ATPase activity of the DEAD-box protein Dhh1 controls processing body formation

1Rapid modulation of gene expression is critical for cells to respond to environmental challenges and to initiate developmental programs. Eukaryotic cells have developed a variety of mechanisms to achieve tight regulation of gene expression. This includes post-transcriptional control of messenger RNA (mRNA) levels by the regulation of translation or by varying the rates of mRNA degradation. Many of these post-transcriptional regulatory mechanisms, including the transition from mRNA translation to storage or decay, are not well characterized.

2Cytoplasmic mRNAs are marked by a 7-methylguanosine cap at the 5’ end and by a polyA tail at the 3’ end. These modifications enable interaction with translation factors, including the cap-binding complex (eIF4F) and the polyA binding protein (Pab1) and protect the mRNA against degradation. Given their impact on both translation and mRNA decay, the status of the 5’ and 3’ ends of the mRNA, as well as the complement of proteins that bind the mRNA termini, are tightly controlled. In budding yeast, a key event for the entry of mRNAs into the degradation pathway is the removal of the polyA tail, which is predominantly accomplished by the CCR4-NOT complex. While deadenylated mRNAs can also be degraded from the 3’ end by the 10-subunit exosome complex, mRNA decay in S. cerevisiae occurs predominantly via removal of the 5’ cap by the Dcp1-Dcp2 decapping enzyme, followed by degradation by the 5’-3’ exonuclease, Xrn1.

3Under certain stress conditions, such as glucose starvation or osmotic shock, protein factors involved in mRNA turnover can assemble into larger mRNP foci, known as processing bodies (PBs). PBs are dynamic, membrane-less structures that appear to form from multivalent interactions between proteins and RNA in a liquid-liquid phase separation phenomenon. Remarkably, PBs and several other related types of mRNP granules, including stress granules, germ granules, and neuronal transport granules, form in a number of different species and cell types, and in a variety of different biological contexts, suggesting these structures are important for cellular function. There is increasing evidence that the ability to form PBs is critical for survival under various stress conditions. For example, cells unable to form PBs show a severe loss in viability in stationary phase. Furthermore, ectopic expression of highly expressed mRNAs in cells that cannot form PBs is toxic. Because of their composition, PBs are postulated to be sites of mRNA storage and/or mRNA degradation. Yet, how the cell regulates PB assembly and disassembly, and how PBs modulate gene expression, has remained elusive.

4It is likely that PB formation requires factors that can either remodel the translating mRNP complex or stimulate the formation of a decay-competent or repressed mRNP. The DEAD-box ATPase Dhh1 stimulates mRNA decay and translation repression and is thought to function at an early step in PB formation, making it a good candidate to facilitate mRNA inactivation. Similar to other DEAD-box proteins, Dhh1 possesses N- and C-terminal RecA-like domains connected by a short linker, and can bind RNA with high affinity in a sequence-independent manner through the phosphate backbone. In vitro, Dhh1 has a significantly lower ATPase activity than other well characterized DEAD-box proteins such as eIF4A or Ded1. This is likely due to intramolecular interactions between its N- and C-terminal RecA lobes that hold Dhh1 in a conformation that is not competent for ATP hydrolysis suggesting the ATPase activity of Dhh1 is stimulated by factors that can alter the conformation of its two RecA domains.

5Several recent studies have revealed that DEAD-box proteins can be stimulated or inhibited by trans-acting factors. These interacting partners appear to share a common 3D architecture, namely, the presence of a MIF4G fold – a highly alpha helical HEAT repeat-like structure found in a number of different DEAD-box-interacting proteins, including eIF4G (with eIF4A), Gle1 (with Dbp5) and CWC22 (with eIF4AIII in higher eukaryotes). Intriguingly, DDX6, the mammalian homolog of Dhh1, binds directly to CNOT1 (Not1 in S.c.), the central scaffold subunit of the CCR4-NOT deadenylase complex, through its MIF4G domain and CNOT1 binding activates the ATPase of DDX6. The binding surface between these two proteins is conserved between yeast and human suggesting that the interaction between Not1 and Dhh1 is also important for modulating the activity of Dhh1 in budding yeast.

6In this study, we examine the ATPase activity of Dhh1 in vitro and in vivo, and demonstrate that the ATPase cycle of Dhh1 is a critical regulator of PB nucleation and disassembly. Cells expressing a Dhh1 variant carrying a mutation in the DEAD motif (E195Q, or Dhh1DQAD) that disrupts ATP hydrolysis form constitutive granules with both the behavior and composition of PBs induced during glucose starvation. Using recombinant proteins, we show that Not1 stimulates the ATPase activity of yeast Dhh1, similar to its function in mammals. Disruption of the interaction between Dhh1 and Not1 in vivo leads to the formation of PBs in the absence of stress, similar to the catalytically dead Dhh1DQAD allele. Furthermore, we demonstrate that Dhh1, ATP, and RNA, are sufficient to form liquid droplets in vitro with the dynamic behavior of PBs, and that these droplets can be dissolved by addition of purified Not1. Overall, these results reveal that the ATPase activity of Dhh1 is a critical regulator of PB dynamics.

7Previously, our lab demonstrated that abrogation of the ATPase activity of Dhh1 through mutation of the conserved DEAD motif (Dhh1E195Q, henceforth Dhh1DQAD; see Supplementary file 2C for a list of all Dhh1 mutants in this study) mislocalizes Dhh1 to large Dcp2-positive foci in the absence of stress. To differentiate whether loss of Dhh1 ATPase activity triggers formation of genuine processing bodies or whether these Dhh1DQAD-induced foci are anomalous granules, we monitored the localization of several PB components – namely Dcp1, Edc3, and Xrn1 – in both DHH1 and dhh1DQADmutant cells. Similar to the PB composition in glucose starvation conditions, all three GFP-tagged proteins colocalized with Dcp2-mCherry in Dhh1DQAD-expressing cells in glucose-rich conditions (Figure 1A). In contrast, the stress granule marker Pab1 did not assemble into foci in dhh1DQAD cells (Figure 1—figure supplement 1A) demonstrating that Dhh1DQAD granules are composed of proteins found in bona fide PBs.

