Decapping complex is essential for functional P-body formation and is buffered by nuclear localization

mRNA decay is a key step in regulating the cellular proteome. Cytoplasmic mRNA is largely turned over in processing bodies (P-bodies). P-body units assemble to form P-body granules under stress conditions. How this assembly is regulated, however, remains still poorly understood. Here, we show that the translational repressor Scd6 and the decapping stimulator Edc3 act partially redundantly in P-body assembly by capturing the Dcp1/2 decapping complex and preventing it from becoming imported into the nucleus by the karyopherin ß Kap95. Nuclear Dcp1/2 does not drive mRNA decay and might be stored there as a ready releasable pool, indicating a dynamic equilibrium between cytoplasmic and nuclear Dcp1/2. Cytoplasmic Dcp1/2 is linked to Dhh1 via Edc3 and Scd6. Functional P-bodies are present at the endoplasmic reticulum where Dcp2 potentially acts to increase the local concentration of Dhh1 through interaction with Scd6 and Edc3 to drive phase separation and hence P-body formation.


Introduction
Translational attenuation is among the first lines of defense when a cell encounters stress. Ribosomes will release mRNA and most of the mRNA is captured into processing bodies (P-bodies), which form very quickly, within five minutes after stress encounter. The fast formation of P-bodies might be explained by the notion that the release of mRNAs and their capture into PBs is coordinated in two ways. First, a subset of PB components such as the 5' exonuclease, Xrn1, is associated with polysomes and second, regulators of translation such as Scp160 and Bfr1 negatively regulate PB formation (Weidner et al., 2014;Tesina et al., 2019). Moreover, PBs contain the translational repressor Scd6, which can sequester eIF4G (Nissan et al., 2010; Rajyaguru, She and Parker, 2012). P-bodies were initially thought to represent an mRNA decay compartment (Sheth and Parker, 2003). This decay requires removal of the 5' 7-mG cap by the decapping complex Dcp1/2. The decapping activity is stimulated by DEAD box helicase Dhh1/DDX6 and the RNA binding protein Pat1 (Nissan et al., 2010). Recent data, however, provide evidence that P-bodies do not only act as decay compartments but are also mRNA storage organelles and that the fate of an mRNA in PBs is dependent of the type of stressor (Wang et al., 2018;Luo et al., 2020). But not only the fate of mRNAs in PBs is stress-dependent, also the morphology of PBs can vary according to the stress. For example, under glucose starvation, usually 1-2 large P-bodies are observed, while hyperosmotic stress and defects in the secretory pathway induce numerous smaller P-bodies . The different morphologies also suggest that the protein composition of the Pbodies might be dependent on the stressor. Although most of the research on P-bodies has been performed in Saccharomyces cerevisiae, because of the evolutionary conservation of the of the P-body components (Fig. 1A) and its functions, the results obtained in yeast are highly relevant for all metazoans. P-bodies are membrane-less organelles. Some of the protein components contain unstructured regions or a RecA fold that are able, together with RNA to engage in liquid-liquid phase separation. Indeed, recently it was shown that Dhh1/DDX6 could drive phase separation in vitro and in vivo (Hondele et al., 2019). Other proteins, such Pat1, Edc3 and Scd6 have also been shown to contribute to P-body assembly under at least some stress conditions (Decker, Teixeira and Parker, 2007;Teixeira and Parker, 2007;Sachdev et al., 2019). It appears as if Dhh1 and Pat1 act together, while Scd6 and Edc3 seem to have partially redundant functions in P-body formation (Coller and Parker, 2005;Decourty et al., 2008;Nissan et al., 2010).
Thus, P-body assembly and function might be regulated through different pathways, consistent with the findings that a variety of kinases can regulate granule assembly (Yoon, Choi and Parker, 2010;Ramachandran, Shah and Herman, 2011). While under stress P-bodies are easily detected by light microscopy, they are essentially undetectable in unstressed cells. Yet, smaller P-body degradative units exist in unstressed cells as the major RNA degradation pathway in yeast is Xrn1-dependent, which acts in P-bodies (Parker, 2012). While the pathway by which Dhh1/DDX6 and Pat1 drive P-body formation is relatively well understood, information about the Scd6/Edc3-dependent pathway is still scarce.
Therefore, we decided to analyze the role of Scd6 and Edc3 in P-body assembly. We found that both proteins are required to retain the decapping complex in the cytoplasm. In an edc3∆ scd6∆ mutant, Dcp1 and Dcp2 accumulated in the nucleus through active import by Kap95. Nuclear localized Dcp1/Dcp2, however, did not drive nuclear mRNA decay. We propose that the nuclear decapping complex acts as a reservoir to regulate mRNA decay and decapping activity in the cytoplasm. We show furthermore that P-body assembly happens primarily on the endoplasmic reticulum (ER) and that in this process Edc3 and Scd6 link the decapping complex to Dhh1 (Fig.1B). Therefore, Dcp2 is also essential for P-body formation. We envisage that Dcp1/Dcp2 acts to increase the necessary critical concentration of Dhh1 at the ER to drive phase separation.

