged as described below. To strip hybridized probe off of the membrane, boiling high-stringency wash buffer (0.1% SDS, 0.2x SSC) was poured on the membrane in a hybridization bottle, incubated for 10 minutes at 80 °C, then discarded. The stripping procedure was repeated for a total of two washes, then secondary probing was performed. For experiments with the OPT, CGA, and NONOPT reporters, an oligonucleotide probe for GFP was used as the primary probe and a probe for RFP as the secondary probe. For experiments with minOPT, minCGA, and minNONOPT reporters, a probe for HIS3 was used as the primary probe and a probe for the endogenous yeast 7S RNA SCR1 was used as the secondary probe.
Phosphor imaging, northern quantification, and half-life calculation
Phosphor storage screens were scanned with a typhoon imager and bands were quantified with ImageQuant TL v8.1 software using rolling ball background subtraction. For each blot, the intensity of the primary probe band was normalized to the intensity of the secondary probe band (GFP/RFP for OPT, CGA, and NONOPT; HIS3/SCR1 for minOPT, minCGA, minNONOPT). For galactose shutoff experiments, intensities and timepoints for three or four replicates were fit to a single-exponential decay by least-squares fitting to estimate reporter RNA half-lives. For steady state reporter experiments, reporter/control ratios in Figures 1D and 2B were further normalized to WT OPT and WT CGA within each replicate set, respectively, to correct for any variation between individual blots.
Ribosome profiling
Sample preparation
Ribosome profiling was carried out based on previously published protocols (McGlincy and Ingolia 2017; Guydosh and Green 2014; C. C.-C. Wu et al. 2019).
Culture and ribosome RNA isolation
Cultures were grown to saturation in appropriate media as described above and diluted to OD600 0.1 in 1 L culture. When cells reached OD600 0.4-0.6, cells were harvested by vacuum filtration and pellets were frozen in liquid nitrogen. A portion of each pellet was ground in a SPEX SamplePrep 6870 Freezer/Mill (8 cycles, 10 hz, 1 min run, 1 min cool) with 1 mL pre-frozen lysis buffer (20 mM Tris pH 8, 140 mM KCl, 5 mM MgCl2, 1% Triton X-100, 0.1 mg/mL cycloheximide, 0.1 mg/mL tigecycline) and thawed into 15 mL lysis buffer. Lysates were cleared by centrifugation (5 min, 3000 xg, 4 °C) and supernatants were loaded onto 3 mL sucrose cushion (20 mM Tris pH 8, 150 mM KCl, 5 mM MgCl2, 500 µM DTT, 1 M Sucrose) in a Ti70 ultracentrifuge rotor tube. Samples were centrifuged for 106 minutes at 60,000 RPM, 4 °C to pellet ribosomes. Supernatant was removed, and the ribosome pellet was rinsed once with lysis buffer excluding cycloheximide and tigecycline (drug-free lysis buffer). Pellet was resuspended by pipetting in 1 mL drug-free lysis buffer. RNA concentrations were measured by Qubit RNA High Sensitivity Assay Kit, 350 µg of RNA was added to a microcentrifuge tube, and volume was increased to at least 400 µL with drug-free lysis buffer. 5 µL Ambion RNaseI was added per 400 µL of RNA solution, and samples were incubated at 25 °C in a thermomixer shaking at 500 RPM too digest free RNA. Samples were placed on ice and 10 µL Superase•In RNase inhibitor was added and mixed to stop the RNase digestion. Sucrose gradients were prepared by a Biocomp Gradient Master (15-40% sucrose gradient containing 20 mM Tris pH 8, 150 mM KCl, 5 mM MgCl2, 500 µM DTT) in SW41 ultracentrifuge rotor tubes and RNase reactions were loaded in top of the gradients. Gradients were centrifuged at 40,000 RPM for 2.5 hr at 4 °C. Gradients were fractionated on a Biocomp Triax gradient fractionator and fractions containing monosomes and disomes were individually pooled and processed in the rest of the downstream protocol. RNA was extracted from samples by SDS-hot phenol/chloroform extraction and isopropanol precipitated with GlycoBlue as co-precipitant.
