{"paper_id":"2c8af8ae-622f-48ac-a249-39e0e99fd7fd","body_text":"1\n1\n2\n3 Rgf1 GEF activity toward Rho1 defines a new actin-dependent signal to determine growth sites \n4 independently of microtubules and Tea1\n5\n6\n7\n8 Patricia Garcia*, Ruben Celador, Tomas Edreira and Yolanda Sanchez*\n9\n10 Instituto de Biología Funcional y Genómica (IBFG), CSIC/Universidad de Salamanca and \n11 Departamento de Microbiología y Genética, Universidad de Salamanca. C/ Zacarías González, \n12 s/n. 37007 Salamanca, Spain.\n13\n14 Short title: Rgf1 and actin define growth sites \n15 * Correspondence should be addressed to: pgr@usal.es ; ysm@usal.es\n16 Key words: polarity, actin, Rho-GTPases, yeast\n17\n18\n19\n20\n21\n22\n23\n24\n25\n26\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n2\n27 ABSTRACT\n28 Cellular asymmetry begins with the selection of a discrete point on the cell surface that triggers \n29 Rho-GTPases activation and localized assembly of the cytoskeleton to establish new growth \n30 zones. The cylindrical shape of fission yeast is organized by microtubules that deliver the \n31 landmark Tea1–Tea4 complex at the cell tips to define the growth poles. However, only a few \n32 tea1Δ cells mistaken the direction of growth, indicating that they manage to detect their growth \n33 sites. Here we show that Rgf1 (Rho1-GEF) and Tea4 are components of the same complex and \n34 that Rgf1 activity toward Rho1 is required for strengthen Tea4 at the cell tips. Moreover, in cells \n35 lacking Tea1, selection of the correct growth site depends on Rgf1 and on a correctly polarized \n36 actin cytoskeleton, both necessary for Rho1 activation at the pole. We propose an actin-\n37 dependent mechanism driven by Rgf1–Rho1 that marks the poles independently of \n38 microtubules and the Tea1–Tea4 complex.\n39\n40\n41\n42\n43\n44\n45\n46\n47\n48\n49\n50\n51\n52\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n3\n53 INTRODUCTION\n54 Cell polarity is the primary mechanism for generating cellular asymmetry, which is critical for \n55 most cell and tissue functions such as development, cell migration and differentiation in a wide \n56 variety of organisms including humans. It typically begins with a signal on the cell surface that \n57 triggers a cascade of molecular events that induce the localized assembly of cytoskeletal and \n58 signaling networks, which subsequently direct the formation of a new growth area (1,2). Fission \n59 yeast has been used over the last decades as a simpler and more accessible model for studying \n60 this complex process (3). Its cells have a cylindrical shape that is maintained throughout the \n61 entire cell cycle, changing in length but not in diameter. This phenomenon is achieved by \n62 restricting growth to the cell poles, a process that is still not well understood. Growth occurring \n63 at the cell ends has been mainly studied in the transition from monopolar to bipolar growth, \n64 termed New End Take-Off (NETO) (4–7). NETO depends on specific polarity determinants, the \n65 kelch-repeat protein Tea1, the SH3 domain-containing protein Tea4 and the DYRK kinase, Pom1 \n66 among others (8–11). In the absence of Tea1–Tea4 complex, cells grow monopolarly but \n67 maintain their cylindrical shape. However, under certain stresses, the cells often choose the \n68 wrong growth site, forming bulged and T-shaped cells (10,12). \n69 Tea1 and Tea4 ride on growing microtubule (MT) plus ends to the cell tips, where they \n70 are released as discrete “dots” at the cortex, being Tea4 totally dependent on Tea1 for its \n71 location (12–15). At poles, the prenylated protein Mod5 and the ERM (Ezrin-Radixin-Moesin) \n72 family protein Tea3 anchor Tea1 to the cell cortex (16,17). Tea1 and Tea4 colocalize at the cell \n73 tips to form clusters or nodes, (18) and it is assumed that this association promotes the binding \n74 of other polarity factors in large protein complexes that organize polarized growth (4,10,19). \n75 One of these proteins is the formin For3, whose association with polarity markers likely brings \n76 it into the proximity of activators, stimulating the formation of F-actin cables that will deliver \n77 growth cargo to the tip (9). \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n4\n78 Establishing polarized growth involves hundreds of proteins; however, a constant from \n79 yeast to humans involves local accumulation of active GTP-bound forms of Rho-family GTPases \n80 at the cell cortex (20–22). While polarity is mostly associated with the functions of Rac and Cdc42 \n81 in mammals or Cdc42 in yeast, other GTPases such as RhoA or Rho1 (yeast) can also play a role \n82 in the development of polarity. Active RhoA is found at the leading edge as the edge advances \n83 in migrating cells, whereas Cdc42 and Rac1 are activated later (23). In budding and fission yeast, \n84 Cdc42 displays the ability to polarize spontaneously (22,24–26). However, the role of Rho1, the \n85 other essential GTPase, in polarized growth remains undefined. Depletion of Rho1 in growing \n86 cells induces shrinking and death via a kind of “apoptosis” that is accompanied by the \n87 disappearance of polymerized actin (27,28). Rho1 activity is regulated by three GEFs, Rgf1, Rgf2, \n88 and Rgf3 that catalyze the exchange of GDP for GTP, rendering the GTPase in an active state \n89 (28–34). The main activator of Rho1, Rgf1, regulates cell integrity through Rho1 by activating the \n90 β-glucan-synthase complex (28) and gene expression via the Pmk1 MAPK cell integrity-signaling \n91 pathway (28,35,36). Moreover, Rgf1, like Tea1, Tea4 and other polarity factors, is required for \n92 the actin reorganization necessary to switch from monopolar to bipolar growth during NETO \n93 (28). \n94 Here we have studied this phenomenon to show that Rgf1 interacts with the cell end \n95 marker Tea4 and binds to the plasma membrane (PM) through its PH domain. Both, PM binding \n96 and Rho-GEF activity are required for stable accumulation of polarity markers at the cell poles. \n97 In addition, we have uncovered a new role for Rgf1 in restricting growth to the poles in the \n98 absence of polarity markers. Most tea1Δ cells maintain their cylindrical morphology unless \n99 subjected to stresses, suggesting that these cells detect the location of its poles by an unknown \n100 mechanism. Here we show that this mechanism depends on the actin cytoskeleton and Rho1 \n101 activation by Rgf1. Therefore, we propose two parallel pathways to define the growth poles in \n102 fission yeast: the canonical one dependent on MTs and Tea1–Tea4 and another one dependent \n103 on actin and Rgf1–Rho1, both necessary to maintain a straight shape when the other is impaired. \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n5\n104\n105\n106 RESULTS\n107 Rgf1 is required for proper localization of the Tea1–Tea4 complex \n108 We have previously shown that rgf1Δ strain displays defects in bipolar growth, with ~80% of the \n109 cells showing monopolar pattern of growth compared to ~25% of the wild-type cells (28). This \n110 growth defect has been described in mutants affected in polarity factors such as Bud6, Tea1 or \n111 For3 (37–39), suggesting that similarly to these proteins, Rgf1 triggers NETO, and thus rgf1Δ cells \n112 may have problems when choosing the right end of growth. This prompted us to examine the \n113 localization of the landmark proteins Tea4 and Tea1 in cells lacking Rgf1. While Tea4-GFP was \n114 concentrated at both cell tips in wild-type cells (9,11), in rgf1Δ cells the signal detected was \n115 visibly diminished. We compared the fluorescence intensity of Tea4-GFP in wild-type and rgf1Δ \n116 cells in the same preparation, in which we grew tea4-GFP rgf1+ sad1-dsred (spindle pole body \n117 marker) cells and tea4-GFP rgf1Δ cells separately, and then mixed and imaged them at the same \n118 time (Fig 1A). In rgf1Δ cells, the Tea4-GFP signal was dispersed in small dots that spread out at \n119 the ends. The average fluorescence of Tea4-GFP dots at the tips of rgf1Δ cells was approximately \n120 half of that seen at the tips of the wild-type cells. Because the rgf1Δ cells grew in a monopolar \n121 fashion, we examined whether the Tea4 dots were more prominent at one end or whether they \n122 were scattered similarly at both ends. We observed that 61% of the  rgf1Δ cells accumulated \n123 Tea4 at the non-growing end (revealed by calcofluor staining) (Fig 1B). Thus, Rgf1 is more \n124 important to localize Tea4 to the growing tip, the one where Rgf1 concentrates in wild-type \n125 monopolar cells (Fig S1A). \n126\n127 Fig 1: Rgf1 is required for proper localization of the Tea1-Tea4 complex at the cell tip\n128 (A) Cells expressing tea4-GFP rgf1 + sad1-dsred and tea4-GFP rgf1Δ were grown in YES liquid \n129 medium separately, and then mixed and imaged in the same preparation. The maximum-\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n6\n130 intensity projection of six deconvolved Z-slides (0.5 µm step-size) is