# Formation of S. pombe Erh1 homodimer mediates gametogenic gene silencing and meiosis progression

Ditipriya Hazra, Vedrana Andrić, Benoit Palancade, Mathieu Rougemaille, Marc Graille

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Ditipriya Hazra, Vedrana Andrić, Benoit Palancade, Mathieu Rougemaille, Marc Graille. Formation of S. pombe Erh1 homodimer mediates gametogenic gene silencing and meiosis progression. Scientific Reports, 2020, 10 (1), pp.1034. 10.1038/s41598-020-57872-4. hal-02456264

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# Formation of *S. pombe* Erh1 homodimer mediates gametogenic gene silencing and meiosis progression

Ditipriya Hazra<fnref n="1" />,⁴, Vedrana Andrić²,⁴, Benoit Palancade³, Mathieu Rougemaille²* &
Marc Graille<fnref n="1" />*

Timely and accurate expression of the genetic information relies on the integration of environmental cues and the activation of regulatory networks involving transcriptional and post-transcriptional mechanisms. In fission yeast, meiosis-specific transcripts are selectively targeted for degradation during mitosis by the EMC complex, composed of Erh1, the ortholog of human ERH, and the YTH family RNA-binding protein Mmi1. Here, we present the crystal structure of Erh1 and show that it assembles as a homodimer. Mutations of amino acid residues to disrupt Erh1 homodimer formation result in loss-of-function phenotypes, similar to erh1Δ cells: expression of meiotic genes is derepressed in mitotic cells and meiosis progression is severely compromised. Interestingly, formation of Erh1 homodimer is dispensable for interaction with Mmi1, suggesting that only fully assembled EMC complexes consisting of two Mmi1 molecules bridged by an Erh1 dimer are functionally competent. We also show that Erh1 does not contribute to Mmi1-dependent down-regulation of the meiosis regulator Mei2, supporting the notion that Mmi1 performs additional functions beyond EMC. Overall, our results provide a structural basis for the assembly of the EMC complex and highlight its biological relevance in gametogenic gene silencing and meiosis progression.

Members of the ERH protein family are small proteins found in metazoan, invertebrates as well as plants. These proteins are strongly conserved (no amino acid changes between frog and human proteins and only one difference between human and zebrafish proteins), arguing for strict evolutionary constraints and for a highly important function. This gene was originally identified 25 years ago from a genetic screen as a mutant enhancing the truncated wing phenotypes of fruit flies lacking the rudimentary (r) gene<fnref n="1" />, which encodes for the enzyme catalyzing the first three steps of the pyrimidine biosynthesis pathway. However, the biochemical function of ERH remains unclear despite its strong abundance in tumors compared to human normal cells, making it a very interesting candidate for functional characterization².

Growing evidence points towards a role of ERH in mRNA synthesis, maturation and nuclear export but the precise mechanism(s) involving this protein is (are) still unknown. This nuclear protein has been shown to interact with: (1) FCP1, the specific phosphatase for RNA PolII C-terminal domain³; (2) the transcription factor SPT5Î; (3) PDIP46/SKAR (now named POLDIP3, for Polymerase delta-interacting protein 3), which localizes to nuclear speckles, regions enriched in pre-mRNA splicing factors⁴,⁵; (4) SNRPD3, a subunit of the Sm complex, which is involved in mRNA splicing. ERH also interacts with CIZ1, a zinc finger protein acting as a DNA replication factor and present in replication foci⁶,⁷. ERH is necessary for chromosome segregation during mitosis, probably through its role on CENP-E mRNA splicing⁸,⁹. Indeed, in the absence of ERH, the CENP-E mRNA, encoding a kinetochore protein, is incorrectly spliced and pre-mRNAs are rapidly eliminated by the nonsense-mediated mRNA decay pathway⁹. Such link between splicing defects, cell cycle arrest and mitotic defects has already been observed for the depletion of other splicing factors¹⁰. In *Xenopus*, ERH has been shown to act as a transcriptional

<footnote n="1">¹BIOC, CNRS, Ecole Polytechnique, IP Paris, F-91128, Palaiseau, France. ²Université Paris-Saclay, CEA, CNRS, Institute for Integrative Biology of the Cell (I2BC), 91198, Gif sur yvette, France. ³Institut Jacques Monod, CNRS, UMR7592, Univ Paris-Diderot, Sorbonne Paris Cité, 75013, Paris, France. ⁴These authors contributed equally: Ditipriya Hazra and Vedrana Andrić. *email: mathieu.rougemaille@i2bc.paris-saclay.fr; marc.graille@polytechnique.edu</footnote>
repressor and to interact with DCoH/PCD (dimerization cofactor of HNF1/pterin-4a-carbinolamine dehydratase), a positive cofactor of the HNF1 homeobox transcription factor, by yeast two-hybrids¹¹. ERH protein is absent in *S. cerevisiae* yeast but expression of human ERH in budding yeast stimulates filamentous growth in low nitrogen media¹². Interestingly, this phenotype is reminiscent of the phenotype observed upon expression of the RBP7 subunit of the human RNA polymerase II in yeast¹³, arguing again for a potential role of ERH proteins in the control of mRNA metabolism.