Ribosome footprint isolation and reverse transcription
RNA pellets were resuspended in 10 mM Tris pH 7.5 and RNA formamide loading dye and run on a 15% TBE-urea gel, taking care to leave empty lanes between samples to minimize cross-contamination. Monosome libraries between 15 and 35 nt (monosomes) or 40 and 70 nt (disomes) were cut out of the gel, frozen, eluted overnight in RNA extraction buffer (300 µM NaOAc pH 5.5, 1 mM EDTA pH 8, 0.25% SDS), and precipitated by isopropanol precipitation. RNA pellets were resuspended in 5 µL dephosphorylation reaction mix (7 mM Tris pH 8, 1x NEB T4 PNK buffer, 10 units Superase•In, 5 units T4 PNK) and incubated at 37 °C for 1 hr. To these reactions, 5 µL of linker ligation mixture was added (38% PEG-8000, 1x NEB T4 ligase buffer, 2 µM oBZ407_preA preadenylated linker, 100 units NEB T4 RNA ligase 2, truncated) and they were further incubated at 37 °C for 3 hrs. Reactions were cleaned up with Zymo Oligo Clean & Concentrator kit and eluted in 10 µL nuclease free water. Samples were supplemented with 1 µL of 10 µM oBZ408 and denatured at 65 °C for 5 mins, then placed on ice. To each sample, 8 µL of reverse transcription reaction was added (2.5x Protoscript II buffer, 12.5 µM DTT, 1.25 mM dNTPs, 20 units Superase•In), samples were mixed, then 1 µL (200 U) Protoscript II reverse transcriptase was added. Samples were incubated 30-60 mins at 50 °C, then RNA templates were hydrolyzed by adding 2.2 µL 1 M NaOH and incubating at 95 °C, 5 min. Samples were again purified with Zymo Oligo Clean & Concentrator kit and eluted in 5 µL nuclease free water.
Ribosomal RNA depletion
A biotinylated subtraction oligo pool from Guydosh & Green, 2014, Cell was prepared as in that publication. To each sample, 1 µL of subtraction oligo pool, 1 µL of 20x SSC, and 2 µL water was added. Oligos were annealed in a thermocycler, denaturing 90 s at 100 °C, then dropping 0.1 °C/s to 37 °C and incubating 15 mins. MyOne Streptavidin C1 magnetic beads were prepared for RNA binding per the manufacturers protocol and annealed oligo solutions were transferred to the beads. Solutions were incubated for 15 mins at 37 °C, beads were pelleted and supernatants were transferred to new tubes. Samples were cleaned up using Zymo Oligo Clean & Concentrator kit and eluted in 6 µL nuclease free water.
Final sequencing library preparation
Loading dye was added to samples, and they were run on a 10% TBE-urea gel. With the aid of marker oligos, appropriate sizes were cut out from the gel for each sample and DNA was extracted from gel slices as before with DNA extraction buffer (300 µM NaOAc pH 5.5, 1 mM EDTA pH 8, 10 mM Tris pH8). DNA was isopropanol precipitated, resuspended in 20 µL circularization reaction mix (7.75 mM Tris pH 8, 1x Epicentre CircLigase buffer, 50 µM ATP, 2.5 mM MnCl2, 50 units CircLigase), incubated at 60 °C for 2 hr and 80 °C for 10 min. Relative cDNA library abundances were tested by qPCR with BioRad iTaq Universal SYBR Green Supermix to identify an appropriate number of PCR amplification cycles for each library. PCR reactions were then performed for the determined number of cycles to introduce sequencing barcodes and amplify libraries (1x Phusion HF buffer, 200 µM dNTPs, 0.5 µM oBZ287 universal forward PCR primer, 1 µM reverse barcode PCR primer, 7.5% v/v cDNA template, 1 unit Phusion polymerase). Samples were mixed with loading dye and loaded on an 8% TBE PAGE gel. Gel was run 60 min, 200 V and each library was cut from the gel, frozen, and extracted from the gel slice as above using DNA gel extraction buffer. Libraries were isopropanol precipitated, resuspended in 6 µL 10 mM Tris pH 8, and assessed for quality and concentration using an Agilent BioAnalyzer 2100 High Sensitivity DNA assay. Libraries were pooled and sequenced at the Johns Hopkins University Genetic Resources Core Facility on an Illumina NovaSeq 6000 instrument.