shown. The graphic \n131 represents the mean ± standard deviation (SD) of the relative fluorescence intensity of Tea4-\n132 GFP measured at the cell tips in the wild-type (WT, n = 47) and rgf1Δ (n = 41) cells. WT levels \n133 were used for normalization. (B) Calcofluor white (CW, 20 µg/mL) staining and GFP fluorescence \n134 in cells expressing tea4-GFP rgf1+ and tea4-GFP rgf1Δ. The maximum-intensity projection of six \n135 Z-slides (0.5 µm step-size) of Tea4-GFP fluorescence is shown. The red arrowheads indicate non-\n136 growing poles. The graphic represents the mean ± SD of the relative fluorescence intensity of \n137 Tea4-GFP measured at the tips of the rgf1Δ (n = 50) cells. Calcofluor staining, which marks sites \n138 of cell growth, was used to differentiate growth from non-growth poles (right). (C) \n139 Representative images of Tea4-GFP  (green) and Tea1-tdTomato  (red) localization in the rgf1Δ \n140 cells. The maximum-intensity projection of six Z-slides (0.5 µm step-size) is shown. (D) \n141 Representative images of the indicated cells expressing tea4-GFP (green) and mCherry-atb2 \n142 (red). The maximum-intensity projection of six Z-slides (0.5 µm step-size) is shown. (E) \n143 Kymographs of time-lapse fluorescence movies of tea4-GFP and mCherry-atb2 expressed in \n144 wild-type or rgf1Δ strains. The maximum-intensity projection of seven Z-slides (0.6 µm step-size) \n145 of images taken every 15 s was used to draw a line along an MT from the middle of the cell to \n146 the tip. The orange arrowheads indicate the moment of MT retraction. (F) Super-Resolution \n147 Radial Fluctuations (SRRF) images of Tea4-GFP (green) and mCherry-Atb2 (red) in “head-on” cell \n148 tips of the indicated strains. One focal plane image was taken every minute. The time projection \n149 of the three images at different time points is shown to follow Tea4 cluster movement (right). \n150 Note that in the wild-type strain Tea4 nodes remain stable (white arrowheads) for longer than \n151 in the rgf1Δ mutant (blue arrowheads). (G) Kymographs of time-lapse fluorescence movies of \n152 GFP-Atb2 producing in the WT and rgf1Δ cells. The graph shows the mean ± SD of the time during \n153 which the MT is touching the pole in both strains (n = 75). (H) Representative images of the \n154 indicated cells producing mCherry-Atb2. The maximum-intensity projection of six Z-slides (0.5 \n155 µm step-size) is shown. The graph shows the mean ± SD of the percentage of curved MTs found \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n7\n156 in the indicated strains (n > 500). Statistical significance was calculated using two-tailed unpaired \n157 Student’s t test. ****P < 0.0001; ns = non-significant. Scale bar 2 µm. \n158\n159 We also evaluated whether the localization of Tea1, which functions in a complex with \n160 Tea4 (9), is affected in the Rgf1 mutant. The wide co-localization between Tea1 and Tea4 in the \n161 absence of Rgf1 indicates that both proteins displayed similar localization defects (Fig 1C), and \n162 that similarly to wild-type cells, in rgf1Δ cells Tea1 and Tea4 were still bound and formed a \n163 complex. In tea4Δ cells, Tea1 is concentrated at the non-growing cell tip (9). Given that in rgf1Δ \n164 cells, Tea1 also localized mainly to the non-growing end (Fig 1B and C), our results suggest that \n165 the Tea1 mislocalization could be a consequence of Tea4 mislocalization. We noticed that rgf1Δ \n166 cells showed a greater number of Tea4 discrete dots in the middle zone of the cell (Figs 1A, 1B, \n167 and S1B). Examination of Tea4-GFP in wild-type and rgf1Δ cells with α–tubulin labeled in red \n168 (mCherry-Atb2) showed co-localization of Tea4 cytoplasmic dots with MTs (Fig 1D). Thus, in the \n169 rgf1Δ cells, a larger free cytoplasmic pool of Tea4, which is not properly sequestered at the poles, \n170 could now be available to be redirected to the cell cortex by MTs. \n171\n172 Rgf1 functions to integrate Tea4 in big clusters at the cell tip \n173 Next, we determined whether Tea4 was accurately delivered to the cell cortex in rgf1Δ cells by \n174 taking time-lapse images every 15 seconds. We did not observe appreciable differences in the \n175 delivery of Tea4 to the cell cortex between rgf1+ and rgf1Δ cells. However, the Tea4-GFP signal \n176 failed to remain in the pole in rgf1Δ cells (Movies S1 and S2). This can be better observed in the \n177 kymographs shown in Fig 1E, where the fluorescence of Tea4 vanished from the cell cortex of \n178 the rgf1Δ cells in a few seconds, whereas it remained stable in control cells. \n179 Polarity factors such as Tea1 and Tea4 localize to the cellular cortex in discrete clusters \n180 (18), which are not easily observable when taking conventional lateral cell images. To better \n181 perceive the formation of Tea4 nodes at the cell poles we used Super-Resolution Radial \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n8\n182 Fluctuations (SRRF) microscopy for “head-on” imaging of cell tips. We performed short time-\n183 lapse experiments (three minutes) at the cellular tip cortex on “head-on” wild-type and rgf1Δ \n184 cells (tagged with Tea4-GFP and mCherry-Atb2). In wild-type cells, Tea4 nodes deposited by MTs \n185 remained stable for an interval of time even after MT catastrophe (Fig 1F, white arrows). \n186 Moreover, clusters of Tea4 not associated with microtubules could be observed stable \n187 throughout the time-lapse, and were of the same size as those associated with MTs. However, \n188 in rgf1Δ cells, Tea4 dots of a similar size to those observed in the wild-type cells appeared \n189 exclusively while they were associated with MTs (Fig 1F,  blue arrows). Once the MT retracted, \n190 the Tea4 cluster became gradually smaller until it eventually disappeared. These observations \n191 confirmed the results obtained with conventional microscopy methods, suggesting that Rgf1 \n192 was not necessary for the delivery of Tea4 to the cell poles, but it is required for its stable \n193 maintenance once it was released there. In addition, we ruled out a defect in the stability of the \n194 Tea4 protein in the rgf1Δ mutant because the half-life of the protein in the wild-type and rgf1Δ \n195 cultures treated with cycloheximide was comparable (Fig S1C).\n196 During the course of these experiments, we noticed that some MTs curled around the \n197 tips in the rgf1Δ cells. We confirmed that the MT dynamics was not affected because the \n198 polymerization and depolymerization rates were similar in the wild-type and rgf1Δ cells, with a \n199 slight increase in the polymerization rate in the mutant (Fig S1D). However, the mean time that \n200 the MT stayed at the tip was ~43 seconds in the rgf1+ cells but ~70 seconds in the rgf1Δ cells (Fig \n201 1G). Therefore, once an MT reached the cortex in the rgf1Δ cells, it remained there longer than \n202 in the wild-type cells. Curved MTs have already been described in tea1Δ cells growing at a high \n203 temperature (10). We incubated the rgf1Δ and tea1Δ mutants at 37°C for 4 hours and observed \n204 MT organization under these conditions (Fig 1H). In the rgf1Δ mutant, ~10% of the cells \n205 possessed at least one MT curled around the end, similarly to the ~12% found in the tea1Δ \n206 mutant, while this type of curly MT was rarely observed in the wild-type (1.7%). It is possible \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n9\n207 that a limited amount of Tea4–Tea1 at the growing pole underlies the curly phenotype seen in \n208 the absence of Rgf1. \n209\n210 Rgf1 cooperates with Mod5 in Tea4 anchoring to the cellular poles.\n211 It has been shown that in cells lacking Mod5, Tea1 and Tea4 fail to accumulate to wild-type levels \n212 at the cell tips (9,16,40). Given that the rgf1Δ cells showed a similar defect, we analyzed the \n213 localization of Tea4 in the double mutant rgf1Δmod5Δ compared with the single mutants. In the \n214 mod5Δ and rgf1Δ cells, most of the Tea4-GFP dots were associated with MTs, although was still \n215 some signal mainly at one of the poles (Fig 2A, yellow arrows ). However, in the rgf1Δmod5Δ \n216 double mutant, the Tea4 signal was depleted at both poles  (Fig 2A, orange arrows), suggesting \n217 that Mod5 and Rgf1 share a function in Tea4 anchoring. \n218\n219 Fig 2: Rgf1 cooperates with Mod5 in Tea4 anchoring to the cellular poles\n220 (A) Representative images of the indicated strains producing Tea4-GFP (green) and mCherry-\n221 Atb2 (red). The maximum-intensity projection of six Z-slides (0.5 µm step-size) is shown. Scale \n222 bar 2 µm. (B) The percentage of cells forming branches in the indicated strains. The cells were \n223 grown to the stationary phase for 3 days, and then were treated with DMSO (-MBC) or MBC \n224 (+MBC; 50 µg/ml). The mean ± SD of > 200 cells from three independent experiments is shown. \n225 (C) The percentage of T-shaped cells in the indicated strains after incubation for 4 hours at 36°C \n226 in YES liquid medium. Statistical significance was calculated using a two-tailed unpaired \n227 Student’s t test. ns = non-significant.