8Despite their identification nearly 15 years ago, the precise functional role of PBs in S. cerevisiae remains poorly understood. Therefore, it is unclear whether PB formation in cells expressing dhh1DQAD is caused by a loss or gain of Dhh1 function. If Dhh1DQAD PB formation is caused by a loss of Dhh1 function, then the presence of a wild-type copy of DHH1 should abolish constitutive granule formation. To test this, we expressed wild-type Dhh1-GFP and Dhh1DQAD-GFP in either DHH1 or dhh1∆ cells in glucose-rich conditions and observed the localization of Dhh1DQAD-GFP (Figure 1B). While Dhh1DQAD-GFP – but not Dhh1-GFP – robustly formed PBs in dhh1∆ cells, PBs were no longer present in cells expressing an additional DHH1 copy, indicating that Dhh1DQAD PB formation is a recessive phenotype, and that the presence of enzymatically active Dhh1 is sufficient to prevent PB formation.

9Given that Dhh1DQAD PB formation appeared to be due to a loss-of-function rather than a gain-of-function, one possible explanation for the constitutive formation of PBs could be a block in mRNA decay in dhh1DQADcells, similar to dcp1∆ or xrn1∆ cells. In order to directly interrogate whether loss of the ATPase activity of Dhh1 disrupted mRNA turnover, we tested the functionality of Dhh1DQAD in mRNA decay using a previously established tether-based functional assay. We and others have observed that tethering Dhh1 to a reporter mRNA using the bacteriophage PP7 or MS2 systems is sufficient to stimulate the decay of a tethered mRNA. We expressed wild-type Dhh1, Dhh1DQAD, or GFP as a PP7 coat protein (PP7CP) fusion protein in dhh1∆ cells containing a single stem loop (PP7L) engineered into the 3’UTR of the FBA1 gene (Figure 1C) and assessed steady state mRNA levels by qPCR. As expected, tethering Dhh1-PP7CP to FBA1 mRNA caused an 80% reduction of FBA1 mRNA levels compared with GFP-tethered mRNA. In comparison, tethering Dhh1DQAD showed a partial attenuation of mRNA decay, with FBA1 levels decreasing by 55%, (Figure 1C), indicating that Dhh1DQAD is capable of stimulating mRNA decay.

10While tethering Dhh1DQAD to an mRNA demonstrated that this variant can function in mRNA decay, it does not address whether Dhh1DQAD PBs can degrade mRNAs. Therefore, we performed single molecule mRNA fluorescence in situ hybridization (smFISH) to examine if Dhh1DQAD PBs show hallmarks of mRNA decay. Log-phase DHH1, dhh1DQAD, and xrn1∆ cells were shifted into glucose starvation media, and the mRNA localization of FBA1, an essential glycolytic gene, was analyzed (Figure 1D). In xrn1∆ cells, 54% of FBA1 mRNAs colocalized with a Dcp2-GFP PB marker in cells grown in glucose-rich conditions and this colocalization was further increased to 75% following glucose starvation. In contrast, only 11% of FBA1 mRNAs colocalized with PB foci in glucose-starved cells expressing wild-type DHH1, consistent with the notion that PBs are sites of mRNA decay, rather than mRNA storage. In cells expressing dhh1DQAD, FBA1 mRNA showed a modest overlap with Dcp2-GFP – around 20% in glucose-rich conditions, and 22% following glucose starvation. Similar results were obtained in smFISH experiments with mRNAs coding for GFA1, which functions in chitin biosynthesis (Figure 1E), PAT1, which is involved in mRNA decapping (Figure 1—figure supplement 1B), and the phosphoglycerate kinase PGK1 (Figure 1—figure supplement 1C) in DHH1, dhh1DQAD, and xrn1∆ cells. Our tethering experiments, together with the difference in mRNA accumulation between Dhh1DQAD PBs and PBs in xrn1∆ cells, suggest that Dhh1DQAD PB formation is likely not due to a complete failure to degrade mRNAs. However, some mRNAs show slower turnover in the presence of Dhh1DQAD. Therefore, reduced decay kinetics may cause mRNAs to persist for longer in PBs, which may in part contribute to the formation of Dhh1DQAD PBs in the absence of stress.

11Our data so far indicate that loss of ATPase activity by Dhh1 triggers formation of bona fide PBs, suggesting that Dhh1 is ATP bound in PBs. We therefore tested whether ATP binding is required for PB localization of Dhh1. Wild-type Dhh1 or a previously characterized ATP-binding mutant of Dhh1 (Dhh1F66R, Q73A, henceforth Dhh1Q-motif) were co-expressed along with Dcp2-mCherry and localization was monitored in glucose-rich or glucose starvation conditions. Dhh1Q-motif showed a strong defect in PB formation (Figure 2A), consistent with prior observations, and also as evidenced by a reduction in PB localization of Dcp2, Xrn1, Dcp1, and Edc3 (Figure 2—figure supplement 1A–C) demonstrating that ATP binding by Dhh1 is required for robust PB formation. We also tested the functionality of Dhh1Q-motif using our tethering assay, and saw that Dhh1Q-motif did not show any obvious defects in mRNA decay when tethered to FBA1 mRNA (Figure 2—figure supplement 2A).

12How does the catalytic activity of Dhh1 contribute to PB formation? Given that ATP-bound Dhh1 binds mRNA in a sequence-independent manner with nanomolar affinity, one plausible model is that Dhh1DQAD binds to mRNA, but is unable to dissociate from it in the absence of ATP hydrolysis, ultimately leading to constitutive PB formation. To test this possibility, we generated an RNA binding mutant of Dhh1, Dhh13X-RNA, with alanine substitutions at three residues in the C-terminal RecA domain that are important for RNA binding (R322A, S340A, R370A). Wild-type Dhh1, Dhh1DQAD, Dhh13X-RNA, and a Dhh1DQAD/3X-RNA-GFP double mutant were co-expressed with Dcp2-mCherry in glucose-rich conditions and PB formation was monitored. While Dhh1DQAD cells formed PBs as expected (Figure 2B, left panel), combining ATPase-dead and RNA-binding mutations in cis in the Dhh1DQAD/3X-RNA mutant abolished constitutive PB formation. In addition, both Dhh13X-RNA and Dhh1DQAD/3X-RNA mutants showed a strong reduction of PB formation in glucose starvation conditions (Figure 2B, right panel). In addition, several other PB components showed strong defects in PB localization in cells expressing dhh13X-RNA (Figure 2—figure supplement 1A–C). Notably, all Dhh1 mutant proteins were expressed to similar levels in these experiments (Figure 2—figure supplement 2C). Next, we examined whether disruptions in RNA binding by Dhh1 also affected Dhh1 function in mRNA decay using our tethering assay (Figure 2—figure supplement 2B). Dhh13X-RNA caused only a ~30% reduction in tethered FBA1 mRNA levels, demonstrating that disruption of the mRNA decay activity of Dhh1 per se is not sufficient to trigger PB formation. Overall, we conclude that Dhh1 in its ATP-bound state promotes PB formation and that PB assembly requires RNA binding by Dhh1.

