A prion-like protein regulates the 2-dimensional to 3-dimensional growth transition in the mossPhyscomitrium patens

38 The colonization of the land by plants coinci ded with the evolution of 3-dimensional (3D) 39 growth; the acquisition of apical cells with the capacity to rotate the plane of cell division. The 40 moss Physcomitrium patens has recently been developed as a model to dissect the genetic 41 basis of 3D growth, an invariable and unifying feature of all land plants. The cytokinin-42 unresponsive Ppnog1-R mutant incorrectly orients division planes in developing buds and 43 thus fails to make the transition to 3D grow th. To reveal the genetic interactors of the 44 PpNOG1 gene, which encodes a protein with a C-terminal UBA domain, we performed a 45 screen and identified the suppressor of nog1a ( snog1a) mutant. We have mapped the 46 causative mutation to a gene that encodes a prion-like protein related to FLOE2/3 and 47 demonstrated that the mutant phenotypes observed in both a nog1 disruptant mutant 48 (nog1dis) and snog1a can be attributed to changes in cytokinin perception. We present a 49 revised model for 3D growth and suggest that th e 2D-to-3D growth transition is regulated, at 50 least in part, by liquid-liquid phase separation (LLPS). 51 52

53 In the absence of cell movement, plants rely on cell growth processes, combined with 54 asymmetric and precisely orientated cell divisions, to generate new morphologies and 55 diverse cell types with speciali sed functions. The main driving force behind cellular diversity 56 is the formation of apical cells that can divide to self-renew and generate new cell types, and 57 it is the geometry of an apical cell and the way it divides that can greatly influence the 58 pattern of growth and development that follows. Thus, diversification of plant form can 59 largely be attributed to altered division processes in apical cells, which occurred prior to the 60 transition from water to land approximately 470 million years ago (Kenrick & Crane, 1997). 61 Charophycean green algae, the sister lineage to the land plants, can develop apical cells but 62 these only have the capacity for either 1-dim ensional (1D) growth (one cutting face) or 2-63 dimensional (2D) growth (two cutting faces) . As a result, the morphologies represented 64 within the multicellular charophytes are typically filamentous (e.g., Chara braunii) or disc-like 65 (e.g., Coleochaete orbicularis) (Kenrick and Crane, 1997; Wickett et al., 2014; Delwiche and 66 Cooper, 2015; Harrison, 2017). 3-dimensional (3D) growth, the development of apical cells 67 with three or more cutting faces, is a unifying and unvarying feature of all land plants. It was 68 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 3 likely the emergence of 3D growth processes, along with the development of a multicellular 69 sporophyte and the acquisition of vegetative desic cation tolerance, that enabled successful 70 terrestrialization (Delwiche & Cooper, 2015; Harrison, 2017; Moody, 2020). 71 The gametophyte of the moss Physcomitrium patens is well suited to studies of 3D growth. 72 This is because an extensive 2D filamentou s growth phase precedes the transition to 3D 73 growth, and thus the 2D to 3D growth transition can be studied without causing lethality, as 74 the 2D growth phase can be vegetatively propagated in perpetuity (Moody et al., 2018; 75 Moody et al., 2018b; Moody et al., 2021). The gametophyte phase of the P. patens life cycle 76 begins with the germination of a haploid spor e, which gives rise to a 2D branching 77 filamentous network known as the protonema, which extends by tip growth. The protonema 78 consists of two cell types; the first to em erge are the chloroplast-dense chloronemal cells, 79 and subsequently caulonemal cells that form because of an auxin-mediated reprogramming 80 of a chloronemal apical cell into a caulonemal apical cell (Jang and Dolan, 2011; Jaeger and 81 Moody, 2021). Caulonemal cells can undergo a lateral division to produce side branch 82 initials, most of which go on to form secondary protonema (2D fate, approximately 95%) but 83 some acquire 3D fate (gametophore initials) and give rise to gametophores (approximately 84 5%) (Aoyama et al., 2012). The fate of a side branch initial is likely determined by highly 85 localized cues within a caulonemal cell prior to side branch emergence, as a single 86 caulonemal cell can simultaneously divide to give rise to both a filament and a gametophore 87 (Harrison et al., 2009). Elegant work by Aoyama and colleagues has demonstrated that 88 persistent expression of a group of AP2-type transcription factors ( PpAPB1-4) is required to 89 commit a side branch to 3D cell fate. Loss of PpAPB1-4 expression corresponds to the 90 maintenance of 2D cell fate, and consequently, mutants lacking all four genes fail to make 91 the transition from 2D to 3D growth (Aoyama et al., 2012). 