Concomitant loss of Scd6 and Edc3 blocks P-body assembly and drives nuclear accumulation of Dcp2
Previous studies have shown that the individual deletions of EDC3 and SCD6 only partially affect PB formation (Decker, Teixeira and Parker, 2007;Rajyaguru, She and Parker, 2012) and are dispensable for growth (Kshirsagar and Parker, 2004;Decourty et al., 2008). However, since they might have partially overlapping functions, we generated an edc3∆ scd6∆ double mutant and assessed its ability to form P-bodies under stress using Dcp2-GFP as a marker (Fig. 1C). As observed previously, deletion of either SCD6 or EDC3 affected P-body formation under hypoosmotic stress and to a somewhat lesser extent under starvation ( Fig. 1C-E) . This effect was strongly exacerbated in the double mutant, consistent with the notion of a redundant function in P-body granule formation. Surprisingly, we also observed an accumulation of Dcp2-GFP in the nucleus in the single mutants, which was again strongly increased in the double mutant under normal growth conditions ( Fig.1 C and F). In fact, in edc3∆ scd6∆ cells, the nuclear Dcp2 localization was maintained even under stress conditions, while this effect was less noticeable in edc3∆ or scd6∆ cells. These data suggest that there might be a correlation between Dcp2 localization and the ability of the cell to form P-body granules.
Next, we asked whether the nuclear accumulation was specific for Dcp2 or whether other P-body components would behave in a similar manner in the absence of Edc3 and Scd6. While Dcp1, which is part of the decapping complex, acted similarly to Dcp2, the exonuclease Xrn1 and the Lsm-associated protein Pat1 remained cytoplasmic ( Fig. 1G-I). Therefore, loss of Edc3 and Scd6 causes the selective nuclear accumulation of the decapping complex. Nevertheless, Pat1 had been shown previously to become nuclear localized in an lsm1∆ mutant ). In addition, in mammalian cells, Pat1's interaction with both the splicing machinery in the cell nucleus and cytoplasmic P-bodies has been demonstrated (Vindry et al., 2017). Yet, Dcp2 was not enriched in the nucleus under the same conditions (Fig. 1I). Our data indicate that at least two independent pathways exist to control the cytoplasmic/nuclear distribution of a subset of P-body components.

Dcp1/Dcp2 requires active import by Kap95 for nuclear localization
It is conceivable that Dcp2 depends on Dcp1 to be localized to the nucleus in edc3∆ scd6∆ cells. To test this possibility, we generated a triple deletion edc3∆ scd6∆ dcp1∆.
In this strain, Dcp2 still reached the nucleus ( Fig. 2A), indicating that nuclear import does not depend on the assembled decapping complex and that Dcp2 itself must contain a nuclear import signal. We analyzed the Dcp2 sequence with the NLStradamus model (Nguyen Ba et al., 2009) for NLS prediction and found three potential monopartite NLSs in the amino acid regions 458-467, 562-576 and 687-708.
The last region was also identified using cNLS Mapper, a web-based application for prediction of importin substrates (Kosugi et al., 2009), as a part of a moderately strong bipartite NLS spanning over amino acids 673-707. The presence of putative NLSs in the Dcp2 sequence suggests the import might be mediated by the karyopherin α/β (Kap60/Kap95) complex. Both KAP60 and KAP95 are essential for cell viability. Therefore, we tagged KAP95 C-terminally with an auxin-inducible degron (AID) in the edc3∆ scd6∆ background to acutely deplete Kap95 upon addition of auxin. We observed, however, that import of Dcp2-GFP into the nucleus was already impaired in the absence of auxin, indicating that the addition of the degron resulted in a hypomorphic kap95 allele ( Fig. 2B and C). Therefore, we conclude that the nuclear localization of Dcp2 is dependent on karyopherin ß.