Closely related proteins, sharing around 30% sequence identity with human ERH, are also present in *Schizosaccharomyces* such as *S. pombe* and in few other fungi¹⁴. Recent studies performed in *S. pombe* have clarified the role of Erh1, the ortholog of human ERH. Initially, the *ERH1* gene was identified as a suppressor of the meiotic arrest phenotype in *sme2Δ* fission yeasts, whereby Mm1l is not properly inactivated during meiosis¹⁵. Mm1l is a YTH-family protein¹⁶, which selectively recognizes RNA hexanucleotide motifs (e.g. UNAAAC) present in meiotic transcripts and triggers their nuclear elimination by the nuclear exosome during mitosis¹⁷. Upon meiosis onset, Mm1l is sequestered within a nuclear dot by a ribonucleoparticle (RNP) composed of the *sme2* + -encoded long non-coding RNA meiRNA and the master regulator of meiosis Mei2¹⁸.

Recent works showed that Erh1 and Mm1l form a 2:2 stoichiometric complex dubbed EMC (for Erh1-Mm1 complex) whereby two Mm1l peptides encompassing residues 95 to 122 are physically bridged by an Erh1 homodimer¹⁷,¹⁹,²⁰. EMC localizes to scattered nuclear foci in vegetative cells and associates with two distinct complexes¹⁹,²¹. The first one known as MTREC (for Mtl1-Red1 core) is composed of the zinc-finger protein Red1, the Mtr4-like RNA helicase Mtl1 and Pir1/Iss10 among others subunits¹⁵,¹⁹. MTREC cooperates with Mm1l to mediate degradation of meiotic mRNAs by recruiting the Rrp6 subunit of the nuclear exosome²². The second complex known to interact with EMC is the CCR4-NOT complex but despite its known function as a mRNA deadenylase, it is not involved in Mm1l-dependent meiotic mRNA clearance¹⁹,²¹,²³⁻²⁵. Instead, it is required for the integrity of heterochromatin and regulates the abundance of Mei2 protein during mitosis through the action of its E3 ubiquitin ligase subunit Mot2/Not4²¹. Interestingly, both MTREC (PAXT in human cells) and CCR4-NOT complexes are conserved in human cells, suggesting that ERH may also interact with these complexes in human cells. Furthermore, human ERH can partially rescues the sensitivity to sorbitol, but not to SDS or hydroxyurea, of *S. pombe* erh1Δ cells¹⁴, suggesting that the function of Erh1 and human ERH proteins is partially conserved.

Here, we describe the crystal structure of *S. pombe* Erh1 protein and compare it to the structures of metazoan ERH proteins that have already been solved as well as to the structure of the *S. pombe* Erh1-Mm1l complex that has been solved while this work was in progress²⁰. We observe that Erh1 organizes as a homodimer in which the two monomers contact each other via hydrophobic interactions, consistent with recent work²⁰. Structure-guided mutational analysis shows that formation of Erh1 homodimer is critical for cell growth at low temperatures and for its functions in meiotic mRNA degradation and meiosis progression. Interestingly, an Erh1 mutant (Erh1<sub>III<sub>R</sub>LL<sub>3R</sub></sub>) defective for dimerization still associates with Mm1l *in vivo*, suggesting that Erh1 monomer is sufficient for interaction with Mm1l while formation of Erh1 dimer is essential for EMC function. We also show that Erh1 does not contribute to the Mm1l-dependent down-regulation of Mei2 in mitotic cells, indicating that Mm1l exerts functions beyond its partnership with Erh1. Overall, our results provide a structural basis for Erh1 dimerization and underscore the biological importance of Erh1 homodimer formation during both the mitotic and meiotic cell cycles.

## Results and Discussion

**S. pombe Erh1 crystal structure.** Initial polycrystals of Erh1 protein with a 6-branches star shape were obtained from an initial large screen of crystallization conditions in the following condition (0.8 M ammonium sulfate; 0.1 Na citrate pH 4). Thanks to the use of a micro-focus beamline, a complete dataset of moderate quality could be collected by shooting on a single branch of the star. Larger crystals could be obtained by increasing the drop volume, varying the ammonium sulfate concentration and the buffer. From one of these crystals, we could collect a dataset of better quality (see Table 1 for dataset statistics) from which we could determine Erh1 structure by molecular replacement using the structure of human ERH as initial model and refine it to 1.95 Å resolution.

Erh1 monomer folds as an α-β protein composed of a four stranded anti-parallel β-sheet and three α helices packed onto the same face of the β-sheet (Fig. 1A). This fold is similar to that of metazoan ERH (rmsd values of 1.1–1.9 Å over 80 Cα atoms¹²,²⁶⁻²⁸). The loop connecting helices α1 and α2 is not visible in our structure of Erh1. This most likely results from the intrinsic flexibility of this loop as shown for the corresponding region of metazoan ERH proteins by X-ray crystallography or NMR¹²,²⁶⁻²⁸. Three copies (protomers A, B and C) of Erh1 protein are present in the asymmetric unit. Protomers A and B are virtually identical (rmsd values of 0.27 Å over 84 Cα atoms) while protomer C slightly differs as illustrated by its higher rmsd value when compared to the two other chains (1.3–1.4 Å over 84 Cα atoms; Fig. S1A). The largest differences between protomer C and the two other Erh1 molecules present in the asymmetric unit occur at helix α2 (2.5 Å translation along the helix longitudinal axis), at the hinge between helix α2 and the N-terminal extremity of strand β3, and to a lesser extent on helix α3 (1.3 Å translation). It is noteworthy that human ERH adopts a conformation similar to that observed for Erh1 protomer C.