92 A filament initial is readily distinguishable from a gametophore initial: (i) the angle of the 93 lateral “transition” division can dictate whether a caulonemal cell gives rise to a filament or a 94 gametophore (Tang et al., 2020); and (ii) gametophore initials swell diffusely and divide in a 95 characteristically oblique manner during the specif ication of 3D growth, at the first, second 96 and third divisions of the developing gametophore. Successive rotating divisions then lead to 97 the establishment of a tetrahedral apical cell at the apex of the shoot. The apical cell 98 continuously self-renews and divides in three planes to form phyllid initials that go on to form 99 the leaf-like phyllids that are organized aroun d the central axes of developing gametophores 100 in a spiral phyllotaxy (Harrison et al., 2009). Mature gametophores bear both archegonia and 101 antheridia, which house eggs and sperm respectively. Following fertilization, a mitotic 102 programme initiates, leading to the formation of a multicellular diploid sporophyte, which 103 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 4 undergoes meiosis to produce haploid spores to restart the life cycle (Cove and Knight, 104 1993). 105 It has long been known that cytokinin induces the formation of gametophore initials but is 106 insufficient to maintain 3D growth, as treatment with high levels of cytokinin produces buds 107 that develop into callus-like tissue, rather than structurally organized gametophores 108 (Brandes and Kende, 1968; Ashton et al., 1979). In a variety of developmental contexts, 109 auxin has been consistently implicated as the si gnal required to ‘break symmetry’ in plants 110 (Petricka et al., 2009; Shao and Dong, 2016). Perhaps unsurprisingly, the formation of 111 gametophore initial cells and the specification and maintenance of 3D growth also relies 112 upon auxin signaling, although the application of high levels of auxin can antagonize 113 cytokinin and reverse the effects of cytokinin treatment (Brandes and Kende, 1968). 114 Nevertheless, auxin levels are notably high during the specification of 3D growth but 115 diminish once a tetrahedral apical cell has been established (Thelander et al., 2018). It is 116 likely that auxin acts to promote cell differ entiation and that cytokinin acts to continually 117 maintain apical cell proliferation (Hata and Kyozuka, 2021). Thus, it appears that a highly 118 regulated balancing act between auxin and cytokini n is required for both the specification 119 and maintenance of 3D growth. 120 In recent years, functional studies have demonstrated that the transition to 3D growth is 121 complex and regulated at many levels; both epigenetically (Mosquna et al., 2009; Okano et 122 al., 2009; Raquid et al., 2023) and transcriptionally (Aoyama et al., 2012), and an ever-123 expanding number of studies have begun to connect complex cell signaling pathways to 124 post-translational regulation (Girod et al., 1999; Perroud et al., 2014; Demko et al., 2014; 125 Johansen et al., 2016; Hoernstein et al., 2016; Schuessele et al., 2016; Whitewoods et al., 126 2018; Moody et al., 2018; Perroud et al., 2020; Moody et al., 2021; Cammarata et al., 2022). 