\n228\n229 Because the localization of Tea4 is entirely dependent on Tea1 and the localization of \n230 Tea1 is partially dependent on Tea4 (9), we wondered whether the phenotype of the \n231 rgf1Δmod5Δ cells (which mislocalize Tea4) resemble the tea1Δ phenotype. To this end, we \n232 scored the percentage of T-shaped cells in polarity re-establishment assays. In these assays, we \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n10\n233 first grew cells to the stationary phase for 3 days, and then diluted them in fresh medium for 3 \n234 hours. This treatment increases the penetrance of polarity mutant phenotypes, inducing T-\n235 shapes and bulges and was performed either in the presence or in the absence of the MT \n236 inhibitor methyl-2-benzimidazole carbamate (MBC), to prevent or allow the continuous delivery \n237 of Tea1–Tea4 to the poles by microtubules(12,16). Consistent with the “poor” localization of \n238 Tea4 in the cell cortex,  the rgf1Δ and mod5Δ cells showed polarity defects when treated with \n239 MBC and marginal defects in the absence of MBC, whereas ~80% of the tea1Δ cells displayed \n240 the characteristic T-shaped pattern, with or without MBC treatment (Fig 2B and S2A )(16). \n241 Interestingly, the rgf1Δmod5Δ cells behaved similar to the tea1Δ mutant: reaching ~80% of T-\n242 shaped cells with MBC and showing ~45% even in the presence of MT (-MBC) (Fig 2B and S2A). \n243 In addition, we combined the rgf1Δmod5Δ deletion with the temperature sensitive (ts) septation \n244 mutant cdc11-119. At the restrictive temperature, cdc11-119 cells show a defect in cytokinesis, \n245 but the nuclear and growth cycles continue and cells grow at both ends after each mitosis. \n246 Presumably, after each mitosis, cdc11-119 mutant cells must decide where to reinitiate growth. \n247 In the cdc11-119 tea1Δ double mutant, these events are often aberrant, leading to the \n248 formation of highly branched or T-shaped multinuclei cells (10,41). Only ~5% of the cdc11rgf1Δ \n249 and cdc11mod5Δ cells were T-shaped after incubation for 4 hours at 36°C ( Fig 2C and S2B). \n250 However, the cdc11rgf1Δmod5Δ triple mutant showed a similar percentage of T-shaped cells \n251 (~17%) as the cdc11tea1Δ double mutant (~21%). These results indicate that Rgf1 and Mod5 \n252 collaborate to position the polarity markers at the cell tips to prevent mislocalization of growth \n253 machinery in successive cell cycles.\n254\n255 Rgf1 interacts with the cell-end marker Tea4 and binds to phosphatidylinositol-4-phosphate \n256 through its PH domain.\n257 Tea1 and Tea4 reside in large protein complexes (10,19). We used different approaches to \n258 determine whether Rgf1 acts locally to retain Tea4 at the cell tips. First, we examined the in vivo \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n11\n259 localization of endogenous Tea4-GFP together with Rgf1-tdTomato. In newly divided and \n260 interphase cells, a subset of Rgf1 dots colocalized with Tea4 dots at cell tips, indicating a close \n261 proximity with each other (Fig 3A). Subsequently, we tested for the coprecipitation of these two \n262 proteins from yeast extracts by using epitope-tagged strains. Indeed, endogenously expressed \n263 GFP-tagged Tea4 led to the co-purification of HA-tagged Rgf1(Fig 3B). Tea1 forms a stable \n264 complex with Tea4 (9); however, we could not detect the interaction between Tea1 and Rgf1 \n265 (Fig S3). To validate these associations, we purified GST-Rgf1 from Escherichia coli and \n266 conjugated it with Glutathione Sepharose (GS) beads; then, we used those beads in a pull-down \n267 assay to trap Tea4-GFP or Tea1-GFP from Shizosaccharomyces pombe  protein extracts. We \n268 detected binding of Tea4 when using Rgf1-GS beads but not with GS beads alone  (Fig 3C). In \n269 addition, we could observe a slight precipitation of Tea1, which might be the result of Tea4 \n270 interaction with Tea1. We confirmed the biochemical interaction in a two-hybrid assay. \n271 Consistently, Rgf1 could interact with Tea4 but not with Tea1 (Fig 3D). Taken together, these \n272 results indicate that Rgf1 associates with the Tea1–Tea4 complex through its binding with Tea4. \n273\n274 Fig 3: Rgf1 interacts with the cell end marker Tea4 and binds to phosphatidylinositol-4-\n275 phosphate through its PH domain.\n276 (A) Colocalization of Rgf1 and Tea4. Representative images of wild-type cells producing Tea4-\n277 GFP endogenously (green) and Rgf1-tdTomato from a plasmid under the control of its own \n278 promoter (red). The maximum-intensity projection of six Z-slides (0.5 µm step-size) is shown. (B) \n279 Coprecipitation of Rgf1 and Tea4. Cell extracts from cells producing Tea4-GFP, Rgf1-HA, or Tea4-\n280 GFP and Rgf1-HA were precipitated with GFP-trap beads and blotted with anti-HA or anti-GFP \n281 antibodies (co-immunoprecipitation and immunoprecipitation). Western blot was performed on \n282 total extracts to visualize total Tea4-GFP and Rgf1-HA levels (whole cell extracts). (C) Cells \n283 expressing Tea1-GFP or Tea4-GFP were pulled down from cell extracts with GST-Rgf1 purified \n284 from E. coli bound to GS-beads or with GS-beads alone, and blotted with anti-GST or anti-GFP \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n12\n285 antibodies (Pull-down). Total Tea1-GFP or Tea4-GFP levels (WCE) were visualized by western \n286 blot; tubulin was used as a loading control. (D) Two-hybrid analysis of the interaction between \n287 Tea1 (pGR135) and Tea4 (pGR106) with Rgf1 (pRZ97). The interaction was assessed on YNB \n288 plates without histidine (YNB -H). (E) Protein-lipid overlay assay. Membrane lipid strips were \n289 overlaid with 1 ug/ml of the purified GST, GST-Rgf1, and GST-Rgf1ΔPH respectively, and the \n290 interaction was detected with an anti-GST antibody. Lipids to which GST-Rgf1 showed a \n291 significant association are shown in red. Note that the interaction with PI4P disappears with \n292 GST-Rgf1ΔPH. (F) Representative images of cells producing Rgf1-GFP  or Rgf1ΔPH-GFP. The \n293 maximum-intensity projection of six Z-slides (0.5 µm step-size) is shown. The graphic represents \n294 the mean ± SD of the relative fluorescence intensity measured at the cellular tips of Rgf1-GFP \n295 and Rgf1ΔPH-GFP (n>120). (G) Representative images of cells producing Rgf1-GFP in WT or efr3Δ \n296 mutant. The maximum-intensity projection of six Z-slides (0.5 µm step-size) is shown. The \n297 graphic represents the mean ± SD of the relative fluorescence intensity measured at the cellular \n298 tips of Rgf1-GFP (n > 120). (H) Protein extracts from cell producing Rgf1-GFP in the WT or efr3Δ \n299 mutant and Rgf1ΔPH-GFP were analyzed by western blot with an anti-GFP antibody to visualize \n300 Rgf1 levels. An anti-tubulin antibody was used as a loading control. The graphic represents the \n301 mean ± SD of the relative Rgf1 proteins levels from two independent experiments. Statistical \n302 significance was calculated using a two-tailed unpaired Student’s t test. ****P< 0.0001; ***P< \n303 0.001; **P < 0.01. Scale bar 2 µm.\n304\n305 Next, we evaluated whether Rgf1 acts as a linker between Tea4 and the PM in the \n306 anchoring process. Rgf1 is a large (~150 KDa) multi-domain protein, including a pleckstrin \n307 homology (PH) domain (34).  PH domains could act as a “membrane-targeting device” by \n308 anchoring GEFs to phosphoinositides and directing them towards their partner GTPases on the \n309 cellular cortex (42,43). To test the ability of Rgf1 to bind different membrane lipids, we fused \n310 the Rgf1 protein to GST (without its carboxi-terminal CNH domain to purify it more easily) and \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n13\n311 purified it from bacteria. We used membrane arrays spotted with different kind of lipids to \n312 detect binding of GST-Rgf1 to lipids. As shown in Fig 3E, recombinant-purified GST-Rgf1 \n313 preferentially bound to phosphatidic acid (PA) and cardiolipin and also bound to \n314 phosphatidylinositol 4-phosphate [PI(4)P], and 3-sulfogalactosylceramide more subtly. GST-\n315 Rgf1∆PH (additionally lacking the PH domain) could not interact with PI(4)P, but it behaved like \n316 the wild-type protein in terms of its binding to the other lipids. This result contrasts with that \n317 described for the budding yeast Rgf1 homolog, Rom2, whose PH domain specifically binds to \n318 phosphatidylinositol 4,5-bisphosphate [PI(4,5)P2] (44). Thus, in S. pombe Rgf1 might bind to the \n319 PM through the interaction between its PH domain with the phospholipid PI(4)P. Consistently, \n320 the PH domain was required for proper anchoring of Rgf1 to the cellular cortex. A GFP-tagged \n321 Rgf1∆PH mutant showed large defects in its localization at both the poles and the septum (Fig \n322 3F). The fluorescence detected at the rgf1∆PH-GFP cell ends was ~25% of that exhibited by the \n323 full-length protein Rgf1-GFP (Fig 3F, right).