**An evolutionarily conserved homodimer.** Among the three copies present in the asymmetric unit, protomers B and C associate to form a tight homodimer with a butterfly-like shape (Fig. 1B) while the protomer A forms a similar homodimer with a symmetry-related molecule (rmsd of 1.1 Å over 160 Cα atoms). This dimeric state is consistent with the molecular weight determined in solution by size-exclusion chromatography coupled to multi-angle laser light scattering (SEC-MALLS; Fig. 2A). In the homodimer, the Erh1 β-sheet face from each monomer, which is not packed against α helices, interacts to form a β-barrel (Fig. 1B). Each monomer engages an area of 850 Å² mostly formed by hydrophobic residues, which are strongly or strictly conserved within Erh1/ERH
<table><caption>**Table 1.** Data collection and refinement statistics.</caption><thead><tr><th>Space group</th><th>C222<sub>1</sub></th></tr></thead><tbody><tr><td colspan="2"><strong>Data collection</strong></td></tr><tr><td>Unit cell parameters</td><td>71.9 Å; 123.9 Å; 68.2 Å;<br>90°; 90°; 90°</td></tr><tr><td>Wavelength (Å)</td><td>0.97857</td></tr><tr><td>Resolution (Å)</td><td>50–1.95 (2.07–1.95)</td></tr><tr><td>R<sub>merge</sub> (%)</td><td>7.7 (190.8)</td></tr><tr><td>I/σI</td><td>14.3 (1.0)</td></tr><tr><td>Completeness (%)</td><td>99.6 (98)</td></tr><tr><td>CC<sub>1/2</sub> (%)</td><td>99.9 (50.4)</td></tr><tr><td>Redundancy</td><td>7.9 (7.55)</td></tr><tr><td>Observed reflections</td><td>176672 (26429)</td></tr><tr><td>Unique reflections</td><td>22390 (3501)</td></tr><tr><td colspan="2"><strong>Refinement</strong></td></tr><tr><td>Resolution (Å)</td><td>50–1.95</td></tr><tr><td>R/R<sub>free</sub> (%)</td><td>20/23.2</td></tr><tr><td colspan="2"><strong>Number of atoms</strong></td></tr><tr><td>Protein</td><td>2308</td></tr><tr><td>SO<sub>4</sub><sup>2-</sup>/Ethylene glycol/Tris/Acetate</td><td>5/20/8/4</td></tr><tr><td>Water</td><td>73</td></tr><tr><td colspan="2"><strong>B-factors (Å<sup>2</sup>)</strong></td></tr><tr><td>Protein</td><td>61</td></tr><tr><td>SO<sub>4</sub><sup>2-</sup>/Ethylene glycol/PEG</td><td>90.2/70.1/90/97</td></tr><tr><td>Water</td><td>58.6</td></tr><tr><td colspan="2"><strong>R.m.s. deviations</strong></td></tr><tr><td>Bond lengths (Å)</td><td>0.010</td></tr><tr><td>Bond angles (°)</td><td>0.94</td></tr></tbody></table>

orthologues from fungi, plants, insects, worm and animals (Fig. 1C). This rationalizes that the Erh1 homodimer is reminiscent of those observed for metazoan ERH proteins (rmsd 1.4 Å over 160 Cα atoms<sup>12,26,28</sup>).

An evolutionarily highly conserved region present on the side of the homodimerization surface has been shown to participate in the interaction with Mm1l (a short region encompassing residues 95–122<sup>20</sup>) while this work was in progress (Fig. 1D). Comparison between the Erh1-Mm1l and our apo-Erh1 structures reveals that protomer C is strongly similar to the Erh1-bound structure and hence compatible with Mm1l binding. On the contrary, significant differences support that protomers A and B are incompatible with Mm1l binding (Fig. S1B,C). Most interestingly, the crystal structure of Erh1-Mm1l complex reveals a 2:2 stoichiometry<sup>20</sup>, confirming a previously proposed model of Erh1-mediated Mm1l self-interaction<sup>17</sup>.

As in this complex, each Mm1l-[95–122] peptide interacts with Erh1 on a region spread at the homodimer interface (Fig. S1B), we have decided to investigate the functional role of Erh1 homodimerization. We then simultaneously mutated two residues located at the homodimer interface (Ile11 and Leu13) into Arg to generate the Erh1<sub>I11R,L13R</sub> mutant with the aim of disrupting Erh1 homodimer. First, we have purified this mutant upon expression in *E. coli*. During purification, the Erh1<sub>I11R,L13R</sub> mutant proved much less soluble than the wild-type protein and a fraction of this mutant had a tendency to aggregate. Yet, SEC-MALLS analysis of purified proteins clearly indicates that the Erh1<sub>I11R,L13R</sub> mutant is monomeric in solution while the wild-type protein is homodimeric (Fig. 2A). Circular dichroism analyses further support that the mutant adopts an overall fold similar to that of the wild-type protein (Fig. S2). Together, these data indicate that recombinant Erh1<sub>I11R,L13R</sub> is monomeric and well-folded *in vitro*.