127 Using a forward genetics approach, we previously demonstrated that the NO 128 GAMETOPHORES 1 ( PpNOG1) gene is essential for the transition to 3D growth in P. 129 patens. Notably, the PpNOG1 gene encodes a protein with a prominent C-terminal ubiquitin-130 associated (UBA) domain, which has been shown to be associated with protein degradation 131 processes (Hofman and Bucher, 1996; Su and Lau, 2009; Moody et al., 2018). Mutants 132 lacking a functional copy of PpNOG1 (the ‘ no gametophores 1 – Reference’ mutant; 133 Ppnog1-R) produce significantly fewer gametophore initials than wild type, even in the 134 presence of cytokinin. Those gametophore initial cells that do form cannot correctly orient 135 the characteristically oblique plane of the first cell division. Cell division planes are then 136 misplaced thereafter, leading to the formation of defective gametophores that undergo very 137 early developmental arrest (Moody et al., 2018). Since the PpAPB genes are downregulated 138 in the Ppnog1R mutant, we previously proposed that PpNOG1 may positively regulate 3D 139 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 5 growth by degrading a repressor of PpAPB transcriptional activation, although the identity of 140 the target(s) remains unknown (Aoyama et al., 2012; Moody et al., 2018). To build on our 141 understanding of the role played by PpNOG1 in both the initiation and specification of 3D 142 growth, we generated a Ppnog1 disruption mutant ( Ppnog1dis) and then performed a 143 suppressor screen to identify mutations that alleviated the Ppnog1dis mutant phenotype (i.e., 144 reversion to 3D growth). Here, we describe the screen, along with the detailed 145 characterization of the suppressor of nog1a ( snog1a) mutant and the identification of the 146 causative mutation within a gene encoding a prion-like protein. 147 148

149 150 Disruption of the PpNOG1 locus recapitulates the Ppnog1-R mutant phenotype. 151 Previously, we performed a UV-mediated forward genetic screen that led to the identification 152 of the PpNOG1 gene (Moody et al., 2018). To explore the PpNOG1 genetic interaction 153 network underpinning the 2D to 3D growth transition, we designed a suppressor screen to 154 identify mutations that alleviated the 3D-defective phenotype caused by loss of PpNOG1 155 function (i.e., the reacquisition of 3D grow th). A preliminary suppressor screen of 156 approximately 3,000 UV-mutagenized lines of the original Ppnog1-R mutant, generated in 157 our original forward genetic screen, identified eight mutants that exhibited a complete 158 reversion to 3D growth. In each of these mutants, the phenotype was caused by correction 159 of the previously described mutation; a T> C transition that converted the premature 160 termination codon back to an arginine residue (data not shown). We therefore decided to 161 disrupt the PpNOG1 locus in such a way that would prevent repair simply through the 162

of UV-induced SNPs. To that end, we set out to generate a nog1 knockout 163 mutant in which the entire coding sequence of PpNOG1 had been replaced with a 164 hygromycin resistance cassette. Several attempts were made to obtain a line in which the 165 entire PpNOG1 sequence had been removed. However, as was the case in previous 166 attempts (Moody et al., 2018), we could achieve disruption of the PpNOG1 locus but 167 unfortunately not a full deletion. Nevertheless, a nog1 null mutant was generated in which a 168 portion of the PpNOG1 promoter had been excised following recombination (Fig. S1A). 169 Furthermore, we were consistently unable to detect a PpNOG1 transcript in this line, and 170 thus had confirmed that we had generated a bona fide PpNOG1 disruptant mutant (nog1dis) 171 (Fig. S1B). Consistent with the original Ppnog1-R mutant, the nog1dis mutant completely 172 and consistently failed to make the transition to 3D growth (Fig. 1). Thus, disruption of the 173 PpNOG1 locus recapitulated the original Ppnog1-R mutant phenotype and generated a 174 suitable line for mutagenesis. 175 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 6 176 177 The suppressor of nog1a ( snog1a) mutant can specify 3D growth and responds to 178 cytokinin. 179 A suppressor screen of 2,864 UV-mutagenized lines of the nog1dis mutant yielded two 180 ‘suppressor of nog1 ’ ( snog1) mutants that exhibited a restoration of 3D growth. In one of 181 these mutants, suppressor of nog1a (snog1a), the formation of gametophores was partially 182 restored to approximately 45% of the frequency of wild type (Fig. 2A,B), although these were 183 somewhat stunted and emerged later than those formed in the wild type (Fig. 2C,D; Fig. S2). 