\n324 Kathleen Gould’s group previously reported that Rgf1 displays septum localization \n325 defects in cells lacking the gene efr3+, a PM scaffold for the PI(4)P kinase Stt4. Cells lacking efr3+ \n326 do not properly position Stt4 and display altered levels of PM phosphoinositides (45). We found \n327 that the localization of Rgf1 at the cell poles was compromised in the efr3∆ cells ( Fig 3G). \n328 Moreover, the PM binding of Rgf1 might be crucial for protein stability. We detected a 3–4-fold \n329 decrease in the protein level of Rgf1, both without the PH domain or in efr3∆ cells, when Rgf1 \n330 was not properly bound to the PM (Fig 3H). These results support a role for PM phospholipids \n331 in the anchoring of Rgf1 to the cellular cortex and in the maintenance of Rgf1 protein level. \n332\n333 Tea4 accumulation at the cell ends depends on Rgf1 anchoring to the PM and Rho1 activation.\n334 Next, we determined whether Tea4 is bound to the cell cortex in the rgf1∆PH-GFP mutant, which \n335 shows compromised Rgf1 localization. Cortical localization of Tea4 was greatly reduced (~35%) \n336 in the mutant lacking the PH domain compared with the wild-type (P < 0.0001)  (Fig 4A). \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n14\n337 However, in the rgf1∆PH-GFP mutant the ability to join and anchor Tea4 to the cortex could be \n338 reduced due to a significant drop in the protein level compared with the Rgf1-GFP protein (Fig \n339 3H). GTPase activation by its GEFs usually takes place when both the GTPase and the GEF are \n340 close to the PM; thus, a low level of Rgf1∆PH-GFP at the PM could promote inefficient activation \n341 of the Rho1 GTPase. To examine whether this is the case, we analyzed the in vivo amount of \n342 GTP-Rho1 (active-Rho1) in the rgf1∆PH cells in a pull-down assay with GST-C21RBD, the \n343 rhotekin-binding domain (previously purified from bacteria) (28). We found that the level of \n344 active-Rho1 detected in the rgf1∆PH cells was similar to that seen in the rgf1∆ cells, and much \n345 less than the amount detected in control cells (Fig 4B). Both, the localization of Rgf1 to the PM \n346 and the activation of Rho1 were impaired in the rgf1∆PH mutant; hence, we could not determine \n347 which one is behind Tea4 mislocalization. To distinguish between these two possibilities, we \n348 utilized the rgf1-ΔPTTR mutant expressing a protein without four amino acids in the RhoGEF \n349 domain. This mutant displays significantly reduced GEF activity toward Rho1,(36) while the \n350 protein remained attached to the growing end (Fig S4). Tea4-GFP was also mislocalized in the \n351 rgf1-ΔPTTR mutant (Fig 4A), indicating that the stable association of Tea4 to the membrane is \n352 dependent on Rgf1 GEF activity. \n353\n354 Fig 4: Tea4 accumulation at the cell ends depends on Rgf1 anchoring to the PM and Rho1 \n355 activation.\n356 (A) Representative images of Tea4-GFP localization in the WT, rgf1Δ, rgf1ΔPH, and rgf1ΔPTTR \n357 cells. The maximum-intensity projection of six Z-slides (0.5 µm step-size) of Tea4-GFP \n358 fluorescence is shown. Scale bar 2 µm. The graphic represents the mean ± SD of the relative \n359 fluorescence intensity of Tea4-GFP (n > 130) measured at the cellular tips. (B) Extracts from cells \n360 producing Rho1-HA (pREP4X-Rho1-HA) in the WT, rgf1Δ, and rgf1ΔPH cells were pulled down \n361 with GST-C21RBD and blotted against anti-HA antibody (Rho1-GTP). Total Rho1-HA was \n362 visualized by western blot (WCE). The relative units indicate the fold-differences in Rho1 levels \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n15\n363 in the mutants compared with the WT strain, with an assigned value of 1 (bottom) from three \n364 independent experiments. (C) Extracts from cells producing Rgf1-GFP or Rgf1ΔPH-GFP were \n365 pulled down with GST-Tea4 purified from E. coli bound to GS-beads or with GS-beads alone and \n366 blotted against anti-GST or anti-GFP antibodies (Pull-down). The total Rgf1 levels (WCE) were \n367 visualized by western blot; tubulin was used as a loading control. (D) Extracts from cells \n368 producing Rgf1-HA or Rgf1ΔPTTR-HA were pulled down with GST-Tea4 purified from E. coli  \n369 bound to GS-beads or with GS-beads alone and blotted against anti-GST or anti-GFP antibodies \n370 (Pull-down). Total Rgf1 levels (WCE) were visualized by western blot; tubulin was used as a \n371 loading control. (E) The percentage of cells WT, tea1Δ, rgf1Δ, rgf1ΔPH and rgf1ΔPTTR cells \n372 forming branches 3 hours after release to growth after 3 days in stationary phase, in the absence \n373 and in the presence of MBC (50 µg/ml). The mean ± SD of > 200 cells from two independent \n374 experiments is shown. Statistical significance was calculated using a two-tailed unpaired \n375 Student’s t test. ****P< 0.0001; **P < 0.01; *P < 0.05.\n376\n377 We wondered whether the  Rgf1-ΔPH and  Rgf1-ΔPTTR mutant proteins retained the \n378 ability to bind Tea4 in vitro. Both proteins proficiently bound Tea4 in an in vitro pull-down assay \n379 (Fig 4C and D). In vivo, we analyzed the percentage of T-shaped cells after re-entry from the \n380 stationary phase to fresh medium. With MBC treatment, the percentage of T-shaped cells was \n381 approximately 50% for the rgf1∆ mutant; this percentage was similar for the rgf1-ΔPH (~55%) \n382 and rgf1-ΔPTTR (~35%) mutants and much higher than for the wild-type strain (~5%) (Fig 4E). \n383 Taken together, these results indicate that the localization of Rgf1 to the PM and its ability to \n384 activate Rho1 are closely linked: Both are required to maintain Tea4 at the cell tips and to \n385 preserve the growth pattern after refeeding. Given that the Rgf1-ΔPTTR protein localized at the \n386 growing end, where it is incompetent for Rho1 activation, GEF activity appears to be more \n387 critical than GEF localization regarding Tea4 maintenance at the poles. Thus, Rho1 could act by \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n16\n388 promoting the formation of a stable critical mass of Tea4 in the cell cortex that is essential to \n389 activate pole-restricted growth.  \n390\n391 Rgf1 is part of actin-dependent machinery that signals growth poles in the absence of the \n392 Tea1–Tea4 complex\n393 As we have described above, tea1Δ cells experience an exacerbated loss of polarity when \n394 subjected to nutritional stress while keeping their cylindrical shape in normal conditions. We \n395 wondered why this occurs and how cells recognize their poles in the absence of the classical Tea \n396 markers. Since Tea4 localization is entirely dependent on Tea1(9) we utilized tea1Δ mutant to \n397 eliminate both markers at the poles. It has been proposed that the high number of T-shaped \n398 cells observed in the tea1Δ mutant in refeeding experiments (Fig 2B and 4E) is due to a transient \n399 depolarization of the actin cytoskeleton (46). To address this issue, we confirmed the \n400 disorganization of the actin cytoskeleton in wild-type cells grown stress treatments exposure to \n401 KCl, sorbitol or heat, which promoted actin depolarization also induced branching in the tea1Δ \n402 mutant for 3 days in liquid medium ( Fig 5A). In addition, ( Fig S5A and B) (11,47). We thought \n403 that if the mechanism that keeps the identity of the growth sites in the absence of Tea1 is lost \n404 because of transient actin depolarization, then treating tea1Δ cells with latrunculin A (LatA), \n405 which prevents the polymerization of filamentous actin, should increase the number of \n406 branched cells. As expected, 75% of the tea1Δ cells showed polarity defects during recovery \n407 from the LatA treatment, compared with ~2% of the wild-type and untreated tea1Δ cells (Fig \n408 5B). Thus, a properly polarized actin cytoskeleton is required to position growth sites at opposite \n409 cell poles in the absence of Tea1–Tea4 markers. \n410\n411 Fig 5: Rgf1 is part of an actin-dependent machinery that signals growth poles in the absence \n412 of the Tea1–Tea4 complex \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n17\n413 (A) Images of LifeAct-GFP (actin) localization in the WT cells growing in liquid medium in the log-\n414 phase or after 3 days in the stationary phase (s-phase). The maximum-intensity projection of six \n415 Z-slides (0.5 µm step-size) of fluorescence is shown. (B) Morphology and quantitation of the T-\n416 shaped wild-type and tea1Δ cells treated for 2 hours with DMSO (untreated) or 50 µM of \n417 Latrunculin A (LatA) and then washed to allow growth for 3 hours. The graph represents the \n418 mean ± SD of > 200 cells from three independent experiments. (C) Morphology and quantitation \n419 of the T-shaped cells in the WT, tea1Δ, tea1Δ mod5Δ, and tea1Δ rgf1Δ cells grown to log phase \n420 in YES liquid medium at 28°C. The graph represents the the mean ± SD of > 500 cells from two \n421 independent experiments. (D) Representative images of Tea1-GFP, Tea4-GFP, GFP-Mod5 and \n422 