To obtain additional insights into EMC complex assembly, we sought to determine whether Erh1 dimerization contributes to its association with Mm1l *in vivo*. We first generated *erh1Δ* strains expressing GFP-tagged versions of the dimeric wild-type Erh1 or the monomeric Erh1<sub>I11R,L13R</sub>. Analysis of total protein levels under denaturing conditions revealed that plasmid-encoded GFP-Erh1 and GFP-Erh1<sub>I11R,L13R</sub> were similarly expressed, indicating that the mutation does not affect the stability of the protein *in vivo* (Fig. 2B, compare lanes 3 and 4). In addition, both proteins accumulate at much higher levels than endogenous wild type Erh1-GFP (Fig. 2B, compare lane 2 to lanes 3 and 4). When preparing extracts for co-immunoprecipitation assays, however, we observed a significant decrease in the total amount of full-length Erh1<sub>I11R,L13R</sub> when compared to plasmid-borne wild-type Erh1 (Fig. 2C, compare lanes 3 and 4). This was likely due to a further susceptibility of the mutant protein to proteolysis in these conditions, as shorter, fast-migrating proteolytic fragments concomitantly accumulate. These proteolytic fragments were also detected when analyzing the wild type protein, albeit to a lower extent, strongly suggesting that Erh1<sub>I11R,L13R</sub> is more prone to degradation by cellular proteases present in native extracts (Fig. 2C, compare lanes 3 and 4). Importantly, in denaturing conditions, whereby endogenous proteases are also inactivated, these
<figure><img src="image_2.png" /><figcaption>**Figure 1.** *S. pombe* Erh1 structure. (A) Cartoon representation of Erh1 monomer. The protein is colored from blue (N-terminus) to red (C-terminus). The loop encompassing residues 47 to 54, which is not defined in electron density maps, probably due to high flexibility, is depicted as a dashed line. Panels A, B and D have been generated using the Pymol software version 1.7.2.2 Schrödinger, LLC (http://www.pymol.org/). (B) Homodimer representation of Erh1. The Ile11 and Leu13 residues mutated in this study are shown as sticks. (C) Sequence alignment of Erh1/ERH proteins from *Schizosaccharomyces* fungi (*S. pombe*, *S. cryophylus*, *S. octosporus* and *S. japonicus*), insects (*Apis mellifera* and *D. melanogaster*), plants (*Zea mays* and *Glycine max*), *C. elegans* worm and animals (*Danio rerio*, *Salmo salar* and *Homo sapiens*). Strictly conserved residues are in white on a black background. Partially conserved residues are boxed and in bold. This panel was generated using the ESPript server³⁷. Secondary structure elements detected from Erh1 apo structure are shown above the alignment. Residues involved in homodimer formation are indicated by black stars below the alignment. (D) Sequence conservation score mapped at the surface of Erh1 homodimer. The conservation score has been calculated using the Consurf server³⁸ from the alignment shown in Fig. 1D. Coloring is from white (low conservation) to red (strictly conserved). The Mmi1 region interacting with Erh1 in the Erh1-Mmi1 crystal structure²⁰, is shown as a cyan cartoon. The side chain for the Mmi1 tryptophan residue (W112) present at the heart of the interface is shown as sticks.</figcaption></figure>

degradation products were barely detectable and both forms of Erh1 were indeed expressed at similar levels (Fig. 2B).

With this caveat in mind, we next sought to determine whether Erh1<sub>I11R,L13R</sub> can self-associate *in vivo*. We co-transformed erh1Δ cells with two plasmids expressing TAP- and FLAG-tagged versions of wild type or mutant Erh1 and performed TAP pull-down prior to Western blot analyses. Consistent with the above results, total and immunoprecipitated TAP-Erh1<sub>I11R,L13R</sub> levels were significantly reduced compared to wild-type TAP-Erh1 (Fig. 2D, compare lanes 2 and 5). To ease comparison, we blotted diluted samples of the wild-type total extract and immunoprecipitate. While FLAG-Erh1 was efficiently immunoprecipitated with TAP-Erh1, FLAG-Erh1<sub>I11R,L13R</sub> was not detectable following TAP-Erh1<sub>I11R,L13R</sub> pull-down (Fig. 2D, compare lanes 2 and 3 to lane 5). These data indicate that, unlike the wild type protein, the Erh1<sub>I11R,L13R</sub> mutant is impaired for self-association *in vivo*. Yet, surprisingly, GFP-Erh1<sub>I11R,L13R</sub> still interacted with Mmi1-TAP (Fig. 2E, IP TAP panel), suggesting that Erh1 dimerization is not a prerequisite for its association with Mmi1, contrary to what could be assumed from the crystal structure of Erh1-Mmi1-[95-122] complex²⁰. However, we cannot formally exclude that residual dimer formation of Erh1<sub>I11R,L13R</sub> may contribute to its interaction with Mmi1 (Fig. 2E).
<figure><img src="image_3.png" /><figcaption>B.</figcaption></figure>

<figure><img src="image_4.png" /><figcaption>C. Immunoprecipitation conditions</figcaption></figure>