The decapping complex is stored in the nucleus as a readily releasable pool
Next, we asked what could be the role of the decapping complex in the nucleus. 2D and E). While high levels of Dcp2 CD reduced cellular fitness over a range of temperatures, confining Dcp2 CD to the nucleus, rescued this phenotype. Likewise, a high nuclear concentration of Dcp2 was not toxic (Fig. 2F). Our data suggest that Dcp2 has essential decapping functions in the cytoplasm but not in the nucleus.
Moreover, high levels of nuclear Dcp2 are well tolerated, suggesting that Dcp2 may not be active in the nucleus.
Decapped RNA is unstable in the nucleus (Kufel et al., 2004). Our results above indicate that Dcp2 should not enhance RNA degradation in the nucleus. To this end, we carried out an mRNA degradation assay in a nup145∆C mutant, which is deficient for mRNA nuclear export at the restrictive temperature (Kufel et al., 2004).
Logarithmically growing cells were first deprived of methionine at the permissive temperature (23°C) to induce the expression of the MET3 gene and then shifted to the restrictive temperature (37°C) to block nuclear mRNA export. MET3 expression was then shut off by addition of excess methionine to the medium, and the progress of degradation of the MET3 mRNA trapped in the nucleus was assessed by Northern blot analysis. MET3 mRNA decay was not accelerated when we trapped Dcp2 in the nucleus by either using the edc3∆ scd6∆ mutant or by overexpressing NLS SV40 -Dcp2, or NLS SV40 -Dcp2 CD as a control (Fig. 2G-I). Therefore, we conclude that Dcp2 is not actively decapping nuclear mRNAs for decay under these conditions. Taken together, our data do not support a nuclear function for Dcp2 and are consistent with the notion that the nuclear Dcp2 pool may serve as a buffer to control cytoplasmic decapping activity.

Dcp1/Dcp2 performs essential functions on the cytoplasmic face of the ER
If the nuclear pool of Dcp2 operates as a reservoir, Dcp2's essential function should be in the cytoplasm, in particular under stress conditions, when P-bodies are formed.
Indeed, an edc3∆ scd6∆ mutant, in which Dcp2 becomes sequestered in the nucleus, showed impaired growth under stress (Fig. 3A). If this growth impairment was solely due to the Dcp2 localization, then Dcp2 overexpression ( Fig. 3B and C) should rescue the growth defect. Overexpression of Dcp2 alleviated the growth phenotype of the edc3∆ scd6∆ mutant strain (Fig. 3D). These data indicate that Edc3 and Scd6 collaborate to regulate cytoplasmic Dcp2 levels and that excess Dcp2 can be stored in the nucleus until needed.
The simple overexpression, however, has two drawbacks. First, it increases the entire cytoplasmic pool of Dcp2 without providing spatial information and second the overexpressed protein can still be imported and trapped in the cell nucleus. To circumvent these potential pitfalls, we locked Dcp2 in the cytosol. To this end, we anchored Dcp2-GFP on the cytosolic face of the endoplasmic reticulum (ER) and of mitochondria (mito) by linking Dcp2-GFP to Dpm1 (Dcp2 ER ) or Fis1 (Dcp2 mito ) ( Fig. 3E and F). This approach allowed Dcp2 expression from its endogenous promoter and to be anchored at specific locations in the cytoplasm. We chose the ER because we have previously shown that P-bodies associate with ER membranes ( Fis1 localized efficiently to mitochondria, and the construct carrying the Dpm1 colocalized with the ER. To ensure that the endogenous Dcp2 would not interfere with our assay, we acutely depleted endogenous Dcp2 using an auxin-inducible degron. This depletion worked efficiently as even under normal growth conditions Dcp2 depleted edc3∆ scd6∆ cells were unable to grow (Fig. 3G). Under these conditions, however, both mitochondrial and ER-localized Dcp2 rescued the growth phenotype.
Under stress conditions, the ER-sequestered Dcp2 allowed much better survival when compared to Dcp2 on mitochondria (Fig. 3G). Therefore, ER-localized Dcp2 is chiefly responsible to cope with stress.
Nevertheless, we observed also some rescue by the mitochondria-localized Dcp2. To understand these results better, we first tested, whether P-bodies could be formed on mitochondria under stress conditions. Indeed, Dcp2 mito formed foci, resembling P-bodies under glucose starvation (Fig. 3H). These foci were also positive for two other bona fide P-body components, Dhh1 and Pat1 (Suppl. Fig. 3B and C), indicating that P-bodies can also form on mitochondria. Moreover, our data suggest that Dcp2 alone is sufficient to determine the location of P-body formation.
The ER and mitochondria are connected via contact sites to allow the exchange of lipids and ions (Elbaz and Schuldiner, 2011;Prinz, 2014). In yeast, the tethering complex ERMES stabilizes these contacts (Kornmann et al., 2009). We wondered whether the P-bodies formed containing Dcp2 mito would be localized close to ERmitochondria contact sites. Indeed, P-bodies were detected next to or at the same site as the ERMES component Mdm34 ( Fig. 3I and J). This presence at ER-mitochondria contact sites was essential for the ability of Dcp2 mito to mount an appropriate stress response because destruction of the contacts by deleting the ERMES component Mmm1 abolished growth of Dcp2 mito expressing cells under stress (Fig. 3K). Taken together, our data are consistent with the notion that functional P-body formation under stress takes place at the ER. Moreover, our data provide evidence that Dcp2 and Pbodies can also act in trans to cope with stress at ER-mitochondrial contact sites, albeit somewhat less efficiently.