184 In contrast to the nog1dis mutant, in which responses to cytokinin are impaired in a similar 185 manner to that of the Ppnog1-R mutant, buds can be induced by cytokinin treatment in the 186 snog1a mutant, although to a lesser extent than in wild type (Fig. 3). Thus, the mutant 187 phenotypes observed in both the nog1dis and snog1a mutants can be attributed to changes 188 in cytokinin perception. 189 To determine whether the cell division or ientation defects had been repaired in the snog1a 190 mutant, we obtained z-stack projections of developing buds stained with propidium iodide. In 191 wild-type P. patens , the first division of the gametophore initial cell was invariably oblique 192 and yielded an apical and a basal cell (Fig. 3A). Two additional oblique divisions of the apical 193 and basal cells then occurred rather synchronous ly. Although the angle of the division plane 194 was consistent in each case, the order in which the cells divided was inconsistent; we 195 generally observed that the apical cell divided befor e the basal cell as often as the basal cell 196 divided before the apical cell (Fig. 3B,C). Successive rotating divisions specified a 197 gametophore apical cell with a characteristic tetrahedral shape, which self-renewed and 198 divided to give rise to the phyllids, which wrap around the central axis of the gametophore in 199 a spiral phyllotaxy (Fig. 3D) (Harrison et al., 2009). In the nog1dis mutant, similarly to the 200 previously described Ppnog1-R mutant, significantly fewer gametophore initial cells formed, 201 and the first division plane of the gametophore init ial cell was not characteristically oblique. 202 In most cases the initial division pattern followed that of a filament initial cell, in which the 203 division plane was positioned roughly parallel to the parental cell from which it was derived 204 (Fig. 3E). Cell plates were then positioned randomly during subsequent divisions, which 205 prevented the specification and maintenance of a tetrahedral apical cell. In some cases, the 206 gametophore initial cells swelled and elongated excessively but could then divide in a 207 somewhat oblique manner at the first and second divisions (Fig. 3F). However, cell plates 208 were invariably misplaced from the onset of the third division (Fig. 3G). Most developing 209 gametophores arrested early in development, but occasionally callus-like buds appeared 210 because of uncontrolled proliferation (Fig. 3H ). Moreover, bifurcation events were 211 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 7 occasionally observed, which were a result of confused cell fate acquisition, and often 212 associated with supernumerary apical cell formation. However, none of these apical cells 213 were successfully maintained long enough to produce a gametophore. In the snog1a mutant, 214 the formation of gametophore initial cells was partially restored, and these largely followed a 215 wild-type pattern of development, with some exceptions. In some cases, we observed that 216 the angle of the first division plane had been corrected to some extent, but not fully, and in 217 these instances a mature gametophore was not formed (Fig. 3I). However, in most cases, 218 the first division plane was characteristically obl ique and thus the orientation of the division 219 plane had been fully corrected (Fig. 3J). This was followed by two further correctly oriented 220 oblique divisions (Fig. 3K), which formed t he prerequisites for the specification of a 221 conspicuous tetrahedral apical cell at the apex of developing gametophores (Fig. 3L). Thus, 222 restoration of gametophore initial cell formation in the snog1a mutant was accompanied by 223 the reversion of the cell division orientation defects observed in the nog1dis mutant. 224 Notably, the phenotype observed in the snog1a mutant only constituted a partial restoration 225 of the nog1dis phenotype. Nevertheless, a tetrahedral apical cell was established and 226 maintained in the snog1a mutant, and the gametophores formed were fully viable. 227 228 The causative mutation of snog1a resides in a gene that encodes a prion-like protein. 229 To identify the causative mutation in the reproductively viable snog1a mutant, we obtained 230 phenotypically segregating populations by performing a cross between the snog1a mutant 231 and the highly fertile non-mutagenized Reute::mCherry strain (Perroud et al., 2020). 