13The requirements of ATP and RNA binding by Dhh1 for robust PB formation would predict that deletion of DHH1 should also cause a reduction in PB formation. However, previous reports suggested that dhh1∆ cells did not show strong defects in PB assembly. To carefully assess the effects of the deletion of DHH1 on PB formation, we utilized the Diatrack particle tracking software which allows for an unbiased, automated, and accurate quantitation of foci formation (see Materials and methods). Analysis of greater than 1000 cells per experiment revealed a nearly 80% reduction of the Dcp2-mCherry foci number per cell in dhh1∆ compared to wild-type cells during glucose starvation (Figure 2B). Together, our results demonstrate that Dhh1 is required for robust PB formation.

14Given that loss of Dhh1 ATPase activity drives constitutive PB assembly and abolishes Dhh1 recycling from PBs, we asked if Dhh1DQAD also affects the dynamic localization of other mRNA decay factors to PBs. To address this question, we performed fluorescence recovery after photobleaching (FRAP) experiments (Figure 3A). Cells expressing either wild-type Dhh1 or Dhh1DQAD were shifted to glucose-free media to allow PBs to form, and the recovery of GFP-tagged mRNA decay factors within photobleached PBs was measured over time. Consistent with our previous work, Dhh1-GFP PB fluorescence recovered to roughly 80% within 1 min, while the Dhh1DQAD-GFP signal did not, suggesting that Dhh1 ATP hydrolysis is required for Dhh1 to shuttle in and out of PBs. In contrast, the dynamics of several mRNA decay factors, namely Dcp1, Dcp2, Edc3, and Xrn1, remained unchanged in cells expressing Dhh1DQAD compared with Dhh1. The mRNA decay factors observed showed two distinct FRAP profiles: Xrn1-GFP showed dynamic PB localization in DHH1 and dhh1DQAD cells, while Dcp1, Dcp2, and Edc3 showed a static PB localization profile. The limited recovery seen for Dcp1 and Dcp2 is in agreement with previous FRAP measurements in mammalian cells, and indicates that these factors are likely resident PB proteins. Thus, with the exception of Dhh1 itself, the dynamics of all tested PB components were not significantly altered by the loss of Dhh1’s ATPase activity.

15FRAP experiments allowed characterization of Dhh1DQAD and wild-type Dhh1 PB dynamics on a sub-minute time scale. To examine the dynamicity of these granules over a longer period, we also treated cells with cycloheximide, which disrupts PB formation, likely by trapping mRNAs on polysomes and preventing their entry into PBs. Cells expressing either Dhh1-GFP or Dhh1DQAD-GFP and Dcp2-mCherry were grown to mid-log phase and shifted to glucose-free media for 30 min to allow PBs to form, and then treated with cycloheximide for up to 2 hr and PB disassembly was monitored over time (Figure 3B, Figure 3—figure supplement 1A). While Dhh1-GFP showed roughly 60% disassembly of PBs after 20 min following cycloheximide treatment versus no treatment or solvent-only (Figure 3B, Videos 1 and 2 ), Dhh1DQAD PB disassembly occurred significantly slower, with 60% disassembly occurring around 80 min after cycloheximide treatment (Figure 3B, Videos 3 and 4). Notably, 2 hr cycloheximide treatment did not adversely affect cell viability (Figure 3—figure supplement 1B). The disassembly of Dhh1DQAD PBs following cycloheximide treatment suggests that these structures, like wild-type PBs, are RNA-dependent structures. However, the slower disassembly kinetics of Dhh1DQAD PBs, coupled with the dampened recycling of Dhh1DQAD, indicates that ATPase activity of Dhh1 is critical for normal PB disassembly, for instance, by facilitating release of Dhh1 from its mRNA client.

16So far, our data reveal that the ATPase cycle of Dhh1 is a critical regulator of PB dynamics, and that Dhh1 in its ATP-bound state promotes PB formation. Interestingly, Dhh1 alone is a very poor ATPase in vitro. However, DDX6, the mammalian homolog of Dhh1, can be stimulated by CNOT1, the central scaffold subunit of the CCR4-NOT complex. Based on our data, we would therefore predict that Not1 should promote the disassembly of PBs by stimulating the ATPase cycle of Dhh1. To test this prediction, we first examined whether S. cerevisiae Not1, like its mammalian homolog, stimulated the ATPase activity of Dhh1 in vitro. We recombinantly expressed and purified full-length Dhh1 and Dhh1DQAD and performed in vitro ATPase assays to assess the enzymatic activity of Dhh1 in the presence or absence of polyU RNA and recombinant Not1MIF4G (amino acids 754–1000). Similar to previous observations, we could not detect an intrinsic ATPase activity for Dhh1 alone. Dhh1 was weakly stimulated by polyU RNA (Figure 4—figure supplement 1A), whereas addition of Not1MIF4G alone had little effect (Figure 4A). However, addition of polyU RNA and increasing concentrations of Not1MIF4G robustly stimulated the ATPase activity of Dhh1, but not Dhh1DQAD (Figure 4A). In contrast, Gle1, another MIF4G-fold protein that stimulates the activity of Dbp5, a related DEAD-box ATPase that functions in mRNA export, had no effect on Dhh1 (Figure 4—figure supplement 1B). Furthermore, Not1 was unable to stimulate the catalytic activity of Dbp5 (Figure 4—figure supplement 1B), demonstrating that Not1 specifically activates the ATPase cycle of Dhh1 in vitro.