Scd6 and Edc3 bridge the interaction of Dcp2 with Dhh1 during P-body assembly
Our data above suggest that Dcp2 localization determines where P-bodies assemble.
Dcp2 can interact with both Scd6 and Edc3 (Fig. 4A), which in turn also interact with Dhh1, although their binding to Dhh1 was reported to be mutually exclusive (Fromm et al., 2012). Importantly, Dhh1 has been shown to drive phase separation, which is essential during P-body formation (Mugler et al., 2016;Hondele et al., 2019). First, we wanted to determine which parts of Scd6 and Edc3 are required to keep Dcp2 in the cytoplasm and to promote functional stress response. To this end, we carried out a domain analysis of Scd6 and Edc3 by overexpressing their individual domains alone or in combination in the edc3∆ scd6∆ background and determined the nuclearcytoplasmic distribution of Dcp2 and fitness at 37°C (Fig. 4B-E). As reported previously, overexpressing full-length of Scd6 impaired growth due to constitutive Pbody formation, even without stress (Nissan et al., 2010). Removal of the C-terminal region of Scd6 or Edc3 not only alleviated these growth defects, but restored growth to wild-type levels and reversed the P-body phenotype. Scd6 or Edc3 constructs lacking either the Dhh1 or the Dcp2 interaction site failed to restore growth and Dcp2 became enriched in the nucleus. Our data demonstrate that both Scd6 and Edc3 can bridge the interaction between Dcp2 and Dhh1, and that this connection is needed to keep Dcp2 from being transferred into the nucleus.

Linking Dcp2 and Dhh1 drives P-body formation and functional stress response
So far, our data suggest that Dcp2 and Dhh1 must come together to drive P-body formation under stress and that this process is mediated through interaction with either Scd6 or Edc3. To further corroborate our findings, we fused the N-terminal Dcp2 binding domain of Edc3 (Edc3(1-86), Edc3 N ) to Dhh1 and expressed the construct in the edc3∆ scd6∆ Dcp2-GFP strain (Fig. 5A). Consistent with the data described above, Dcp2 is mostly nuclear in edc3∆ scd6∆ cells (Fig. 5B). This localization did not change when either Dhh1 or the Edc3 N were expressed separately. However, the expression of the Edc3 N -Dhh1 fusion protein sequestered Dcp2 in the cytoplasm. More importantly, when we repeated the experiment under glucose starvation, expression of only Edc3 N -Dhh1 was sufficient to induce P-body formation ( Fig. 5C and D). These P-bodies appear to be functional as they completely rescued the growth phenotype of edc3∆ scd6∆ (Fig. 5E).