Data processing
Reads from raw FASTQ files were trimmed and aligned using a custom Python script to run software from the BBtools suite (https://jgi.doe.gov/data-and-tools/bbtools/) and the STAR aligner (https://github.com/alexdobin/STAR). Subsequent analyses were performed by custom Python scripts. Briefly, reads on start codons from all genes in monosome libraries were used to calculate distances from the 5’ end of a read to the ribosomal A site (17 nt for monosomes, 50 nt for disomes). Reads per million mapped reads (RPMs) were calculated at each position of the reporter RNAs by dividing the number of A site shifted 5’ ends at a given position by the total number of reads mapped to the genome (not including those that mapped to ncRNA). Reads with lengths 19 to 26 nt were considered part of the 21-mer population and reads with lengths 27 to 35 nt were considered part of the 28-mer population. Ratios of 21-mers/28-mers were calculated by dividing read numbers of 21-mers on the reporter (excluding the FLAG tag, five codons upstream of the top codon and the common binding region of the northern blotting probe) by read numbers of 28-mers on the reporter and normalizing to the 21-mer/28-mer ratio for all genes in a sample (Fig 5E). Ratios of 21-mers/28-mers in the CGA region of the minCGA reporter were calculated by re-aligning unaligned reads allowing multimapping (STAR option --outFilterMultimapNmax 999), then excluding any reads outside the CGA region and counting each unique read only once.
Gene set enrichment analysis (GSEA)
Using data from the CGA screen ranked by per-plate Z-score without LOESS normalization, ranked GSEA was performed using the GSEApy library for Python, querying the GO Biological Process annotation (“GSEApy,” n.d.; Xie et al. 2021; Subramanian et al. 2005). The top five most enriched terms in each direction were selected for plotting. Full results are available in the supplemental files.
Affinity purification-mass spectrometry (AP-MS)
TAP-tagged Syh1 and Smy2 strains were purchased from Dharmacon and grown as described above. TAP tag purifications were performed as previously published (Amberg, Burke, and Strathern 2006) replacing NP-40 for Triton X-100 and excluding TCA precipitation. Samples were submitted to the Johns Hopkins University Mass Spectrometry and Proteomics Core facility and processed by facility personnel as follows. Samples were reduced with DTT, alkylated with iodoacetamide and FASP digested on a 30 kDa filter with 10 ng/µl trypsin in 25mM TEAB at 37° C overnight. Peptides were step-fractionated by basic reverse phase chromatography on a µ-HLB Oasis plate. Samples were loaded in 0.1% TFA, eluted with 10 mM TEAB containing 5%, 15%, 20%, 25%, or 50% acetonitrile and fractions were dried. Each fraction was reconstituted in 2% acetonitrile and 0.1% formic acid and injected for MS/MS.
Raw data produced by the core facility was analyzed by MaxQuant (Tyanova, Temu, and Cox 2016) searching against the UniProt yeast database and LFQ values for identified proteins were calculated without imputation, combining data from all fractions of each sample.
Multiple Sequence Alignment
Structure-aware multiple sequence alignment for human GIGYF1 (NCBI accession: O75420.2) , GIGYF2 (NCBI accession: Q6Y7W6.1), and yeast Syh1 (NCBI accession: NP_015220.1) and Smy2 (NCBI accession: NP_015220.1) was performed by T-Coffee Expresso (Notredame, Higgins, and Heringa 2000). An additional alignment was performed by EMBL-EBI MUSCLE (https://www.ebi.ac.uk/Tools/msa/muscle) to independently verify alignment results. T-Coffee Expresso alignment results were processed with ESPript 3.0 (Robert and Gouet 2014) and output was included as Figure S2F. Regions corresponding to the DDX6 binding motif identified in Weber et al. 2020 were shaded.
Data Availability
Ribo-seq data is available in the NCBI Gene Expression Omnibus (GEO) (https://www.ncbi.nlm.nih.gov/geo/) database with the accession GSE189404. The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE (Perez-Riverol et al. 2019) partner repository with the dataset identifier PXD030076.
Funding
Canadian Institutes of Health Research Foundation Grant FDN-159913
Grant W. Brown
National Institutes of Health (R37GM059425)
Rachel Green
National Institutes of Health (5T32GM135131-02)
Juliette Lecomte
Acknowledgements
We thank Allen R. Buskirk, Niladri Sinha, and Nicolle Rosa Mercado for careful reading of the manuscript and all Green lab members for helpful discussions throughout this study. High throughput sequencing was performed at the Johns Hopkins Genetic Resources Core Facility (RRID: SCR_018669) and the Johns Hopkins Single Cell and Transcriptomics Core. Mass spectrometry was performed by the Johns Hopkins Mass Spectrometry Core.