17If Not1 also stimulates Dhh1 ATPase activity in vivo then our model would predict that disruption of the Dhh1-Not1 interaction should lead to constitutive PB formation. To interfere with the Dhh1-Not1 interaction, we generated a mutant with amino acid substitutions in conserved residues on three distinct surfaces of Dhh1 (Dhh1R55E, F62E, Q282E, N284E, R355E, henceforth Dhh15X-Not) that are predicted to affect binding to Not1, based on previous structural data. Indeed, Dhh15X-Not showed a marked reduction in Not1 binding in immunoprecipitation experiments compared with wild-type Dhh1 (Figure 4B), indicating that these amino acid residues are important for the interaction between Dhh1 and Not1. Given that ATP binding by Dhh1 is likely a prerequisite for PB formation, we also examined whether ATP binding by Dhh1 was needed for the interaction with Not1. As shown in Figure 4—figure supplement 1C, Dhh1Q-motif was defective in Not1 binding, suggesting that ATP-bound Dhh1 is needed for robust interaction with Not1.

18To examine the importance of the Dhh1-Not1 interaction in PB formation, we co-expressed GFP-tagged Dhh1, Dhh1DQAD, or Dhh15X-Not along with Dcp2-mCherry, grew cells into mid-log phase and examined Dhh1 localization. Dhh15X-Not triggered Dhh1 and Dcp2 colocalization in cytoplasmic granules in glucose-rich conditions, similar to catalytically dead Dhh1DQAD(Figure 4C). All Dhh1 mutant proteins were expressed to similar levels in these experiments (Figure 4—figure supplement 2B). Two lines of evidence suggest that these granules are indeed bona fide PBs. First, Dhh15X-Not granules contained several other known PB proteins – including Xrn1, Dcp1, and Edc3 – in both glucose-rich and glucose starvation conditions (Figure 4—figure supplement 1D–F). Second, Dhh15X-Not granule assembly required robust RNA binding activity, as a Dhh15X-Not/3X-RNA mutant showed a dramatic defect in PB formation in both glucose-rich and glucose starvation conditions (Figure 4C). Of note, the Dhh15X-Not mutant did not show a defect in mRNA decay using our tethering assay (Figure 4—figure supplement 2A), suggesting that PB formation in cells expressing Dhh15X-Not is not due to a block in mRNA degradation.

19Several other decay factors have been shown to interact with the C-terminal RecA domain of Dhh1. Thus, it was conceivable that the Dhh15X-Not mutant not only disrupted the interaction between Dhh1 and Not1, but also with additional factors, which may contribute to PB formation. We therefore generated a NOT1 allele, not19X-Dhh1 – with substitutions at conserved amino acid residues that were previously shown to mediate interaction between CNOT1 and DDX6 (F791A, N795A, K804A, E832R, N834A, Y835A, K962A, F967A, and E970A). We co-expressed Not1 or Not19X-Dhh1 along with Dhh1-GFP and Dcp2-mCherry and shifted cells into media with and without glucose to evaluate PB formation. While cells expressing wild-type Not1 showed diffuse Dhh1 and Dcp2 localization, the Not19X-Dhh1 mutant triggered colocalization of Dhh1 and Dcp2 into distinct foci in both glucose-rich and glucose starvation conditions (Figure 5). While granule induction in these cells was less pronounced than in Dhh1DQADor Dhh15X-Not cells (Figure 4C) in glucose-rich conditions, these foci contained other known PB components (Figure 5—figure supplement 1A–C), suggesting they are bona fide PBs. Additionally, Not19X-Dhh1 was expressed at wild-type Not1 levels (Figure 5—figure supplement 1D). In summary, we conclude that the ATPase cycle of Dhh1 is a critical regulator of PB formation, and that Not1 regulates the ATPase activity of Dhh1 in vivo, preventing PB formation in glucose-rich conditions.

20To better understand how the ATPase cycle of Dhh1 regulates PB formation, we attempted to reconstitute granule formation in vitro. Remarkably, recombinant Dhh1, in the presence of RNA and ATP, readily formed droplets in solution (Figure 6A and B). These droplets showed hallmarks of liquid-liquid phase separation, undergoing growth and fusion events and reversible deformation (Video 5), consistent with the reported biophysical behavior of PBs. Dhh1 droplet formation was RNA-dependent, as no droplets formed when RNA was omitted (Figure 6A), and the number and size of droplets rapidly decreased upon addition of RNase A (Figure 6C, Video 6), but not with buffer alone (Figure 6C, Video 7).

21Next, we examined the role of ATP binding by Dhh1 in droplet formation. Despite numerous attempts, we were unable to purify Dhh1Q-motif with sufficient quality for analysis. However, we successfully purified a single Q-motif point mutant, Dhh1F66R for use in our in vitro assay. While Dhh1F66R showed only a minor defect in PB localization following glucose starvation (Figure 6—figure supplement 1A), this mutant showed a dramatic loss of droplet formation in vitro (Figure 6—figure supplement 1B). Additionally, Dhh1 droplets did not form in the absence of ATP (Figure 6B), demonstrating that Dhh1 in its ATP-bound form promotes liquid droplet formation.

22Given that Not1 promotes PB disassembly in vivo by stimulating the ATPase activity of Dhh1, we next examined whether the presence of Not1 also antagonizes Dhh1 liquid droplet formation in vitro. Consistent with our in vivo data, addition of Not1MIF4G triggered the dissolution of pre-formed Dhh1 liquid droplets (Figure 6C, video 8). Furthermore, no assembly occurred when Not1MIF4G was added before polyU during the assembly reaction (Figure 6D).

23To determine whether catalytically active Dhh1 was required for droplet dissolution, we also tested the functionality of the ATPase-dead Dhh1DQAD mutant in our in vitro assay. While Dhh1DQAD formed droplets to a similar extent as wild-type Dhh1 (Figure 6E), these structures did not dissolve in the presence of Not1MIF4G, supporting the specificity of the observed effect (Figure 6E). Interestingly, the Dhh1DQAD droplets slightly increased in size and number upon Not1MIF4G addition. It is likely that the MIF4G domain of Not1, like other MIF4G domains, stabilizes a conformation of the two RecA domains which facilitates nucleotide and RNA loading, which consequently may enhance droplet formation in the absence of Dhh1's ATPase activity. Thus, while other mRNA decay factors contribute to PB formation in vivo, this demonstrates that with a minimal number of constituents, namely Dhh1, RNA, and ATP, higher-order dynamic liquid droplets can be formed in vitro. These droplets recapitulate several properties of PBs formed in vivo such as the dependence on ATP and RNA binding by Dhh1 to form (Figure 2A and B) as well as the requirement of both a functional Dhh1 ATPase and the MIF4G domain of Not1 for dissolution (Figures 4C and 5).