Dcp2 is essential for P-body assembly
Thus, our data suggest that Dcp2 is essential for P-body formation. However, it is assumed that P-body assembly is redundant and no single component is essential . Therefore, we decided to revisit the issue and determined P-body formation in our Dcp2-AID degron strain. Similarly, to what had been described previously, upon Dcp2 depletion, P-body formation was strongly impaired (Fig. 6A and   B). Instead of 1-3 bright foci/cell, either only diffuse signal or multiple weak foci were observed, which were most conspicuous in the case of Edc3. To test whether these smaller foci might present smaller functional P-body units, we performed colocalization analyses. While Edc3 colocalized with Xrn1 and Pat1 very well in the presence of Dcp2, this level of co-localization dropped drastically in the absence of Dcp2 (Fig. 6C-E). Therefore, we conclude, that even though smaller speckles can be formed in the absence of Dcp2, they do not represent functional P-bodies, indicating that Dcp2 is essential for P-body formation. Moreover, our data are in accordance with previous data (Weidner et al., 2014) that granule formation and phase separation do not necessarily correlate with P-body functionality.

Discussion
Even though P-bodies are an essential part of the cellular stress response, their assembly and cellular location are still debated and not fully understood. Recent studies highlighted the dynamics of individual P-body components during and the removal after stress (Xing et al., 2020) (Lee et al., 2020). The mechanism of their formation, however, is still not entirely clear. In another recent study, it was shown that the phase separation capability of the helicase Dhh1 contributed to P-body formation (Hondele et al., 2019). Yet, the dogma in the field is that no individual P-body component is essential for P-body formation. This reasoning is largely based on a study in which all major P-body components were individually deleted and the formation of foci with individual P-body members was analyzed ) with a recent follow up from the same group (Rao and Parker, 2017). The initial systematic study showed that none of the deletions suppressed completely foci formation. Here, we show that even though some P-body components form speckles in the absence of Dcp2, these appear not to be functional P-bodies as they lack other P-body components. Moreover, our data provide strong evidence that functional Pbodies are formed at the ER under stress. We envisage an assembly pathway in which Upon stress, Scd6 would inhibit translation either through its direct interaction with eIF4G, or with the help of Dhh1 which likewise has translational suppressor activity (Coller and Parker, 2005;Zeidan et al., 2018). However, in order to form a P-body, the local concentration of Scd6-Dhh1-mRNA complexes must be increased. This is achieved through binding to Dcp2 on the ER membrane (Fig. 7). We surmise that on the membrane, which reduces the diffusibility of the Dcp2-Scd6-Dhh1-mRNA complexes, it is much easier to gain a critical concentration to initiate Dhh1-driven phase separation. In a next step, Edc3, which like Scd6, can bind both Dcp2 and Dhh1 is recruited. Because Edc3 is a dimer, it can act as a scaffold to further enhance recruitment of Dhh1 and P-body formation. In our model, the local concentration of Dcp2 on ER polysomes would be key to drive P-body formation. In support of our model, we find that first Dcp2 ER can rescue a scd6∆ edc3∆ mutant, while Dcp2 mito cannot, unless concentrated at ER-mitochondrial contact sites. Second, directly linking Dcp2 to Dhh1 through the N-terminus of Edc3 (Edc3 N -Dhh1) is sufficient for cells cope with stress. Moreover, an mRNA coupled to Scd6 is on the one hand translationally repressed in a Dhh1-dependent manner and on the other hand destabilized by Dcp2 How does our model explain why deletions of for example PAT1 and LSM1 also affect P-body assembly? We propose that there is an initiation phase, and this is largely the process described above in which translational repression is intimately coupled to the initiation of P-body formation. In the next step, these molecular assemblies need to grow and to be stabilized, during which process those P-body components associated with 3' of the client mRNA are needed. In our view, in pat1∆ and lsm1∆ the initial assembly of the 5' P-body members with the RNA at the ER is not defective, but rather stabilization of the assembly and its growth is disrupted. In this phase of P-body stabilization, the numerous partially redundant interactions among different P-body members are important as previously described (Rao and  Supplementary Table 1. Standard genetic techniques were used throughout (Sherman, 1991). All modifications were carried out chromosomally, except where indicated. Chromosomal tagging and deletions were performed as described previously (Knop et al., 1999;Goldstein and McCusker, 1999;Gueldener et al., 2002;Janke et al., 2004). DCP2 and KAP95 were genomically tagged with an auxin-inducible degron AID*-9MYC and AID*-6HA respectively using pNat-AID*-9myc and pHyg-AID*-6HA plasmid templates for generation of the C-terminal tagging cassettes (Morawska and Ulrich, 2013). NUP145 was truncated by genomic Cterminal tagging with 3MYC to yield the mutant nup145∆C allele (nucleotides 1-1815 of the original ORF). TRP1 marker in the genomically integrated YIp204-ADH1p-AFB2 was disrupted using kanMX or hphMX4 cassettes.