232 Because a conventional cross between two di fferent haploid strains was carried out, and at 233 least two genetic loci were mutated (the mutation within the PpNOG1 gene, and the 234 unknown mutation that caused the snog1a phenotype), we expected one quarter of the 235 progeny to exhibit the original nog1dis mutant phenotype and the remainder to exhibit 236 varying capacities for 3D growth (Fig. S3 A). The frequencies that we observed were 237 generally consistent with the mutation of a single genetic locus in the snog1a mutant, 238 although this is not statistically significant (Fig. 5A). One possible explanation for the 239 reduced nog1dis mutant phenotype frequency observed was that disruption of the PpNOG1 240 gene affects haploid spore germination and/or the viability of young sporelings. 241 Nevertheless, the frequencies observed did not support the idea that causative mutations 242 resided in two or more genetic loci. We therefore prepared genomic DNA from 80 individuals 243 that exhibited the nog1dis phenotype and pooled these in equimolar amounts, and this 244 became the ‘wild-type pool’ (all individuals resembled the non-mutagenized parental line, 245 nog1dis). We also prepared genomic DNA from 98 individuals with the capacity for 3D 246 growth, showing a preference for those that most closely resembled the snog1a mutant, and 247 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 8 this became the ‘mutant pool’ (Supplementary Figure S3A). These two pools were 248 sequenced at 44X coverage alongside both parental lines; the nog1dis mutant (generated in 249 the Villersexel wild-type strain) and the Reute::mCherry line. To identify the genomic region 250 containing the snog1a mutation, bulk segregant analysis was performed. Single nucleotide 251 polymorphisms (SNPs) that differed between the two parental lines were identified as 252 markers and the frequency of each SNP variant was mapped across the chromosomes to 253 show the parental origin of each region. For regions not associated with the phenotypes in 254 the two pools, the expected SNP frequency was around 0.5, showing equal contribution from 255 both parents. The expected snog1a mutant allele frequency was 0 in the wild-type pool and 256 0.67 in the mutant pool (Fig. S3B,C). When the allele frequencies for the mutant individuals 257 were plotted across all 27 chromosomes in the P. patens genome assembly, a peak of the 258 expected allele frequency was revealed on chromosome 8, which provided a region of the 259 genome to interrogate for causative mutations (F ig. S3D). Our analysis revealed that two C 260 > T transitions generated two distinct in-frame termination codons (Gln 335Ter and Gln374Ter) 261 in a single gene (Pp3c8_19720) (Fig. 5B,C). We cloned and sequenced the coding 262 sequence of this gene and confirmed that both point mutations were present in the snog1a 263 mutant, but absent from both the nog1dis mutant and the Reute::mCherry line 264 (Supplementary Data S1). In addition, sequenci ng of the corresponding transcript confirmed 265 the presence of the point mutation but did not reveal any splice variants in the snog1a 266 mutant, a phenomenon observed in both previously described Ppnog1-R and Ppnog2-R 267 mutants (Supplementary Data S2) (Moody et al., 2018; Moody et al., 2021). 268 The gene mutated in the snog1a mutant encodes a protein with a C-terminal UBA, and thus 269 the protein domain architecture resembles that of the PpNOG1 protein (Moody et al., 2018). 270 To infer phylogenetic relationships for the putative SUPPRESSOR OF NOG1A protein, we 271 set out to retrieve orthologous sequences from the genomes of representatives of the 272 chlorophytes, charophytes, bryophytes, lycophytes, monilophytes, gymnosperms and 273 angiosperms (Supplementary Data S3). Remarkably, Pp3c8_19720 shares homology with 274 an Arabidopsis prion-like protein (AtFLOE1 ) that can undergo hydration-dependent phase 275 separation (PS) to regulate seed germination, and related proteins (AtFLOE2 and AtFLOE3) 276 that have been shown to undergo PS in a transient expression system (Dorone et al., 2021). 