24The DEAD-box ATPase Dhh1 and its orthologs play a critical role in translational repression and degradation of cytoplasmic mRNAs. However, how the catalytic activity of Dhh1 contributes to its function has not been well defined. Here, we show that the ATPase activity of Dhh1 regulates the dynamics of PBs in an RNA-dependent manner. Point mutations in Dhh1 that prevent ATP hydrolysis or disrupt the interaction surface with the ATPase activator Not1 were sufficient to trigger aberrant PB formation in vivo in the absence of stress (Figure 4C, Figure 5). Furthermore, we can recapitulate this process in vitro, as Dhh1 forms dynamic liquid droplets in the presence of RNA and ATP that are dissolved upon addition of the purified MIF4G ATPase activation domain of Not1 (Figure 6).

25The central scaffold of the CCR4-NOT complex, Not1, similar to its mammalian homolog CNOT1, is shown here to be an activator of the catalytic cycle of Dhh1 in vitro. Like other known DEAD-box cofactors, Not1 possesses a MIF4G domain that is critical for stimulation of Dhh1. In the absence of Not1MIF4G and RNA, we could not detect ATPase activity of Dhh1. Both Dhh1 and DDX6 alone adopt an unusual closed conformation, characterized by extensive intramolecular contacts that are not present in other members of the DEAD-box protein family. Binding of CNOT1 causes a dramatic structural rearrangement, shifting DDX6 into an ATPase-competent state. However, even the Not1-stimulated activity of Dhh1 remains low (Figure 4A). Although the rate-limiting step in the catalytic cycle of DEAD-box ATPases often appears to be the release of substrate RNA and ADP/Pi, it is conceivable that the large conformational change that Dhh1 must undergo in order to bind both Not1 and substrate significantly contributes to the slow ATPase cycle of Dhh1.

26While mutations in conserved residues that mediate the interaction between Dhh1 and Not1 triggered constitutive PB formation, these mutants did not completely recapitulate the degree of PB formation seen in cells expressing catalytically dead Dhh1DQAD. This may perhaps be due to only a partial loss of stimulation of Dhh1 by Not1 in these mutants. Unfortunately, we were unable to purify these variants as recombinant proteins, and therefore could not determine their effect on ATPase stimulation in vitro. Alternatively, there may be additional cellular factors that modulate the catalytic cycle of Dhh1.

27In addition to demonstrating a critical role for ATP hydrolysis by Dhh1 in the regulation of PB formation, we also show that both RNA and ATP binding by Dhh1 are critical for PB assembly, consistent with prior observations. Neither Dhh1Q-motif nor Dhh13X-RNA mutant cells robustly form PBs following glucose starvation. In addition, mutations that disrupt RNA binding also prevent constitutive PB formation of catalytically dead Dhh1 (Figure 2B). Remarkably, Dhh1 forms liquid droplets in vitro that require both ATP and RNA (Figure 6), indicating that multimeric assembly of Dhh1 in its ATP-bound state with RNA may drive PB formation. Since DDX6 can oligomerize in both an RNA-dependent and RNA-independent manner, and given that both Dhh1 and DDX6 exist in molar excess over cytoplasmic mRNA, it is conceivable that an ATP-bound conformation of Dhh1 multimerizes on RNA in vivo, thereby delivering mRNAs to PBs and seeding PB assembly. Upon ATP hydrolysis, Dhh1 could then return to the cytoplasmic pool to bind and deliver the next mRNA target. However, when ATP hydrolysis is inhibited, such as in Dhh1DQAD or Dhh15X-Not-expressing cells, Dhh1 remains associated with its mRNA client, triggering the formation of constitutive PBs.

28Despite an increasing understanding of PB composition, the precise functional role of PBs remains unclear. Given the large number of mRNA decay factors present in PB assemblies, as well as the accumulation of Xrn1-protected polyG-tract-containing mRNAs, PBs were initially proposed to be sites of mRNA decay. However, several studies have shown that mRNAs can also stably localize within PBs, raising the question of whether these granules are sites of active mRNA decay or rather of mRNA storage. It should be noted, however, that in many cases mRNAs were localized to PBs using either the MS2 or PP7 coat protein system, whereby multiple stem loops are engineered into the 3’UTR of transcripts and then visualized using fluorescently tagged coat-protein fusions that recognize these stem loops. Yet, recent data indicate that these stem loop systems may inhibit mRNA decay in budding yeast, and that primarily these stem loop structures – but not the body of transcripts – persist in PBs, and cannot be degraded by Xrn1 (,;; Heinrich and Weis, unpublished). Our smFISH data are consistent with the hypothesis that active decay occurs within PBs, since none of the four tested mRNAs were enriched in wild-type and Dhh1DQAD PBs in contrast to PBs in xrn1∆ cells (Figure 1D–E, Figure 1—figure supplement 1B–C). Thus, selective delivery of mRNAs to PBs could enhance their degradation because of the high local concentration of the mRNA decay machinery in PBs. Alternatively, the sequestration of mRNA decay factors and selected mRNAs into PBs could also allow for spatial separation of translation factors from mRNA, preventing translation of messages that would be unproductive during periods of stress.

29The formation of mRNPs – including PBs – into membrane-less organelles that behave like dynamic liquid droplets has recently emerged as a common mechanism by which cells may further compartmentalize their biochemistry. Furthermore, a variety of different ATP-driven protein machines have also emerged as important regulators of mRNP granule assembly. For example, stress granule (SG) assembly and dynamics are disrupted by loss-of-function alleles in the MCM and RVB helicase complexes, while hypomorphic alleles in the chaperonin-containing T (CCT) complex form more SGs. Additionally, the AAA+ ATPase Cdc48 was also previously shown to facilitate clearance of SGs. Our data demonstrate that the DEAD-box ATPase Dhh1 is a critical regulator of PB disassembly in vivo and that liquid droplets containing Dhh1 multimers form in the presence of RNA and ATP, which can be dissolved upon induction of ATP hydrolysis in vitro. Two biochemical functions are critical for the role of Dhh1 in PB formation both in vivo and in vitro: (1) Dhh1’s affinity for RNA, which may facilitate delivery of mRNA substrates into PBs, and (2) ATP binding, and the tuning of Dhh1’s ATPase activity by factors such as Not1. These features of Dhh1 may ultimately be the critical controllers of PB formation and PB turnover.