Rgf1-GFP localization in wild-type cells in the stationary phase after 3 days of growth in liquid \n423 medium. The maximum-intensity projection of six Z-slides (0.5 µm step-size) of fluorescence is \n424 shown. (E) Rgf1-GFP localization in WT cells growing in liquid medium untreated or treated with \n425 KCl 0.6M, sorbitol 1.2M or 37°C (heat) for 1 hour. The maximum-intensity projection of six Z-\n426 slides (0.5 µm step-size) of fluorescence is shown. The arrowheads point lateral accumulation of \n427 Rgf1-GFP. (F) LifeAct-mCherry (actin in red) and Rgf1-GFP localization (green) in tea1Δ cells \n428 treated with KCl 0.6M for 1 hour and then washed and allowed to grow without stress for the \n429 indicated times. The maximum-intensity projection of four Z-slides (0.6 µm step-size) of \n430 fluorescence is shown. The arrowheads point to lateral accumulation of Rgf1-GFP and actin (LA). \n431 (G) Morphology of the tea1Δ and gef1Δtea1Δ cells growing in YES liquid medium. (H) \n432 Quantitation of the T-shaped cells in WT and rgf1Δ cells treated with DMSO (-) or with MBC (50 \n433 µg/mL) for 4 hours. The graph represents the mean ± SD of > 200 cells from three independent \n434 experiments. (I) Morphology and quantitation of the T-shaped cells in wild-type cells treated \n435 with DMSO (WT -), MBC 50 µg/mL (WT MBC) for 4 hours, 50 µM of LatA for 2 hours and then \n436 washed and allowed to grow with DMSO for 4 hours (WT LatA) or first treated with LatA, washed, \n437 and then treated with 50 µg/mL of MBC for 4 hours (WT LatA-MBC). The graph represents the \n438 mean ± SD of > 200 cells from three independent experiments. Statistical significance was \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n18\n439 calculated using a two-tailed unpaired Student’s t test. ****P < 0.0001; ns = non-significant. \n440 Scale bar 2 µm.\n441\n442 Given that Rgf1 is required for actin re-organization during NETO (28), we reasoned that \n443 the absence of Rgf1 in the tea1Δ background could uncover polarity defects that would \n444 otherwise remain undetected. This was indeed the case; 50% of the  tea1Δ rgf1Δ cells in \n445 unperturbed conditions were T-shaped compared with fewer than 5% of the tea1Δ and \n446 tea1Δmod5Δ cells (Fig 5C). Thus, in the absence of Rgf1 and polarity markers (Tea1–Tea4), \n447 stresses that induce actin depolarization were not necessary for branching to occur. Moreover, \n448 Rho1 activation was required to maintain polarity, evidenced by the high number of T-shaped \n449 cells observed in the tea1Δrgf1ΔPTTR mutant (Fig S5C). Therefore, in the absence of Tea1, Rgf1–\n450 Rho1 probably mark the poles through an actin-dependent mechanism. \n451 To understand why the tea1Δrgf1Δ null cells behaved alike tea1Δ stressed cell, we \n452 studied the localization of Rgf1 under situations that depolarize the actin cytoskeleton. Rgf1-\n453 GFP localization to the cell tips was lost quickly in cells treated with LatA, but was unaffected in \n454 cells treated with MBC (Fig S5D). Thus, we reasoned that because actin is depolarized in \n455 quiescent cells, Rgf1-GFP should behave similarly. Accordingly, Rgf1 was missing from the cell \n456 periphery in stationary phase cells, while Tea1, Tea4, and Mod5 (which depend on MTs to reach \n457 the poles), remained polarized in the same experiment (Fig 5D) (10,11,16). Moreover, Rgf1 \n458 disappeared from the cell tips under osmotic or heat stress but was observed in lateral patches \n459 (Fig 5E). Then, we followed actin reorganization and Rgf1 localization during recovery from \n460 osmotic stress in wild-type and tea1Δ cells. After relieving the stress, wild-type cells quickly re-\n461 concentrated both actin and Rgf1 at the poles ( Fig S5E and Movie S3). However, in tea1Δ cells \n462 actin and Rgf1 frequently localized at the lateral cortex precisely at points where a branch began \n463 to grow (Fig 5F and Movie S4). \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n19\n464 To find out whether other proteins of the growth machinery also define the growth sites \n465 or if it is a specific characteristic of Rgf1, we used a mutant lacking gef1. Gef1 is a GEF of Cdc42 \n466 involved in polarized growth and undergoes a similar relocation from poles to lateral patches \n467 under stress, where it is required to activate Cdc42 (48,49). Interestingly, the gef1Δtea1Δ double \n468 mutant displayed a similar percentage of T-shaped cells as the tea1Δ mutant, indicating that in \n469 the absence of Tea1, Gef1 is not necessary for marking the poles, in contrast to Rgf1–Rho1 (Fig \n470 5G and S5F).\n471 Taken together, these results indicated that Rgf1 localization at the cell tips depends on a \n472 properly polarize actin cytoskeleton but it is independent on MTs. Thus, both actin \n473 concentration and Rho1 activation at the poles are necessary events to define these locations \n474 as growth sites.\n475\n476 Tea1–Tea4-MTs and Rgf1-Rho-actin define two parallel pathways to restrict growth to the cell \n477 tips\n478 Because stresses that induce actin disorganization and Rgf1 delocalization at the poles increase \n479 the percentage of T-shaped cells in the tea1Δ mutant (Fig 5D and S5A), we wondered whether \n480 chemical ablation of the pathway that delivers Tea1 and Tea4 to the cell tips would yield a similar \n481 result. When we treated the rgf1Δ mutant with MBC, ~30% of the cells exhibited a branched \n482 phenotype compared with ~5% of the wild-type cells (Fig 5H and S5G). Therefore, in the absence \n483 of Rgf1 (which lacks the “actin-dependent signal” necessary to recognize the polarity growth \n484 zones) and MTs, the imposition of stress is not necessary for branching to occur. The previous \n485 results suggest that there could be two parallel pathways for positioning the growth poles: one \n486 dependent on MTs and the Tea1–Tea4 markers, and another dependent on actin and Rgf1–\n487 Rho1. To mimic the elimination of both signaling pathways by chemical treatment, we treated \n488 wild-type cells in the log phase with LatA for 2 hours (to block actin-dependent signaling), \n489 washed them, and exposed them to MBC for 4 hours (to remove MTs) to analyze the number of \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n20\n490 T-shaped cells. Only after the treatment with both, LatA and MBC, ~35% of the cells showed \n491 branches (Fig 5I).  \n492 Our results indicate that Rgf1–Rho1 and Tea1–Tea4 are part of the same complex, with \n493 similar functions to delimit the sites of polarized growth of S. pombe. We propose that Rgf1 and \n494 Rho1 could activate an actin-dependent pathway that instructs cells to grow at the tips \n495 regardless of the classical polarity markers.\n496\n497\n498 DISCUSSION\n499 Cross-talk between microtubules (MT) and the actin cytoskeleton is crucial for various cellular \n500 processes, including asymmetric cell division, the establishment of cell growth zones, and cell \n501 migration. In fission yeast, MTs deliver the Tea1–Tea4 complex to the cell tips, where actin \n502 concentrates to promote growth. Tea1–Tea4 act as end markers, defining the organization of \n503 cell-growth zones and, consequently, the direction of growth. While it is known that these \n504 polarity markers are not essential for organizing a growth zone, they become critical for placing \n505 the growth zone correctly, especially under stress conditions. However, certain questions \n506 remain unanswered, such as how and why the Tea1–Tea4 complex remains linked to the plasma \n507 membrane (PM) once the growth direction is established and how cells mark their tips in the \n508 absence of Tea1. In the present study, we addressed some of these questions by demonstrating \n509 that Rgf1 functions as a molecular link between the Tea1-Tea4 complex and the PM. Through \n510 Rho1 activation, Rgf1 stabilizes Tea4 at the cell ends, promoting its accumulation. Additionally, \n511 we described an alternative actin-dependent mechanism, driven by Rgf1 and Rho1, for marking \n512 the poles independently to the known MT- and Tea-dependent pathway.