<figure><img src="image_5.png" /><figcaption>D.</figcaption></figure>

<figure><img src="image_6.png" /><figcaption>E.</figcaption></figure>

<figure><img src="image_7.png" /></figure>

**Figure 2.** Erh1<sub>I11R,L13R</sub> does not self-associate *in vivo* and *in vitro*. (A) Chromatograms resulting from SEC-MALLS analysis of wild-type (experimentally determined molecular weight of 24.38 kDa) and I11R/L13R double mutant (experimentally determined molecular weight of 12.57 kDa) Erh1 proteins. (B) Western blot showing expression levels of endogenous Erh1 (C-terminally fused to GFP at the gene locus) and plasmid-encoded GFP-Erh1 and GFP-Erh1<sub>I11R,L13R</sub> expressed from the P<sub>nmt41</sub> promoter (pREP41 vector) in *erh1Δ* cells, under denaturing conditions. An anti-CDC2 antibody was used for loading control. The asterisks denote proteolytic fragments. Note that for panels B and C, the position of the GFP tag and the length of the linker between the ORF and the tag are the likely reasons why C-terminally and N-terminally tagged versions of Erh1 have slightly different migration patterns. (C) Western blot showing expression levels of endogenous Erh1 (C-terminally fused to GFP at the gene locus) and plasmid-encoded GFP-Erh1 and GFP-Erh1<sub>I11R,L13R</sub> expressed from the P<sub>nmt41</sub> promoter (pREP41 vector) in *erh1Δ* cells, under immunoprecipitation conditions. An anti-CDC2 antibody was used for loading control. The asterisks denote proteolytic fragments. (D) The Erh1<sub>I11R,L13R</sub> mutant does not self-associate *in vivo*. Western blot showing that wild-type FLAG-Erh1 co-immunoprecipitates with wild-type TAP-Erh1, while FLAG-Erh1<sub>I11R,L13R</sub> is not pulled-down by TAP-Erh1<sub>I11R,L13R</sub>. (WCE) Whole Cell Extract; (IP) Immunoprecipitation; (PAP) Peroxidase-conjugated Anti-Peroxidase complex antibody (binds to the proteinA moiety of the TAP tag). (E) Monomeric Erh1<sub>I11R,L13R</sub> protein associates with Mmi1. Western blot showing that both GFP-Erh1 and GFP-Erh1<sub>I11R,L13R</sub> co-immunoprecipitate with Mmi1-TAP.

**Formation of Erh1 homodimer is required for gametogenic gene silencing.** To investigate the role of Erh1 dimerization *in vivo*, we first analyzed its impact on *S. pombe* cell growth. Similar to *erh1Δ* cells, the Erh1<sub>I11R,L13R</sub> mutant displayed growth defects at all tested temperatures, especially 23°C, indicating that Erh1 dimerization is essential for its function (Fig. 3A).

In vegetative cells, Erh1 and Mmi1 assemble in the EMC complex that localizes to nuclear foci<sup>17,19,20</sup>. Live cell microscopy experiments revealed lower and diffuse GFP-Erh1<sub>I11R,L13R</sub> signals when compared to GFP-Erh1 (Fig. 3B). Strikingly, nuclear dots were lost in the mutant, including in cells in which the GFP signal was similar to that of the wild type (Fig. 3B, red arrows). This supports the notion that Erh1 homodimer formation is a prerequisite for its confinement into nuclear bodies.

Erh1 cooperates with Mmi1 to target meiosis-specific transcripts for degradation by the nuclear exosome<sup>17,19,20</sup>. To evaluate the role of Erh1 homodimer in this pathway, we measured the levels of Mmi1 RNA targets by RT-qPCR, including the *mei4*+ and *ssm4*+ meiotic mRNAs as well as the lncRNA *meiRNA*. Cells lacking Erh1 or expressing Erh1<sub>I11R,L13R</sub> showed a strong accumulation of *mei4*+ and *ssm4*+ transcripts, while *meiRNA* levels
<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /><figcaption>**Figure 3.** Erh1 dimerization is required for meiotic mRNAs recognition and degradation by Mmi1. (A) The Erh1<sub>I11R,L13R</sub> mutant phenocopies the deletion of *erh1* + gene. Spotting assays at 23°C, 30 °C and 37 °C. Cells of the indicated genotypes were grown until mid-log phase and plated on EMM-LEU medium at an initial OD = 0.25 followed by 5-fold serial dilutions. (B) Live cell microscopy of GFP-tagged Erh1 and Erh1<sub>I11R,L13R</sub> in strains of the indicated genotypes. Cells were imaged by differential interference contrast (DIC) and with a GFP filter. Red arrows indicate Erh1<sub>I11R,L13R</sub> cells for which GFP signals are similar to that of wild type Erh1. Squares with white dashed lines lying on the bottom right of GFP panels show enlarged images of the small squares. (C) RT-qPCR analysis of the *mei4+*, *ssm4+* and *meiRNA* transcripts in strains of the indicated genotypes. Signals were normalized to *act1* + mRNA levels and expressed relative to wild-type cells. Error bars represent the standard deviation from four independent experiments. (D) RNA-immunoprecipitation experiments in *erh1Δ* cells expressing GFP-Erh1 or GFP-Erh1<sub>I11R,L13R</sub>. Shown are the enrichments (% input) of *act1+*, *mei4+*, *ssm4+* and *meiRNA* transcripts upon pull-down of TAP-tagged Mmi1. Error bars represent the standard deviation of four independent immunoprecipitations from biological duplicates.</figcaption></figure>

were only partially increased when compared to the *mmi1Δ* mutant (Fig. 3C). These results indicate that Erh1 dimerization is required for efficient Mmi1-dependent meiotic RNA degradation. To determine whether this was due to defective recruitment of Mmi1 to its targets, we analyzed the levels of meiotic RNAs co-precipitated with Mmi1. In cells expressing wild-type Erh1, Mmi1 efficiently bound to the *mei4+*, *ssm4+* and *meiRNA* transcripts (Fig. 3D). Instead, in the Erh1<sub>I11R,L13R</sub> mutant, the association of Mmi1 to meiotic mRNAs was abolished, while *meiRNA* still co-precipitated, albeit to a lower extent (Fig. 3D). Altogether, these data strongly suggest that
<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>

**Figure 4.** Erh1 dimerization is required for efficient meiosis progression and meiRNA dot formation. (A) Homothallic strains of the indicated genotypes were spotted on EMM-LEU plates and incubated for 5 days at 30°C. The presence or absence of asci was determined by iodine staining and live cell imaging (bright field). The mating/sporulation efficiency is indicated for each strain and represents the percentage of asci among 500 cells. (B) Representative images of meiRNA (red) detected by Single molecule RNA Fluorescence *In-Situ* Hybridization (SmFISH) in meiotic cells. DNA was stained with DAPI (blue). GFP-tagged wild-type Erh1 or Erh1<sub>I11R,L13R</sub> were visualized in parallel. Images are shown as the maximum-intensity projections of Z-stacks.