Plasmid construction
Shuttle vectors for expression in yeast were prepared from pRS414-ADH and pRS415-GPD backbone plasmids (Mumberg, Müller and Funk, 1995) by Gibson assembly using NEBuilder HiFi DNA Assembly Cloning Kit according to the manifacturer's protocol (New England Biolabs). The fragments for the assembly were prepared by PCR using Q5 High Fidelity polymerase (New England Biolabs). The plasmids used along with details on their preparation are listed in Supplementary   Table 2.

Fluorescence Microscopy
Yeast cells were cultured at 30°C in YPD in case of genomically integrated modifications or in the respective HC-selection media when harboring plasmids.
Cultures were diluted in HC-complete medium, re-grown to log phase, and either

Nuclear mRNA degradation assay
The nuclear degradation of the MET3 mRNA was assayed in a nup145∆C mutant background. nup145∆C mutants are viable at 23°C, but show a strong mRNA nuclear export defect at 37°C (Kufel et al., 2004). Logarithmically growing mutant cells were first shifted to a medium lacking methionine to induce the expression of the methionine-related genes (4 h), then to 37°C for 30 min to inhibit the nuclear mRNA export and finally methionine-related genes expression was shut off by addition of excess of methionine to the medium. Aliquots were taken out, spun down and frozen in liquid nitrogen at specific times. For preparation of total RNA the cell pellets were P13, P100 and S100 were recovered, denatured with Laemmli buffer at 65°C for 5 min and analyzed by Western blotting.  Logarithmically growing cells expressing genomically tagged Dcp2-GFP were imaged without further treatment. Scale bar 5 µm. (B) Dcp2 nuclear import is dependent on Kap95. KAP95 in edc3∆ scd6∆ cells expressing genomically tagged Dcp2-GFP and the nuclear marker Nup84-mCherry was C-terminally tagged with an auxin-inducible degron. Logarithmically growing cells from the starting strain and the KAP95-tagged variant were imaged without further treatment. Scale bar 5 µm. (C) Quantification of the nuclear-cytoplasmic Dcp2-GFP distribution of the data in panel B. (D) and (E) Overexpression of NLS-appended and catalytically dead (CD) variants of Dcp2. Cells expressing Dcp2-GFP from a genomic locus were transformed with a low-copy plasmid expressing different versions of Dcp2-GFP from the strong GPD promoter. Cells from a logarithmically grown culture were imaged by fluorescence microscopy (D, scale bar 5 µm) or GFP expression was analyzed by Western blotting (F). High levels of Dcp2 as well as Dcp2 sequestered in the cell nucleus do not affect cell fitness. Serial dilutions of the strains in panels (D) and (E) were spotted on YPD-agar, and incubated for 2 days at the indicated temperatures. (G), (H) and (I) Northern blot analysis of MET3 mRNA nuclear degradation. nup145∆C strain background was either deleted for EDC3 and SCD6, or transformed with low-copy plasmids expressing the indicated Dcp2 constructs. Logarithmically growing cultures were starved for methionine to induce the expression of MET3 and then shifted to 37°C to inhibit mRNA nuclear export. MET3 transcription was then blocked by addition of methionine and aliquots for Northern blotting analysis were taken at the indicated time points. The scR1 mRNA served as a loading control. Dcp2-GFP was expressed on a GPD promoter from a low-copy plasmid on top of genomically tagged Dcp2-GFP in edc3∆ scd6∆. Logarithmically growing cells were either imaged directly (B, scale bar 5 µm) or GFP expression was analyzed by Western blotting (C). (D) Overexpression of Dcp2 partially rescues growth of edc3∆ scd6∆ under stress. Wild-type and edc3∆ scd6∆ strains expressing Dcp2-GFP on a GPD promoter from a low-copy plasmid were serially diluted and spotted on YPD-agar, and incubated at the indicated temperatures for 2 days. (E) Structure and subcellular localization of the Dcp2-GFP-MITO. edc3∆ scd6∆ with genomically tagged Om45-mCherry were transformed with a