30PBs have been extensively studied, yet the molecular mechanisms driving PB formation and disassembly are poorly understood. Our data show that the ATP- and RNA-bound form of Dhh1 promotes PB formation, while Not1 promotes PB disassembly by stimulating the ATPase activity of Dhh1 (for illustration, see model Figure 7). Still, there are several elements of the PB and Dhh1 ATPase cycle that remain unclear. For example, what leads to PB formation under specific cellular stress conditions? Is this driven by a stress-induced attenuation of translation or increase in mRNA turnover, which leads to an increased number of client mRNAs targeted to PBs? Alternatively, cellular stress may dampen the ATPase activity of Dhh1, for example by regulating the interaction between Dhh1 and Not1, thereby shifting the equilibrium towards the ATP-bound, RNA-bound Dhh1 state. This in turn would then slow down PB disassembly, causing a build up of PB structures.

31In addition, the polyA status of Dhh1-bound mRNAs targeted to PBs is also unknown. While deadenylation by CCR4-NOT was previously placed upstream of Dhh1 in the mRNA decay pathway, it is unclear whether deadenylation is a prerequisite for targeting mRNAs to PBs. Finally, with respect to the hydrolysis step, does Not1 facilitate recycling of Dhh1 from the PB by promoting the release of Dhh1 from RNA, similar to the function of other DEAD-box activators or does it regulate the interaction with other factors such as scaffold proteins of PBs? Intriguingly, the ATPase cycle of Dhh1 does not seem to influence the recycling of other PB components (Figure 3A), consistent with the idea that the regulated interaction between Dhh1 and mRNA shifts the balance between PB formation or disassembly.

32While Not1 has a well-known role as the central subunit of the major cytoplasmic deadenylase, our work defines a novel function for Not1 in PB disassembly and Dhh1 recycling, which presumably occurs at a late stage in mRNA turnover. Interestingly, there is increasing evidence that the CCR4-NOT complex functions at several other steps during gene expression outside of mRNA decay, including transcription and translation. Future work is needed to address how the activity of this multifunctional protein complex is modulated in order to regulate and coordinate multiple steps of gene expression.

33The strains used in this study are derivatives of W303 and are described in Supplementary file 1. Yeast deletion strains and C-terminal epitope tagging of ORFs was done by PCR-based homologous recombination, as previously described. Generation of bacteriophage PP7CP and PP7-loop tagging plasmids was described previously.

34Plasmids for this study are described in Supplementary file 2A. Mutations in Dhh1 were generated using a QuikChange II site-directed mutagenesis kit (Agilent Technologies, Santa Clara, CA) using Pfu Ultra or Pfu Turbo. Mutagenic oligonucleotides were designed using the Agilent Technologies primer design platform. Construction of NOT1-TAP and not19X-Dhh1-TAP integration vectors were made using NOT1 and NOT1(F791A, N795A, K804A, E832R, N834A, Y835A, K962A, F967A, and E970A)gene block fragments ordered from Integrated DNA Technologies (IDT, Coralville, IA) that were ligated into the single-integration vector pNH605 by Gibson Assembly (New England Biolabs, Ipswich, MA). Primer sequences for strain construction are listed in Supplementary file 2B.

35Sample preparation was performed as previously described. Briefly, yeast cells were inoculated overnight in synthetic media containing 2% glucose and grown to saturation. The following morning, cultures were diluted and grown to exponential growth phase (OD600 = 0.4–0.8) then collected by centrifugation and lysed in 1X phosphate-buffered saline (PBS) with 0.1% Tween-20 and protease inhibitors. Lysis was performed using a Mini-Beadbeater-96 (BioSpec Products, Inc., Bartlesville, OK) with a 5-minute cycle. The extract was clarified by centrifugation, and RNA was isolated using the RNeasy RNA isolation kit (Qiagen, Hilden, Germany). Resulting RNA samples were stored at −80°C.

36RNA was isolated as described above and quantified using a NanoDrop spectrophotometer (Thermo Fischer, Waltham, MA). cDNA was generated by reverse transcription of 1 μg of RNA using a random hexamer oligonucleotide (Invitrogen, Carlsbad, CA) and Superscript II (Invitrogen). Quantitative PCR was performed in real time using the StepOnePlus Real-Time PCR System (Applied Biosystems, Foster City, CA) and a SYBR-Green ROX qPCR Master Mix (Thermo Fischer) supplemented with gene-specific primers as reported in.

37The indicated strains were inoculated overnight in synthetic media containing 2% glucose and grown to saturation. The following morning, cultures were diluted and grown in synthetic complete media containing 2% glucose at 25°C to exponential growth phase (OD600 = 0.6-0.8), then shifted to synthetic complete media with or without glucose for 20 min and fixed for 15 min with 4% paraformaldehyde. Samples were processed for single molecule fluorescence in situ hybridization (smFISH) as described in, with the exception of spheroplasting yeast cells for 20 min using 1% 20T zymolyase. Mixtures of DNA probes coupled to CAL Fluor Red 590 (Stellaris, LGC Biosearch, Novato, CA; probes were synthesized by BioCat, Heidelberg, Germany) were used for smFISH, targeting the FBA1, GFA1, PAT1, or PGK1 open reading frame moiety (Supplementary Supplementary file 3). Microscopy was performed with an inverted epi-fluorescence microscope (Nikon Ti) equipped with a Spectra X LED light source and a Hamamatsu Flash 4.0 sCMOS camera using a PlanApo 100 x NA 1.4 oil-immersion objective and the NIS Elements software. Images were processed using FIJI software. Quantification of colocalization was performed on all planes of a 3D stack image using the Colocalization Threshold tool in Fiji. In brief, images were background-subtracted and thresholded with a defined minimum threshold set for each smFISH probe separately. Then, the Colocalization Threshold tool was applied, which highlighted the colocalization between PB and mRNA. The colocalization events and the total number of PBs were then counted manually. The percentage of colocalization was calculated by forming the ratio between the number of PBs colocalizing with mRNA and the total number of PBs.