formation of Erh1 homodimer and hence proper EMC assembly is crucial for meiotic mRNAs recognition and degradation by Mmi1 but dispensable in the case of meiRNA. This different requirement for Erh1 dimer in the binding of Mmi1 to its RNA substrates might relate to the number of Mmi1 binding motifs (i.e. UNAAAC) within transcripts. Given *mei4*+ and *ssm4*+ mRNAs contain 8 and 7 binding sites, respectively, while meiRNA has up to 25<sup>29</sup>, it is possible that a pool of Mmi1 associates with the latter even in the absence of properly assembled EMC. Alternatively, low levels of Erh1<sub>I11R,L13R</sub> dimers may account for the residual binding of Mmi1 to meiRNA.

We previously showed that Mmi1 recruits the Ccr4-Not complex to promote ubiquitinylation and down-regulation of its own inhibitor Mei2, a master regulator of meiosis<sup>21</sup>. To determine whether Erh1 also contributes to this regulatory circuit, we analyzed Mei2 levels in cells lacking Erh1 or expressing Erh1<sub>I11R,L13R</sub>. We found that, contrary to *mmi1*Δ cells, Mei2 levels were not increased in mutants (Fig. S3), indicating that Erh1 does not contribute to Mei2 down-regulation in mitotic cells. This is also consistent with the notion that Mmi1 can exert functions independently of Erh1, as in case of transcription termination<sup>20,30</sup>.

**Formation of Erh1 homodimer contributes to meiosis progression.** Erh1 not only suppresses the meiotic program in vegetative cells but also stimulates meiosis progression<sup>17,19,20</sup>. To examine the requirement for Erh1 dimerization in meiosis, homothallic *erh1*Δ cells expressing wild-type Erh1 or Erh1<sub>I11R,L13R</sub> were exposed to iodine vapor, which stains the spore walls with dark color. Cells expressing wild-type Erh1 displayed strong staining intensity and high sporulation frequency, as indicated by the prevalence of asci (Fig. 4A). On the contrary, cells lacking Erh1 or expressing the Erh1<sub>I11R,L13R</sub> mutant showed reduced intensity in staining, consistent with lower sporulation efficiency and asci formation (Fig. 4A). However, meiosis was not completely abolished as in *mei4*Δ *mmi1*Δ cells, suggesting that the absence of Erh1 might be bypassed at least at low frequency.

Upon entry into meiosis, Mmi1 and Erh1 foci converge to a single nuclear dot associated with the meiosis regulator Mei2 and the lncRNA *meiRNA*<sup>18,19,31</sup>. To determine whether Erh1 homodimer is necessary for dot formation, we probed meiRNA in meiotic cells by fluorescence *in situ* hybridization. Consistent with previous studies<sup>18,19,31</sup>, meiRNA colocalized with wild-type GFP-tagged Erh1 in a unique nuclear dot (Fig. 4B). On the contrary,
cells expressing GFP-Erh1<sub>I11R,L13R</sub> failed to form both meiRNA and Erh1 dots (Fig. 4B). From these experiments, we propose that defects in Erh1 homodimer formation prevent the assembly of the EMC-meiRNA-Mei2 nuclear dot required for Mmi1 sequestration/inactivation during meiosis, thereby rationalizing impaired meiosis progression.

Conclusion

In this study, we report the crystal structure of Erh1 and show that it assembles as a homodimer through hydro-
phobic interactions between N-terminal residues. Consistent with recent work²⁰, the structure also reveals a
highly conserved region lying on the side of the dimerization domain and to which Mmi1 associates to form
the heterotetrameric EMC complex. The strong similarity between Erh1 and human ERH homodimers (this
study²⁰,²⁶,²⁸) raises the possibility that the assembly of ERH-based multimeric complexes has been maintained
throughout evolution for regulatory purposes, including modulation of gene expression. Whether Mmi1-related,
YTH family RNA-binding proteins or other factors bind to ERH homodimers in metazoans to regulate cellular
processes remains to be investigated.

Our functional analyses also highlight the biological relevance of Erh1 homodimer formation in gametogenic gene silencing. Importantly, Erh1 dimerization seems to be dispensable for interaction with Mmi1 but essential for Mmi1 binding to meiotic transcripts, implying that RNA recognition by the YTH domain is not sufficient *per se* and that only fully assembled EMC complexes are functionally competent. This illustrates the cooperation between the C-terminal YTH domain and the N-terminal disordered region of Mmi1, to which Erh1 associates, for optimal binding to RNA substrates. The mechanistic basis for this is presently unclear but it is possible that the assembly of large macromolecular machines (e.g. EMC, MTREC, Ccr4-Not) facilitates protein-RNA interactions. In line with this, it is tempting to speculate that EMC and meiotic mRNA nuclear foci observed in vegetative cells¹⁷,¹⁹ may reflect multimerization driven by Erh1 dimers of such RNP complexes, gathering multiple transcripts in close proximity for efficient degradation.