277 As with the analyses conducted by Dorone et al. (2021), our own phylogenetic analyses, 278 revealed the presence of two discrete clades, a FLOE1-like (FLOE1L) and a FLOE2-like 279 (FLOE2L) clade. However, while it was pr eviously stated that the FLOE1L clade was 280 restricted to seed plants, we found FLOE1L representatives in the seed plants in addition to 281 the monilophytes. Conversely, the FLOE2L clade included Pp3c8_19720, as well as three 282 additional P. patens paralogues, and homologues in most l and plants except for hornworts, 283 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 9 where they appear to have been lost. FLOE2L homologues were also identified in some 284 green algal lineages (Fig. S4, Supplementary Da ta S3). Given the significant homology 285 between Pp3c8_19720 and representatives within the FLOE2L clade, we hereafter refer to 286 Pp3c8_19720 as PpFLOE2L-1. 287 The AtFLOE1, AtFLOE2 and AtFLOE3 proteins contain a predicted folded domain (known 288 as the nucleation domain), a coiled-coil domain (CC), a UBA domain at the C-terminus and 289 two disordered regions; one enriched for asparti c acid and serine (DS-rich domain.) and one 290 enriched for glutamine, proline, and serine (QPS-rich domain) (Dorone et al., 2021). 291 Alignment of PpFLOE2L-1 with AtFLOE1, 2 and 3 confirmed that these protein domains 292 were conserved in PpFLOE2L-1 (Fig. 5D, Fig. S5). As a result of the nonsense mutations 293 introduced into PpFLOE2L-1, a truncated protein was formed in snog1a, which was missing 294 most of the QPS-rich domain in addition to the UBA (Fig. 5D; Supplementary Data S2). 295 To confirm that we had correctly identified th e causative mutation, we used the rice actin 296 promoter to drive the expression of a wild-type version of PpFLOE2L-1 in the snog1a 297 mutant. The resulting line exhibited a reversion to the 3D-defective phenotype seen in the 298 nog1dis mutant (Fig. 6A,B and Fig. S6). We also generated two independent lines in which 299 PpFLOE2L-1 was deleted in the nog1dis mutant. Each line exhibited a restoration of 3D 300 growth, and thus the snog1a mutant phenotype had successfully been recapitulated (Fig. 301 6C-E, Fig. S7). Thus, the causative mutation of snog1a resides in a gene that encodes a 302 prion-like protein. 303 304

305 The acquisition of apical cells with the capacity for 3D growth occurred in the last common 306 ancestor of land plants, and enabled the divers e morphologies seen across the planet today 307 (Kenrick and Crane, 1997; Delwiche and Cooper, 2015). We previously showed that the NO 308 GAMETOPHORES 1 ( PpNOG1) gene is required for 3D growth in P. patens , an extant 309 representative of the bryophytes. Notably, in mutants lacking PpNOG1 function (Ppnog1-R), 310 gametophore initial cell formation is significantly reduced, even in the presence of cytokinin. 311 Furthermore, cell division planes are misoriented in emerging gametophores, which 312 subsequently undergo premature developmental arrest. Thus, PpNOG1 promotes the 313 cytokinin-mediated transition from 2D to 3D growth and positively regulates the orientation of 314 cell divisions required to establish a tetrahedral apical cell (Moody et al., 2018). 315 To reveal new insights into the genetic interaction network underpinning the transition from 316 2D to 3D growth, we first recapitulated the Ppnog1-R mutant phenotype in an independent 317 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 10 Ppnog1 disruption mutant ( nog1dis), in which PpNOG1 was not expressed (Fig. 1). The 318 nog1dis mutant was generated using a targeted approach, and thus lacked the background 319 SNPs that were introduced into the Ppnog1-R mutant by UV-based mutagenesis. We then 320 performed a UV-mediated suppressor screen to identify mutations that alleviated the 321 nog1dis mutant phenotype to permit the reversion to 3D growth. The primary focus of this 322 paper is the characterisation of the ‘ suppressor of nog1a’ (snog1a) mutant identified in this 323 screen. 