38Immunoprecipitation experiments were performed as in. Yeast were inoculated in synthetic media containing 2% glucose and grown overnight to saturation, then diluted the following day in 1 L synthetic media and grown to exponential growth phase (OD600 = 0.4–1.0). Cells were harvested by centrifugation at 3000 x g for 10 min, then resuspended in resuspension buffer (final concentration: 20 mM HEPES-KOH, pH 7.4, 1.2% polyvinylpyrrolidone (molecular weight = 40,000), 1 mM DTT, 0.2 mM PMSF, 10 μg/mL Pepstatin A). Pellets were centrifuged at 2600 x g for 15 min at 4°C to remove extra buffer, then centrifuged again at 2600 x g for 15 min at 4°C and pellets were frozen in liquid nitrogen and stored at −80°C. Frozen yeast pellets were then lysed with a Retsch Planetary Ball Mill MM 301 (Retsch, Newtown, PA) for six cycles at 30 Hz for 3 min with cooling in liquid nitrogen between cycles. 0.5 g of lysate was then resuspended in 14 mL TBT buffer (final concentration: 20 mM HEPES-KOH, pH 7.4, 110 mM KOAc, 2 mM MgCl2, 1 mM DTT, 0.5% Triton X-100, 0.1% Tween-20, 0.2 mM PMSF, 10 μg/mL Pepstatin A, 1:5000 SuperRNasin (Ambion, Austin, TX), 1:5000 Antifoam B (Sigma Aldrich, St. Louis, MO). Lysate was clarified through 2.7 µm and 1.6 µm GD/X Glass Microfiber syringe filters (Whatman, Maidstone, UK), and then incubated with 8 mg rabbit IgG (Sigma Aldrich)-coupled magnetic beads (Thermo Fischer, Waltham, MA) – corresponding to 400 μL bead slurry at 20 mg/μL slurry – and rotated at 4°C for 30 min. The beads were collected using a magnetic rack, washed three times with 1 mL TBT buffer, and a final wash in 1 mL of 100 mM NH4OAc, (pH = 7.4, 0.1 mM MgCl2, 0.2% Tween-20) for 5 min while rotating. Protein complexes were eluted from the beads directly in SDS-PAGE sample buffer and boiled at 95°C, and processed further for Western blot.

39For Western blot analysis, roughly 5 OD600 units of cells were harvested and treated with 5% trichloroacetic acid (TCA) and incubated at 4°C for 10 min. Acid was removed using an acetone wash, and the resulting pellet was dried 2–3 hr. Cell pellets were resuspended in 200 μL breakage buffer (final concentration: 50 mM Tris-HCl pH = 7.5, 1 mM EDTA, 2.75 mM DTT, and protease inhibitors) and disrupted using glass beads and a Mini-Beadbeater-96 (BioSpec Products, Inc. Bartlesville, OK). Samples were cooled on ice for 5 min and SDS sample buffer was added and homogenates were boiled. Proteins were resolved by 4–12% Bolt Bis-Tris SDS PAGE (Thermo Fischer, Waltham, MA), then transferred to nitrocellulose membrane (GE Life Sciences, Marlborough, MA). Membranes were blocked in PBS with 4% non-fat milk, followed by incubation with primary antibody overnight. Membranes were washed four times with PBS with 0.1% Tween-20 (PBS-T) and incubated with secondary antibody for 45 min. Membranes were then analyzed and quantified using an infrared imaging system (Odyssey; LI-COR Biosciences, Lincoln, NE). The following primary antibodies were used for detection of tagged proteins at the indicated dilutions: rabbit-anti-Dhh1 (1:5000) as described in, (Weis Lab ETH Zurich Cat# Weis_001, RRID:AB_2629458), anti-FLAG-M2 (1:2500) (Sigma-Aldrich Cat# F1804, RRID:AB_262044, St. Louis, MO), mouse-anti-HA.11 (1:2000) (Covance Research Products, Inc. Cat# MMS-101P-1000, RRID:AB_291259, Princeton, NJ) mouse-anti-GFP (1:1000) (Roche Cat# 11814460001, RRID:AB_390913), and rabbit-anti-Hxk1 (1:3000) (US Biological Cat# H2035-01, RRID:AB_2629457, Salem, MA). IRdye 680RD goat-anti-rabbit (LI-COR Biosciences Cat# 926–68071, RRID:AB_10956166) and IRdye 800 donkey-anti-mouse (LI-COR Biosciences Cat# 926–32212, RRID:AB_621847) were used as secondary antibodies.

40Samples were grown overnight in synthetic media containing 2% glucose, diluted to OD600 = 0.05 or 0.1 the following day, and grown to mid-log phase (OD600= 0.3–0.8). Cells were harvested by centrifugation and washed in ¼ volume of fresh synthetic media +/− 2% glucose, then harvested again and resuspended in 1 volume of fresh synthetic media +/− 2% glucose and grown 15 min at 30°C. Cells were then transferred onto Concanavalin A-treated MatTek dishes (MatTek Corp., Ashland, MA) and visualized at room temperature using the DeltaVision Elite Imaging System with softWoRx imaging software (GE Life Sciences, Marlborough, MA). The system was based on an Olympus 1X71 inverted microscope (Olympus, Japan), and cells were observed using a UPlanSApo 100 × 1.4 NA oil immersion objective. Single plane images were acquired using a DV Elite CMOS camera. Image processing for PB analysis was performed using Diatrack 3.5 particle tracking software as described below.

41Samples were grown as indicated in 'wide-field fluorescence microscopy' methods section and imaged using an Andor/Nikon Yokogawa spinning disk confocal microscope (Belfast, United Kingdom) with Metamorph Microscopy Automation & Image Analysis software (Molecular Devices, Sunnyvale, CA). The system was based on a NikonTE2000 with inverted microscope, and cells were observed using a PlanApo100 × 1.4 NA oil immersion objective and single plane images were captured using a Clara Interline CCD camera (Andor).

42Samples were grown overnight in synthetic media containing 2% glucose, diluted to OD600 = 0.05 or 0.1 the following day, and grown to mid-log phase (OD600= 0.3–0.8). Cells were harvested by centrifugation and washed in ¼ volume of fresh synthetic media +/− 2% glucose, then harvested again and resuspended in 1 volume of fresh synthetic media +/− 2% glucose and grown 15 min at 30°C. Cells were then transferred onto Concanavalin A-treated MatTek dishes (MatTek Corp., Ashland, MA) and visualized at room temperature. Dhh1-GFP and Dhh1DQAD-GFP photobleaching experiments were performed on a Leica SP8 Laser Scanning Confocal Microscope (Leica, Wetzlar, Germany) using Leica LAS AF SP8 software (version 3.3). The system was based on a LeicaDMI6000B inverted microscope, and cells were observed using a PlanApo 63 × 1.4 NA oil immersion CS2 objective and a conventional photomultiplier tube (PMT) detector. Dcp1-GFP, Dcp2-GFP, Edc3-GFP, and Xrn1-GFP photobleaching experiments were performed on a Andor/Nikon Yokogawa spinning disk confocal microscope with acquisition parameters as described above.