Another important finding from our work is the requirement for Erh1 dimerization in Mmi1 sequestration/
inactivation by the meiRNA-Mei2 dot during meiosis. The residual binding of Mmi1 to meiRNA observed in
Erh1<sub>I11R,L13R</sub> mitotic cells may not be sufficient for proper dot formation in meiosis. Therefore, Erh1 homodi-
mer formation not only promotes Mmi1 function during vegetative growth but also contributes to its inhibition
during meiosis. This cell-cycle dependent duality in functional outcomes may allow rapid changes in the activ-
ity of the complex without altering its expression levels or assembly. Whether this relates to the nature of the
RNA substrate (meiotic mRNAs versus meiRNA) and/or additional factors (e.g. Mei2) remains to be determined.
Regardless the precise mechanism, our study opens new perspectives to study the formation and activity of ERH
homodimer-containing complexes in metazoan and to understand their relevance to human diseases¹⁰.

Methods

**Cloning and protein expression.** The gene encoding for Erh1 was amplified using a *S. pombe* cDNA library by PCR with oligonucleotides oMG511 and oMG512 (Table S1) using the Phusion High-Fidelity DNA Polymerase (Thermo) according to manufacturer's instructions. The PCR products were further cloned into pGEX-6P1 vector using Fast Digest BamHI and XhoI restriction enzymes (Thermo Fisher Scientific) to generate the plasmid pMG921 encoding for a GST-tag fused to the N-terminal extremity of the full-length Erh1 by a 3C protease cleavage site (Table S1). The plasmid encoding for the Erh1<sub>I11R,L13R</sub> double mutant was obtained by one-step site-directed mutagenesis of pMG921 using oligonucleotides oMG629/oMG630 (Table S1) to yield plasmid pMG945.

The Erh1 protein was expressed in *E. coli* BL21 (DE3) Codon+ cells upon transformation with pMG921 plasmid. Cultures were performed in 1 L of auto-inducible terrific broth media (ForMedium AIMTB0260) containing ampicillin (100 µg/mL) and chloramphenicol (25 µg/mL) first for 3 hours at 37°C and then overnight at 18°C. The cells were harvested by centrifugation at 4100 rcf for 45 minutes. The pellet was resuspended in 30 mL lysis buffer (20 mM Tris-HCl pH 7.5, 200 mM NaCl, 5 mM β-mercaptoethanol) in the presence of 100 µM PMSF.

Cell lysis was performed by sonication on ice, followed by lysate clearance by centrifugation at 20000 rcf for 45 minutes. The supernatants were applied on GSH-sepharose resin pre-equilibrated with lysis buffer. After extensive washing steps with lysis buffer and with a high salinity buffer (20 mM Tris-HCl pH 7.5, 2 M NaCl, 5 mM β-mercaptoethanol), the protein was eluted with elution buffer (20 mM Tris-HCl pH 7.5, 200 mM NaCl, 20 mM GSH, 5 mM β-mercaptoethanol). The eluted protein was next incubated overnight at 4 °C with GST-3C protease under dialysis conditions in lysis buffer and then passed through GSH column to remove the GST-tag as well as the GST tagged 3 C protease. The unbound proteins were subjected to size exclusion chromatography using HiLoad 16/60 Superdex 75 column (GE Healthcare Biosciences) pre-equilibrated with lysis buffer on an ÄKTA Purifier system (GE Healthcare Biosciences). The Erh1<sub>I11R,L13R</sub> double mutant was purified using the same protocol.

**Crystallization, data collection and structure determination.** Crystallization conditions were screened by the sitting-drop vapor diffusion method using JCSG⁺ screen (Molecular Dimensions) at 4°C by mixing 150 nL of concentrated protein (7.5 mg/ml) solution with an equal volume of reservoir solution in a 96-wells TTP plates (TTPlabtech). Initial hits corresponding to star shaped crystals were obtained in 0.8 M ammonium sulfate; 0.1 Na citrate pH 4. For crystal optimization, hanging-drop method was used at 4 °C, by mixing 1 µL of concentrated protein with 1 µL of reservoir solution. The best dataset was collected from crystals obtained in 0.3 M ammonium sulfate; 0.1 M Na acetate pH 3.8.

Prior to data collection, the crystals were quick-soaked in cryo-protectant solutions containing 15% (v/v) and then 30% ethylene glycol in corresponding well solutions and flash-frozen in liquid nitrogen. X-ray diffraction datasets were collected on both Proxima-1 and Proxima-2a beamlines at synchrotron SOLEIL (Saint-Aubin,
France) and were processed with the XDS package³². The dataset collected on Proxima-1 showed a higher resolution limit and was then used to determine and refine the structure of Erh1 protein (Table 1). The structure was solved by molecular replacement searching for 3 molecules in the asymmetric unit with the program PHASER³³. The initial model for molecular replacement was generated by the PHYRE2 server³⁴ using the crystal structure of human ERH (30% sequence identify) as template²⁸. In this model, the loop corresponding to residues 44 to 55 was removed as it is known to be highly flexible from the comparison of human and fruit fly ERH structures¹²,²⁶,²⁸.

The final model was obtained by iterative cycles of building and refinement using COOT³⁵ and BUSTER³⁶, respectively (for final statistics, see Table 1). This model encompasses residues 6-46 and 55-100 for protomer A, 1-46 and 55-98 for protomer B (as well as 4 residues from the N-terminal tag) and 1-2, 7-47 and 54-98 for protomer C as well as 5 ethylene glycol molecules, 1 sulphate ion, 1 acetate molecule and 73 water molecules.

The atomic coordinates and structure factors have been deposited into the Brookhaven Protein Data Bank under the accession numbers 6S2W.