\n513\n514 Rgf1 (Rho1 GEF) activity toward Rho1 is required for stable accumulation of Tea4 at the cell \n515 ends \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n21\n516 Failure to accumulate Tea4 at the cell cortex in the rgf1Δ cells is independent of the movement \n517 of Tea4 riding on the tips of polymerizing MTs. Wild-type and rgf1Δ cells exhibited similar rates \n518 of movement for Tea4-GFP dots from the middle of the cell to the cell ends (Fig S1D). However, \n519 once the Tea4-GFP dots had reached the cell ends, their fluorescence faded in the rgf1Δ cells. \n520 The refill mechanism responsible for keeping Tea4 stacked at the growing pole depends on \n521 correct binding of Rgf1 to the PM and on Rgf1 catalytic activity toward Rho1. We provided \n522 evidence that Rgf1 physically binds the polarity marker complex through its interaction with  \n523 Tea4 (Fig 3B-D). Moreover, Rgf1 binds to PM phospholipids, likely through its PH domain \n524 interacting with membrane PI4P. The physical association between Rgf1 and Tea4 is crucial for \n525 the localization of Tea4 and Tea1 at the cell poles. Indeed, a mutant of Rgf1 lacking the \n526 membrane-binding domain, which binds Tea4 in vitro, exhibited similar polarity defects as the \n527 null mutant (Fig 4E). In the  rgf1ΔPH mutant, the reduced activation of Rho1 (Fig 4B) is coupled \n528 to protein instability (Fig 3H) (probably because it is not properly bound to the PM) which makes \n529 the interpretation of this result challenging. Additionally, a catalytic mutant (rgf1-ΔPTTR) also \n530 shows impaired Tea4 anchoring. The Rgf1-ΔPTTR protein binds Tea4 in vitro, localizes to the \n531 growing end and is catalytically deficient (36), indicating that Rho1 activity is required to \n532 maintain a the Tea1-4 complex stable at the pole. Furthermore, the tandem Rgf1–Rho1 also \n533 affects Tea1 functions, promoting MT catastrophe once they reach the cell pole or restricting \n534 growth sites after refeeding (Figs 1G-H and 2B). To maintain polarity markers stably at the poles, \n535 Rgf1 functions together with Mod5. The loss of Tea4 localization at the poles is partial in the \n536 rgf1Δ and mod5Δ single mutants and complete when both mutations are combined, causing a \n537 defect in polarity even in cells with a continuous supply of Tea1-Tea4 (Fig 2B and C) . It is likely \n538 that Mod5 and Rgf1 anchor the Tea1-4 complex to the PM through different proteins: Mod5 \n539 interacts with Tea1 (17), while Rgf1 interacts with Tea4. This mechanism ensures double \n540 anchoring of the Tea1–Tea4 complex to the PM; thus, when one of these connections is lost, the \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n22\n541 other remains available. Only when both are missing do the polarity markers completely lose \n542 their connection with the PM, mimicking the behavior of a the tea1Δ mutant.\n543\n544 Rgf1–Rho1 are involved in defining the growth sites\n545 Another question that must be addressed is how cells detect their growth sites in the absence \n546 of Tea markers. Here we proposed that actin and Rho1 activation by Rgf1 are behind this \n547 process. We observed two ways to induce the formation of branched cells in the tea1Δ \n548 background: one in the presence of certain stresses, “the stress pathway” and the other in the \n549 absence of rgf1, “Rgf1 depletion” (Fig 5C and S5B). The loss of polarity induced by “the stress \n550 pathway” also leads to disorganization of the actin cytoskeleton and, consequently, Rgf1 \n551 delocalization. Therefore, both pathways lead to insufficient activation of Rho1 at the poles. \n552 When the tea1Δ cells recover from stress, Rgf1 and actin appear simultaneously at sites \n553 were ectopic growth occurs. The interdependence of actin and Rgf1 localization (Fig S5D) (28) \n554 suggests a positive feedback loop between Rho1 activation and actin organization at the growth \n555 sites. Therefore, Rgf1–Rho1 would act as Tea markers, defining growth sites without directly \n556 promoting growth, given that the tea1Δrgf1Δ double mutant retains the ability to grow, albeit \n557 at the wrong places. Interestingly, in the absence of Tea1 and Gef1 (Cdc42 GEF), which is also \n558 involved in polarized growth and relies on actin for proper localization (48–50), cells do not form \n559 branches, suggesting that is the activation of Rho1 and not that of Cdc42 which specifically \n560 defines the growth sites. \n561 The interaction between Tea4 and Rgf1–Rho1 indicates their involvement in the same \n562 complex, with each protein essential for the accumulation of the other at the poles. Rgf1 \n563 together with Mod5 acts to link Tea4 to the membrane, and Tea1-4 assists in returning Rgf1 to \n564 the poles after stress-induced actin disorganization (Fig 5F and S5E ), establishing a functional \n565 link between MTs and actin cytoskeletons (Fig 6A).\n566\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n23\n567 Fig 6: Rgf1-Rho1 functions as a molecular link between Tea4 and the PM and marks the growth \n568 sites in an actin-dependent manner\n569 (A) In wild-type cells MT deliver Tea1 and Tea4 to the cell poles, where they bind to the \n570 membrane through their interaction with Mod5 and Rgf1, respectively. Rgf1, in turn, interacts \n571 with the PM due to its affinity for the phospholipid PI4P and activates Rho1, promoting proper \n572 actin cytoskeleton polarization at the cellular tips. (B) Cells lacking Tea1 still recognize the cell \n573 poles correctly because Rgf1 is localized in these regions, where it activates Rho1, thus allowing \n574 the maintenance of polarized actin. (C) In rgf1Δ cells Tea1-Tea4 partially disappears from the \n575 pole, although a remnant remains attached to Mod5. Rho1 would be inactive, leading to actin \n576 cytoskeleton disorganization. However, the continuous supply of Tea1–Tea4 from MTs persists, \n577 allowing the cells to grow correctly. (D) When cells lack tea1 and rgf1, they lose both pathways \n578 that allowed them to distinguish the tips. First, there is no continuous supply of polarity markers \n579 towards the cell ends by the MTs that mark the growth sites. Second, Rgf1 is not at the pole to \n580 activate Rho1, leading to actin disorganization. The elimination of both pathways causes the \n581 cells to direct their growth towards incorrect locations, where growth factors probably \n582 accumulate. \n583\n584 We showed that Rgf1 and Rho1 are required to maintain an actin-dependent signal that \n585 preserves the identity of the cell poles, which becomes evident in the absence of the Tea \n586 markers. This would explain why the rgf1Δtea1Δ double mutant forms T-shaped cells in the log \n587 phase, without stresses or refeeding treatments. We propose that there are two different \n588 pathways to choose the growth sites under different environmental conditions: the canonical \n589 pathway dependent on MT and Tea1–Tea4 and, a novel pathway dependent on actin and Rgf1–\n590 Rho1. Various combinations of mutants and/or chemical ablation of one component from each \n591 pathway (Tea1–Tea4–MT and Rgf1–Rho1–actin) at once, leads to comparable outcomes.  For \n592 example, chemical actin depolymerization in the tea1Δ cells induces branching (Fig 5B), whereas \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n24\n593 treatment of the rgf1Δ cells with MBC to prevent the constant supply of Tea markers generates \n594 T-shaped cells as well (Fig 5H). Furthermore, wild-type cells treated to transiently depolarize \n595 actin and subsequently prevented for refeeding of polarity markers to the tips (via MT de-\n596 polymerization) also experience difficulties in detecting the cell poles (Fig 5I). Thus, cells would \n597 have different pathways to maintain their cylindrical morphology even if one of them is \n598 challenged by internal or external conditions. When the continuous supply of Tea1 from MT is \n599 disrupted (Fig 6B), cells could use an additional cue provided for a different cytoskeletal polymer, \n600 actin. Similarly, when actin becomes disorganized, for example, in the transition from \n601 monopolar to bipolar growth (NETO) or in rgf1Δ mutant (defective in NETO), cells would count \n602 on polarity markers transported by MTs (Fig 6C). Only when both, the actin and MT \n603 cytoskeletons are compromised, fission yeast cells lose their cylindrical shape (Fig 6D). This \n604 sophisticated regulation highlights the importance of maintaining cell morphology throughout \n605 the cell cycle and under changing environmental conditions.\n606\n607\n608 MATERIALS AND METHODS \n609 Media, Reagents and Genetics\n610 S. pombe strains were streaked on plates of complete yeast growth medium (YES), or selective \n611 medium (EMM) supplemented with the appropriate requirements (51), and incubated at 28°C \n612 until colonies formed. For each biological replicate, a single colony was used to inoculate 5 mL \n613 of the respective liquid media. Cultures were incubated at 28°C overnight with shaking (200 \n614 rpm). Each overnight culture was subsequently used as a seed culture to inoculate fresh media. \n615 Fresh cultures were next grown at 28°C, 200 rpm, to OD 600 = 0.5-0.6 at the time of harvest. \n616 Crosses were performed by mixing the appropriate strains directly on sporulation medium \n617 plates. Recombinant strains were obtained by tetrad analysis or the “random spore” method. \n618 For overexpression experiments using the nmt1 promoter, cells were grown in EMM containing \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n25\n619 15 µM thiamine up to the logarithmic phase. Then, the cells were harvested, washed three times \n620 with water, and inoculated in fresh medium (without thiamine) at an optical density at 600 nm \n621 (OD600) of 0.01 for 18-20 hours. Two-hybrid interaction was tested with YNB medium lacking \n622 histidine in Saccharomyces cerevisiae strain AH109 (Clontech).