43Using the Leica SP8 Laser Scanning Confocal Microscope, selected PBs were subjected to 5–10 pulses of an argon laser at 488 nm. Images were collected from a single plane with a 2.92 nm pinhole at 500 ms intervals for 50 s post-bleach. Using the Andor/Nikon spinning disk confocal microscope, selected PBs were pulsed once for 500 ms using a Mosaic 405 nm laser (Andor) and images were collected from a single plane at 3 s intervals for 3 min post-bleach. For all experiments, PB fluorescent intensity and total cellular fluorescence intensity were quantified in ImageJ/FIJI by manual tracing. The background was determined by determining the intensity of an ROI with the same size as either the PB or the total cell. PB intensity was normalized to the total fluorescent intensity of the cell using the equation:

44Recovery curves were generated by normalization to the bleach point, and percent of fluorescent recovery values were determined by curve fitting using the equation:

45To quantify PBs, we used Diatrack 3.5 particle tracking software (; www.diatrack.org). A Matlab script transformed our foci images into a single long sequence that could be fed to Diatrack. This allowed the same image analysis parameters to be applied across all images and experiments. PBs display significant variations in appearance (size and brightness). They were identified in Diatrack based on their intensity and contrast measure. Optimal parameters were selected interactively such that false negative and false positive rates were below 3%. Occasionally, yeast vacuoles pinch the cytoplasm against the cell wall. This tends to create narrow intensity ridges in some of our fluorescence images and can trigger the default Diatrack particle detector. Thus, we used an alternative contrast measure (‘blurred 360’). This only retained particles around which the intensity decreases significantly in all directions (rather than decreasing on average only). A sample movie showing detected PB for a variety of images is provided as supplementary information (movieDetection.avi). For each image, a list of intensities corresponding to each PB in each image was exported from the software and further processed in Microsoft Excel (Microsoft Corporation, Redmond, WA). The sum of particle intensities represents a suitable measure of overall PB abundance. Alternately, the number of PBs per image may also be used. This value was divided by the number of cells in each image to deliver per-cell PB abundances. Automated counting of cells was performed as described in.

46Dhh1 (wild-type, Dhh1DQAD, Dhh1F66R) and Dbp5 were cloned into a pETMCN-based expression vector with a N-terminal 6xHis and V5 tag plus a C-terminal mCherry tag. The MIF4G domain of Not1 (residues 754–1000) was cloned into a pETMCN-based expression vector with a N-terminal 6xHis and V5 tag. Recombinant proteins were expressed in E. coli BL21 DE3 cells grown in rich medium. Cells were grown at 37°C to an OD600 of 0.6 and induced with 300 µM IPTG. Cells were then grown overnight at 18°C, harvested and resuspended in 30 mL lysis buffer (500 mM NaCl, 25 mM Tris-HCl pH 7.5, 10 mM imidazole, protease inhibitors) per cell pellet from 2 L of culture. After cell lysis by sonication, the 6xHis tagged proteins were affinity extracted with Ni2+ sepharose and further purified by size exclusion with a Superdex 200 column (Dhh1 and Dbp5, in the final storage buffer 200 mM NaCl, 25 mM Tris-HCl pH 7.5, 2 mM DTT) or Superdex 75 column (Not1MIF4G, in the final storage buffer 200 mM NaCl, 25 mM Tris-HCl pH 7.5, 2 mM DTT, 10% glycerol) (GE Life Sciences, Marlborough, MA). Gel filtration fractions were analyzed by SDS-PAGE. Clean fractions were pooled, concentrated to about 500 µM and snap frozen in small aliquots.

47ATPase assays were performed according to with the following modifications: 2 µM Dhh1 or Dbp5 was mixed with 2 µL 10x ATPase buffer (300 mM HEPES-KOH pH 7.5, 1 M NaCl, 20 mM MgCl2), Not1 or Gle1 as indicated, 4 µL 10 mg/ml polyU (unless indicated otherwise), RNase inhibitors, 13.3 µL 60% glycerol, 2.7 µL 10 mg/mL BSA, and Not1 storage buffer to compensate for volume differences, in a final volume of 36 µL. Reactions were set up in triplicate in a 96-well NUNC plate. The assay was initiated by the addition of 40 µL of a master mix containing 1x ATPase buffer, 2.5 mM ATP (from a 100 mM stock in 0.5 M HEPES-KOH pH 7.5), 1 mM DTT, 6 mM phosphoenolpyruvate, 1.2 mM NADH (from a 12 mM stock in 25 mM HEPES-KOH pH 7.5) and 125–250 units / mL PK/LDH. NADH absorption was monitored with a CLARIOstar plate reader at 340 nm in 30 s intervals for 400 cycles.

48Dhh1-mCherry (wild-type, Dhh1DQAD, or Dhh1F66R) was diluted at least tenfold to 50 µM with 1x ATPase buffer. From this solution, Dhh1 was added as indicated in Figure 6 to a 20 µL reaction (and pre-incubated with Not1, if applicable) in a 384-well microscopy plate. Reactions were filled to 5 µL with 1x ATPase buffer. A master mix was prepared with 2 µL 10x ATP reconstitution system (40 mM ATP, 40 mM MgCl2, 200 mM creatine phosphate, 70 U/mL Creatine Kinase), 1 µL HEPES-KOH pH 6.6, 1 µL BSA (10 mg/mL), 1.5 µL 1 mg/mL polyU (unless indicated otherwise), 0.2 µL RNase inhibitors and 10 µL buffer (150 mM KCl, 30 mM HEPES-KOH pH 7.4, 2 mM MgCl2) and added to the protein solutions. For the reactions not containing the creatine kinase ATP regeneration system, ATP was supplemented to a final concentration of 5 mM together with 10 mM MgCl2. Reactions were mixed, incubated at 4°C for the indicated length, and microscopy was performed with an inverted epi-fluorescence microscope (Nikon Ti) equipped with a Spectra X LED light source and a Hamamatsu Flash 4.0 sCMOS camera using a PlanApo 60 × NA 1.4 oil-immersion objective and the NIS Elements software.