**S. pombe strains and growth media.** The *S. pombe* strains used in this study are listed in Table S2. Strains were generated by transformation with a lithium acetate-based method. The *mmi1Δ* cells were generated from a parental strain possessing a deletion of *mei4+*, since the absence of Mmi1 leads to severe growth and viability defects due to the deleterious expression of Mei4. All experiments were performed using minimal medium (EMM Broth, Formedium, #PMD0210) supplemented with 150 mg/L of each adenine (Sigma, #A2786), L-histidine (Sigma, #H8000), uracil (Sigma, #U750) and L-lysine (Sigma, #L5501) but lacking L-leucine (EMM-LEU). To assess mating/sporulation efficiency, cells plated on EMM-LEU medium for 5 days at 30°C were exposed to iodine crystals (Sigma, #326143) for 5 min at room temperature.

**Co-immunoprecipitation and Total protein analyses.** Experiments were performed as described in Simonetti *et al.*²¹ except that detection was done with a Vilber Lourmat Fusion Fx7 imager.

**RNA extraction and RT-qPCR.** Experiments were performed as described in Simonetti *et al.*²¹ except that 100 units Maxima Reverse Transcriptase (Thermo Scientific, #EP0743) were used for reverse transcription reactions.

Oligonucleotides used in qPCR reactions are listed in Table S3.

**RNA-immunoprecipitation.** Experiments were performed as described in Simonetti *et al.*²¹ with the following modifications: 40 ODs of cells were grown to mid-log phase at 30°C in EMM-LEU and cross-linked with 0.2% formaldehyde for 20 min. Following quenching with 250 mM glycine for 5 min, cells were harvested by centrifugation. Cell pellets were resuspended in 2 ml RIPA buffer (50 mM Tris-HCl pH 8, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 2 mM EDTA, 2 mM benzamidine, 1X Roche complete EDTA-free protease inhibitor cocktail and 80 U RNaseOUT Ribonuclease inhibitor (Invitrogen, #10777-019)) to make "pop-corn". Lysis was performed using a Ball Mill (Retsch, MM400) for 15 min at 15 Hz frequency. Extracts were cleared by centrifugation before immunoprecipitation with 1 mg of pre-washed rabbit IgG-conjugated M-270 Epoxy Dynabeads (Invitrogen, #14311D) for 1 hour at 4°C. Beads were then washed once with low salt buffer (10 mM Tris-HCl pH7.5, 150 mM NaCl, 0.5% Triton X-100), twice with high salt buffer (10 mM Tris-HCl pH7.5, 1 M NaCl, 0.5% Triton X-100) and once again with low salt buffer for 10 min at room temperature. Total and immunoprecipitated RNAs were de-cross-linked at 70°C for 45 min in the presence of reverse buffer (10 mM Tris-HCl pH 6.8, 5 mM EDTA, 10 mM DTT, 1% SDS) and treated with proteinase K for 30 min at 37°C. RNAs samples were next extracted with phenol:chloroform 5:1 pH4.7 (Sigma, #P1944), precipitated with ethanol and treated with DNase (Ambion, #AM1906) prior to RT-qPCR analyses.

**SmFISH.** Quasar 670-labeled meiRNA probes were designed using Stellaris Probe Designer tool (Table S4) and synthesized by Biosearch Technologies. Single molecule RNA Fluorescence *In-Situ* Hybridization (smFISH) was performed according to the manufacturer's protocol (Biosearch Technologies) with minor modifications.

Vegetative cells were plated on EMM-LEU and grown for 3 days at 30°C. Cells were then resuspended in 1X PBS containing 3.7% formaldehyde to an OD₆₀₀nm of 0.3, treated with Zymolyase 100 T for cell wall digestion and permeabilized in 70% ethanol prior to over-night incubation with meiRNA probes. Stellaris RNA FISH hybridization and wash buffers were obtained from Biosearch Technologies. DAPI stained cells were resuspended in Vectashield antifade mounting medium (Vector laboratories) and imaged using DM6000B Leica microscope with a 100X, numerical aperture 1.4 (HCX Plan-Apo) oil immersion objective and a charge-coupled device (CCD) camera (CoolSNAP HQ; Photometrics). Optical Z sections (0.2 µm step size, 25 sections) were acquired using a piezo-electric motor (LVDT; Physik Instrument) and the MetaMorph 6.1 software prior to maximum-intensity projection into a single plane. Images were processed in ImageJ (NIH).

Received: 19 July 2019; Accepted: 23 December 2019;

Published online: 23 January 2020

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## Acknowledgements

We acknowledge SOLEIL for provision of synchrotron radiation facilities and we would like to thank the beamline scientists for their assistance in using beamlines Proxima-1 and Proxima-2a. We are indebted to Dr Y. Mechulam for his help with data collection and to Pr. C. Gaillardin (INA-PG, Thiverval-Grignon, France) for the gift of the *S. pombe* cDNA library. This work was supported by Ecole Polytechnique, the Institut de Biologie Intégrative de la Cellule, the Centre National pour la Recherche Scientifique and the Agence Nationale pour la Recherche [grants ANR-16-CE11-0003 to M.G and ANR-16-CE12-0031-01 to M.R.]. D.H. was supported by a PhD fellowship from the French Ministère de l'Enseignement Supérieur et de la Recherche (MESR) and a mini post-doctoral fellowship from Ecole Polytechnique.

## Author contributions

D.H. and V.A. performed experiments. D.H., V.A., B.P., M.R. and M.G. designed the study, analyzed the data, drafted the article and approved the version to be published.

## Competing interests

The authors declare no competing interests.
**Additional information**

**Supplementary information** is available for this paper at https://doi.org/10.1038/s41598-020-57872-4.

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