\n623\n624 Plasmid and DNA manipulations\n625 Plasmids used in this study are listed in Key resources table (Recombinant DNA). pREP4x-HArho1 \n626 (with thiamine-repressible nmt1 promoter) and pGEX-C21RBD plasmids (rhotekin-binding \n627 domain) kindly provided by Pilar Pérez (Instituto de Biología Funcional y Genómica, Salamanca, \n628 Spain) were used to detect Rho1-GTP levels. To express proteins in Escherichia coli we used a \n629 pGEX-2T plasmid that contains a GST to tagged genes al 5’. pGEX-rgf1  (pGR152) and pGEX-tea4 \n630 (pGR129) were made by inserting the entire ORF of rgf1 (without introns) or tea4 in frame into \n631 pGEX-2T and purified for be used in pull-down assays. To perform lipid strips assays we \n632 constructed the plasmids pGR128 that contains the ORF of rgf1 (from aminoacid 1-922) without \n633 the last 428 aminoacids containing the CNH domain, and pGR138 that contains the ORF of rgf1 \n634 (from aminoacid 1-722) without the last 578 aminoacids containing the PH and CNH domains. \n635 For two-hybrid experiments we constructed the plasmids pGAD-tea1 (pGR135), pGAD-tea4 \n636 (pGR106), and pGBK-rgf1 (pRZ97) where the entire ORF of the corresponding gene was inserted \n637 in frame into the pGADT7 (GAL4 activation domain) or pGBKT7 (GAL4 binding domain) plasmid \n638 (Clontech).\n639\n640 Protein extracts and immunoblot analysis\n641 S. pombe cultures (5 mL) at an OD600 of 0.5 were pelleted just after the addition of 10% TCA and \n642 washed in 20% TCA. The pellets were resuspended in 100 μl 12.5% TCA with the addition of glass \n643 beads and lysed by vortexing for 5 min. Cell lysates were pelleted, washed in iced acetone, and \n644 dried at 55°C for 15 min. Pellets were resuspended in 50 μl of a solution containing 1% SDS, 100 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n26\n645 mM Tris–HCl (pH 8.0), and 1 mM EDTA. Samples were electrophoretically separated by SDS-\n646 PAGE (4–15% MiniProtein Gel, BioRad) and immunodetected with anti-GFP (Living Colors, \n647 RRID:AB_10013427) and anti-mouse (Bio-Rad, AB_11125547) antibodies. As a loading control, \n648 we used monoclonal antitubulin antibodies (Sigma, RRID:AB_477579).\n649\n650 Immunoprecipitations and pull-down assays\n651 Immunoprecipitation assays were performed as described previously with some modifications \n652 (52). Briefly, logarithmic cultures were pelleted and re-suspended in lysis buffer (10 mM Tris–\n653 HCl pH 7.5, 150 mM NaCl, 0.5 mM EDTA, 0.5% NP40, containing 100 μM PMSF, leupeptin, and \n654 aprotinin) and lysed in a cryogenic grinder. Lysates were centrifuged for 5 min at 6000 g, and \n655 then 10 μL of GFP-Trap magnetic beads (Chromotek) was added to the supernatants an \n656 incubated for 1 hour at 4°C. Immunoprecipitates were washed three times with dilution buffer \n657 (10 mM Tris–HCl pH 7.5, 150 mM NaCl, and 0.5 mM EDTA). Proteins were released from \n658 immunocomplexes by boiling for 5 minutes in sodium dodecyl sulfate (SDS) loading buffer. \n659 Samples were separated by SDS–polyacrylamide gel electrophoresis (4–15% Mini-Protean TGX \n660 gels, Bio-Rad) and detected by immunoblotting with polyclonal anti-HA (Roche, \n661 RRID:AB_514506) or anti-GFP antiserum (Living Colors, RRID:AB_10013427). For pull-down \n662 assays, first GST-tagged Rgf1, Tea4, or C21RBD (rhotekin-binding domain to detect Rho1-GTP \n663 levels) was purified from E. coli (BL21). The fusion proteins were produced by adding 0.5 mM \n664 IPTG at 18°C overnight (or 3 hours at 28°C for C21RBD). Cells were sonicated and proteins \n665 immobilized on glutathione-Sepharose (GS) 4B beads (GE-Healthcare). After incubation for 1 \n666 hour, the beads were washed several times, and the bound proteins were analyzed by SDS-PAGE \n667 and stained with Coomassie brilliant blue. Pull-down assays were performed as described \n668 previously (53). In brief, extracts from the indicated strains were obtained by using 500 μL of \n669 lysis buffer (50 mM Tris–HCl pH 7.5, 20 mM NaCl, 0.5% NP-40, 10% glycerol, 0.1 mM \n670 dithiothreitol, and 2 mM Cl2Mg, containing 100 μM PMSF, leupeptin, and aprotinin) and lysed in \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n27\n671 a cryogenic grinder. Cell extracts (2–3 mg of total protein) were incubated with 2–10 μg of GST-\n672 tagged protein coupled to GS beads for 2 hours, washed four times with lysis buffer, and blotted \n673 with an anti-HA or anti-GFP antibody. Protein levels in whole-cell extracts (80 μg of total protein) \n674 were monitored by western blot. Tubulin and GST were used as loading controls. \n675\n676 Lipid strip overlay assays\n677 Lipid strip overlay assays were performed as described previously (54) using lipid strip \n678 membranes (p-6002, Echelon). Strips were blocked with 3% fatty acid–free bovine serum \n679 albumin (BSA; Sigma) in TBS-T (10 mM Tris–HCl pH 8.0, 150 mM NaCl, and 0.1% Tween-20; TBST-\n680 BSA) at room temperature for 1 hour and then incubated for 2 hours with 1 μg/mL GST, GST-\n681 Rgf1, or GST-Rgf1ΔPH in TBST-BSA. The strips were then washed three times with 5 mL of TBST-\n682 BSA and incubated with anti-GST horseradish peroxidase–conjugated antibody (GE Healthcare, \n683 RRID:AB_771429) diluted in TBST-BSA. Bound protein was detected using an enhanced \n684 chemoluminescence detection kit (BioRad). GST, GST-Rgf1, and GST-Rgf1ΔPH were expressed in \n685 E. coli (BL21) and purified with GS beads (GE-Healthcare) according to the manufacturer’s \n686 instructions as described above. Once attached to the beads, they were washed and eluted with \n687 200 μL of elution buffer (100 mM Tris–HCl pH 8.0, 120 mM NaCl) with 20 mM of L-glutathione \n688 reduced (Sigma) freshly added for 30 minutes at 4°C. Aliquots were frozen in liquid nitrogen with \n689 15% of glycerol and stored at -80°C until use. \n690\n691 Microscopy\n692 Wet preparations were observed with an Andor Dragonfly 200 Spinning-disk confocal \n693 microscope equipped with a sCMOS Sona 4.2B-11 camera (Andor) and controlled with the \n694 Fusion 2.2 acquisition software; or a Personal Deltavision (Applied Precision, LLC) microscope \n695 equipped with a CoolSNAP HQ2 camera (Photometrics) and controlled with softWoRx Resolve \n696 3D. Depending on the experiment, a single focal plane at the centre of the cell or a stack of 4–6 \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n28\n697 images covering the entire volume of the cell (Z-series) with a spacing of 0.5–0.6 μm were \n698 captured, and the maximum projection was generated.\n699 To analyze protein dynamics, time-lapse experiments were performed with cells in μ-\n700 Slide 8-well (Ibidi) coated with soybean lectin (Sigma Aldrich) and imaged at the indicated times \n701 using an Andor Dragonfly Spinning-disk confocal microscope. We used the ImageJ 1.53t software \n702 to calculate the relative fluorescence intensity of each protein at the cell tips. We have designed \n703 an ImageJ macro to automatically select the fluorescence regions (ROI manager) of the cell poles \n704 and to measure the fluorescence intensity of 40–130 cells. We used the integrated density value \n705 of each tip of the different strains versus the average of the integrated density of the wild-type \n706 strain to calculated the relative fluorescence levels. To create kymographs, we drew a line from \n707 the cell’s centre to the pole along a microtubule on a time-lapse movie and utilized the \n708 KymographBuilder plugin of the ImageJ software. For super-resolution radial fluctuations (SRRF) \n709 images of Tea4-GFP (green) and mCherry-Atb2 (red) in “head-on” cell tips, cells were mounted \n710 onto a μ-Slide pre-coated with lectin. Bound cells found to be frontally arrayed (“head-on”) were \n711 visually selected for imaging using a the SRRF module of the Andor Dragonfly Confocal \n712 microscope. Fifty repetitions of one focal plane were taken for each time point. The time \n713 projection of the three images at different time points is shown to follow Tea4 cluster \n714 movement. \n715 Calcofluor white (Blankophor BBH, Bayer Corporation) staining was performed by \n716 adding 1 μL of a stock solution (2.5 mg/mL) to 500 μL of samples for 20 seconds, followed by a \n717 wash with phosphate–buffered saline (PBS). \n718\n719 Quantification and statistical analysis\n720 Statistical analyses and graphs were generated using GraphPad Prism Software version 9.5.1. To \n721 compare two conditions, a two-tailed unpaired Student’s t-test was applied to determine \n722 statistical significance (as detailed in the Fig legends). P < 0.05 were considered significant. The \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n29\n723 graphs show the mean ± standard deviation of the indicated data. Asterisks represent the \n724 following: *P < 0.05 **P < 0.01; ***P < 0.001 ****P < 0.0001.