low-copy plasmid expressing Dcp2-GFP-Fis1 from a DCP2 promoter. Logarithmically growing cultures were imaged directly, scale bar 5 µm. (F) Structure and subcellular localization of the Dcp2-GFP-ER. edc3∆ scd6∆ cells with genomically tagged Pho88-mCherry were transformed with a low-copy plasmid expressing Dcp2-GFP-Dpm1 from a DCP2 promoter. Logarithmically growing cultures were imaged directly, scale bar 5 µm. (G) Dcp2 at the ER is required for growth under stress. edc3∆ scd6∆ with DCP2 tagged genomically with an auxin-inducible degron were transformed with low-copy plasmids expressing the indicated Dcp2-constructs from the endogenous DCP2 promoter. Serial dilutions of the respective logarithmically growing cultures were spotted on YP-agar with 2 or 0.2 % glucose, supplemented with 0.2 M auxin as indicated. Agar plates were incubated either for 2 days at 30°C or first for 2 days at 37°C, and then for 3 days at 30°C. (H) Dcp2-MITO forms foci upon glucose starvation. edc3∆ scd6∆ strain with the endogenous DCP2 tagged with an auxin-inducible degron was transformed with a low-copy plasmid expressing the Dcp2-MITO construct from a DCP2 promoter. Logarithmically growing cultures were treated with 2 mM auxin for 2 h, and either imaged directly or upon 30 min glucose deprivation. Scale bar 5 µm. (I) Dcp2-MITO foci localize in close vicinity of ERmitochondria contact sites. edc3∆ scd6∆ strain with genomically tagged Mdm34-mCherry and Dcp2-Aid was transformed with a low-copy Dcp2-GFP-MITO construct. Logarithmically growing cells were treated as for panel 3H. (J) Quantification of the number of GFP foci in panel 3-I colocalizing or juxtaposed to Mdm34 upon 30 min of glucose starvation. (K) ER-Mitochondria contact sites are essential for the functioning of Dcp2-MITO under stress. edc3∆ scd6∆ with the endogenous DCP2 tagged with an auxin-inducible degron was deleted for MMM1 and transformed with a low-copy plasmid expressing Dcp2-MITO from a DCP2 promoter. Logarithmically growing cultures of the indicated strains were serially diluted and spotted on YP-agar with 2 or 0.2 % glucose, supplemented with 0.2 M auxin, and incubated at 30°C for 3 days. (B) Bridging Dcp2 to Dhh1 is required to keep Dcp2 in the cytoplasm. Wild-type or edc3∆ scd6∆ with genomically tagged Dcp2-GFP were transformed with low-copy plasmids expressing the indicated SCD6 or EDC3 constructs from the strong GPD promoter. Logarithmically growing cells were imaged without further treatment. Scale bar 5 µm. (C) Quantification of the nuclear-cytoplasmic GFP distribution in the cells on panel 4-B. D and E. Scd6 and Edc3 bridge Dcp2 to Dhh1 to cope with stress. Logarithmically growing cultures from the strains in panels B and C were serially diluted and spotted on HC-agar lacking leucine, and incubated at 30 or 37°C for 2 days. The cartoon shows the Dcp2-GFP nuclear-cytoplasmic distribution.  . Dcp2 is essential for P-body assembly. A-B. Depletion of Dcp2 impairs PB formation. All strains were constructed from an isogenic background with DCP2 genomically tagged with an auxin-inducible degron. The respective genes were either genomically appended with GFP or mCherry at the C-terminus (EDC3, SCD6 and XRN1), or the respective mCherry-tagged protein was expressed from a low-copy plasmid on its endogenous promoter (Dhh1 and Pat1). Logarithmically growing cells were treated with 2 mM auxin for 2 h, then deprived from glucose for 30 min and imaged (A, scale bar 5 µm), and the number of mCherry foci was quantified (B). C-E. Dcp2 coordinates the recruitment of Xrn1 and Pat1 to Edc3 during PB formation. The strains expressing Edc3-GFP from panel 6A was genomically tagged with mCherry at the XRN1 locus (C) or transformed with a low-copy plasmid expressing Pat1-mCherry from its endogenous promoter (D). Logarithmically growing cells were treated as in panel 6A. Scale bar 5 µm. (E) Manders coefficients for the fraction Xrn1-mCherry or Pat1-mCherry colocalizing to Edc3-GFP upon 30 min -D.