\n725\n726 Author contributions\n727 Conceptualization, P.G., and Y.S.; Methodology, P.G.; Investigation, P.G., R.C. and T.E.; Writing – \n728 Original Draft, P.G.; Writing – Review & Editing, P.G., and Y.S.; Funding Acquisition, Y.S.; \n729 Supervision, P.G. and Y.S.\n730\n731 Acknowledgments\n732 We thank J.C. Ribas, Phong T. Tran, Sophie Martin, Sergio Rincón, Ken Sawin, Paul Nurse, James \n733 Moseley, Kathleen L. Gould, César Roncero, Henar Valdivieso, Sergio Moreno and Pilar Pérez for \n734 sharing strains and plasmids. We also wish to thank Javier Encinar del Dedo and Sergio Rincón \n735 for their very helpful comments on the manuscript. We are grateful to Jesús Pinto (IBFG \n736 bioinformatics facility) for ImageJ macro used for fluorescence quantification. \n737\n738\n739 REFERENCES\n740 1. Allam AH, Charnley M, Russell SM. Context-Specific Mechanisms of Cell Polarity \n741 Regulation. J Mol Biol. 2018 Sep;430(19). \n742 2. Drubin DG, Nelson WJ. Origins of cell polarity. Cell. 1996;84:335–44. \n743 3. Hoffman CS, Wood V, Fantes PA. An Ancient Yeast for Young Geneticists: A Primer on \n744 the Schizosaccharomyces pombe Model System. Genetics [Internet]. 2015 \n745 Oct;201(2):403 LP-- 423. Available from: \n746 http://www.genetics.org/content/201/2/403.abstract\n.CC-BY 4.0 International licenseperpetuity. 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Stress-\n877 dependent inhibition of polarized cell growth through unbalancing the GEF/GAP \n878 regulation of Cdc42. Cell Rep. 2021 Nov;37(5). \n879 49. Coll PM, Trillo Y, Ametzazurra A, Perez P. Gef1p, a new guanine nucleotide exchange \n880 factor for Cdc42p, regulates polarity in Schizosaccharomyces pombe. Mol Biol Cell \n881 [Internet]. 2003;14(1):313–23. Available from: \n882 https://www.ncbi.nlm.nih.gov/pubmed/12529446\n883 50. Hirota K, Tanaka K, Ohta K, Yamamoto M. Gef1p and Scd1p, the Two GDP-GTP \n884 exchange factors for Cdc42p, form a ring structure that shrinks during cytokinesis in \n885 Schizosaccharomyces pombe. Mol Biol Cell [Internet]. 2003;14(9):3617–27. Available \n886 from: https://www.ncbi.nlm.nih.gov/pubmed/12972551\n887 51. Moreno S, Klar A, Nurse P. [56] Molecular genetic analysis of fission yeast \n888 Schizosaccharomyces pombe. In 1991. p. 795–823. Available from: \n889 https://linkinghub.elsevier.com/retrieve/pii/007668799194059L\n890 52. Calvo IA, García P, Ayté J, Hidalgo E. The transcription factors Pap1 and Prr1 collaborate \n891 to activate antioxidant, but not drug tolerance, genes in response to H <inf>2</inf>O \n892 <inf>2</inf>. Nucleic Acids Res. 2012;40(11). \n893 53. García P, Coll PM, del Rey F, Geli MI, Pérez P, Vázquez de Aldana CR, et al. Eng2, a new \n894 player involved in feedback loop regulation of Cdc42 activity in fission yeast. Sci Rep. \n895 2021;11(1). \n896 54. Fernández-Golbano IM, Idrissi FZ, Giblin JP, Grosshans BL, Robles V, Grötsch H, et al. \n897 Crosstalk between PI(4,5)P2 and CK2 Modulates Actin Polymerization during Endocytic \n898 Uptake. Dev Cell. 2014 Sep;30(6). \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n36\n899  \n900 Supporting Information\n901 S1 Table. List of yeast strains used in this study.\n902 S2 Table. List of plasmids used in this study.\n903 S1 Fig. Rgf1 is required for proper localization of the Tea1-Tea4 complex at the cell tip. \n904 (A) Wild-type cells expressing rgf1-GFP were stained with Calcofluor white (20 μg/mL) to spot \n905 areas of growth. The arrows indicate the localization of Rgf1 at the growing tip in monopolar \n906 cells. (B) Quantitation of the number of Tea4 dots associated to the MTs in WT and rgf1Δ strains \n907 per cell. The mean ± SD of > 80 cells is shown. (C) The wild-type and rgf1Δ cells expressing tea4-\n908 GFP were treated with the translation inhibitor cycloheximide (CHX, 100 μg/mL) for the \n909 indicated times. Proteins were visualized by western blot with antibodies against GFP (Tea4) or \n910 tubuline (Tub), as a loading control (upper). The graphic represents the quantification of Tea4 \n911 levels at different times (hours) after the treatment relative to time 0, which was assigned a \n912 value of 1 (bottom). (D) The graphics show the MT polymerization (left) and depolymerization \n913 (right) rates in wild-type and rgf1Δ cells. The mean ± SD of > 75 cells is shown. Statistical \n914 significance was calculated using two-tailed unpaired Student’s t test. *P < 0.05; ****P < 0.0001; \n915 ns= non-significant.\n916 S2 Fig. Rgf1 cooperates with Mod5 in Tea4 anchoring to the cellular poles. \n917 (A) Morphology of the wild-type, rgf1Δ, mod5Δ and rgf1Δmod5Δ strains after refeeding \n918 treatment (-MBC). Scale bar 2 μm. (B) Cell morphology of the indicated strains after 4 hours at \n919 36°C in YES liquid medium. Scale bar 2 μm.\n920 S3 Fig. Rgf1 interacts with the cell end marker Tea4 and binds to phosphatidylinositol-4-\n921 phosphate through its PH domain. \n922 Coprecipitation of Rgf1 and Tea1. Cell extracts from cells producing Tea1-GFP, Rgf1-HA, and \n923 Tea1-GFP and Rgf1-HA were precipitated with GFP-trap beads and blotted with anti-HA or anti-\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n37\n924 GFP antibodies (co-immunoprecipitation and immunoprecipitation). Western blot was \n925 performed on total extracts to visualize total Tea1-GFP and Rgf1-HA levels (whole cell extracts).\n926 S4 Fig. Tea4 accumulation at the cell ends depends on Rgf1 anchoring to the PM and Rho1 \n927 activation. \n928 Representative images of Rgf1-GFP and Rgf1ΔPTTR-GFP localization. Cells were stained with \n929 calcofluor white (20 μg/mL) to spot areas of growth. Scale bar 2 μm.\n930 S5 Fig. Rgf1 is part of actin-dependent machinery that signals growth poles in the absence of \n931 Tea1–Tea4 complex. \n932 (A) Representative images of LifeAct-GFP (actin) localization in cells untreated or treated with \n933 KCl 0.6 M, sorbitol 1.2 M, or 37°C (heat) for 1 hour. The maximum-intensity projection of six Z-\n934 slides (0.5 μm step-size) of fluorescence is shown. (B) Quantitation of the T-shaped cells in the \n935 tea1Δ mutant treated with DMSO (Unt.), KCl 0.6 M, sorbitol 1.2 M, or 37°C (heat) for 1 hour, \n936 then washed and allowed to grow without the drug for 3 hours. The graph represents the mean \n937 ± SD of > 200 cells from two independent experiments. (C) Quantitation of T-shaped wild-type, \n938 tea1Δ, tea1Δ rgf1Δ, and tea1Δ rgf1ΔPTTR cells grown to log phase in YES liquid medium at 28°C. \n939 The graph represents the mean ± SD of > 200 cells from two independent experiments. (D) Cells \n940 expressing rgf1-GFP and crn1-GFP (actin patches) or mCherry-atb2 (microtubules) cultured \n941 separately; mixed; and then treated with DMSO, LatA 100 μM, or MBC 50 μM for 15 minutes. \n942 (E) LifeAct-mCherry (actin) and Rgf1-GFP localization in wild-type cells treated with KCl 0.6 M for \n943 1 hour and then washed and allowed to grow without stress for the indicated times. The \n944 maximum-intensity projection of four Z-slides (0.6 μm step-size) of fluorescence is shown. \n945 Statistical significance was calculated using a two-tailed unpaired Student’s t test. **P < 0.01; \n946 ****P < 0.0001. Scale bar 2 μm.\n947 S1 Movie. Tea4 and microtubule dynamics in wild-type cells. Tea4-GFP (green) and mCherry-\n948 Atb2 (red) localization in wild-type cells. Protein dynamics was followed for 8 minutes, with \n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n38\n949 pictures taken every 20 seconds. The maximum-intensity projection of six Z-slides (0.5 µm step-\n950 size) is shown. \n951 S2 Movie. Tea4 and microtubule dynamics in rgf1Δ cells. Tea4-GFP (green) and mCherry-Atb2 \n952 (red) localization in rgf1Δ cells. Protein dynamics was followed for 8 minutes, with pictures taken \n953 every 20 seconds. The maximum-intensity projection of six Z-slides (0.5 µm step-size) is shown. \n954 S3 Movie. Rgf1 and actin dynamics in wild-type cells during osmotic stress recovery. Rgf1-GFP \n955 (green) and LifeAct-mCherry (actin in red) localization in wild-type cells treated with KCl 0.6 M \n956 for 1 hour and then washed and allowed to grow without stress. Protein dynamics was followed \n957 for 42 minutes, with pictures taken every 3 minutes. The maximum-intensity projection of four \n958 Z-slides (0.6 µm step-size) is shown. \n959 S4 Movie. Rgf1 and actin dynamics in tea1Δ cells during osmotic stress recovery. Rgf1-GFP \n960 (green) and LifeAct-mCherry (actin in red) localization in tea1Δ cells treated with KCl 0.6 M for 1 \n961 hour and then washed and allowed to grow without stress. Protein dynamics was followed for \n962 66 minutes, with pictures taken every 3 minutes. The maximum-intensity projection of four Z-\n963 slides (0.6 µm step-size) is shown.\n964\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint \n\n.CC-BY 4.0 International licenseperpetuity. It is made available under a \npreprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in \nThe copyright holder for thisthis version posted January 11, 2024. ; https://doi.org/10.1101/2024.01.10.574961doi: bioRxiv preprint","source_license":"CC-BY-4.0","license_restricted":false}