324 Similarly to the original Ppnog1-R mutant, the nog1dis mutant was unable to form 325 gametophores even in the presence of cytokinin (Fig. 1,3). Mutants in which all four PpAPB 326 genes have been disrupted exhibit a similar leve l of cytokinin-unresponsiveness to both 327 Ppnog1-R and nog1dis mutants and fail to make the 3D growth transition (Aoyama et al., 328 2012). Furthermore, we previously observed that PpAPB genes are downregulated when 329 PpNOG1 is absent (Moody et al., 2018). Thus, PpNOG1, along with the PpAPB genes are 330 integral regulators of cytokinin perception during the switch from 2D to 3D growth. Since the 331 snog1a mutant exhibits responsiveness to cytoki nin, this suggests that those cytokinin 332 signaling components, ordinarily repressed in nog1dis , have been reactivated in the snog1a 333 mutant. This demonstrates that the phenotypes observed in both nog1dis and snog1a can 334 be attributed to alterations in cytokinin perception. Thus, this study has begun to shed light 335 on the potential mechanism underlying cytokinin-mediated 3D growth initiation, a topic that 336 has fascinated biologists for several decades (Brandes and Kende, 1968; Ashton et al., 337 1979; Reski and Abel, 1985; Schulz et al., 2000; Schulz et al., 2001; von Schwartzenberg et 338 al., 2007; von Schwartzenberg et al., 2016; Cammarata et al., 2022). 339 Although the formation of gametophore initial cells was not fully restored in the snog1a 340 mutant, those that formed usually followed a wild-type pattern of development to achieve the 341 establishment of a tetrahedral apical cell (Fig . 4). It has previously been shown that the 342 positioning of cell division planes in emerging gametophores is dependent on microtubules 343 (Kosetsu et al., 2017; Kozgunova et al., 2022). Thus, it is likely that the cell division 344 orientation defects observed in the nog1dis mutant are due to aberrant organisation of the 345 microtubule cytoskeleton. Since the correction of division plane orientation observed in the 346 snog1a mutant is accompanied by the restoration of a cytokinin response, we speculate that 347 that microtubule organisation that occurs dur ing the 3D growth transition is dependent on 348 cytokinin. This is consistent with other reports in the literature (Montesinos et al., 2020). 349 There is an increasing volume of literature that describes the regulatory role of cuticle-350 related genes in the 3D growth transition (Renault et al., 2017; Lee et al., 2020; 351 Kreigshauser et al., 2021; Moody et al., 2021). These mutants invariably exhibit 352 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 11 perturbations in the frequencies of gametophore initial cells that form, and those that do form 353 exhibit division orientation defects that arrest gametophore development. Furthermore, the 354 expression of these genes has been shown to be induced by cytokinin (Moody et al., 2021). 355 Although we understand that a cuticle is absent in the protonema but observed in 356 gametophores, the developmental stage at which cuticle biosynthesis initiates remains 357 unclear (Renault et al., 2017; Lee et al., 2020; Kreigshauser et al., 2021). In several of the 358 3D-defective mutants described in th e literature so far, including the Ppnog1-R mutant, the 359 cuticle is invariably absent (Aoyama et al., 2012; Renault et al., 2017; Moody et al., 2018; 360 Kreigshauser et al., 2021; Zhang et al., 2024). Thus , it is possible that the acquisition of the 361 cuticle was a pre-requisite for 3D growth, and that the mechanical constraints imposed by 362 the cuticle alter the division properties of cells formed in the steps toward the establishment 363 of a tetrahedral apical cell. To support this hypothesis, the cuticle is notably absent in the 364 nog1dis mutant but has been partially restored in the snog1a mutant, in which the partial 365 reversion to 3D growth has occurred. 366 Our mapping approach and phylogenetic analyses revealed that the gene mutated in the 367 snog1a mutant (Pp3c8_19720) was a homologue of the previously characterized FLOE 368 genes in Arabidopsis; AtFLOE1, AtFLOE2 and AtFLOE3 (Fig. S4) (Dorone et al., 2021). 369 Dorone et al. have demonstrated that AtFLOE1 undergoes reversible hydration-dependent 370 liquid-liquid phase separation (LLPS) in the embryo, and that this process is dependent on 371 the presence of a QPS-rich disordered domain. LLPS drives the formation of AtFLOE1 372 condensates when a seed is hydrated, but AtFLOE1 remains dispersed in desiccated seeds. 373 Since loss of AtFLOE1 function can promote germi nation during drought or salt stress, it is 374 thought that AtFLOE1 functions to inhibit s eed germination in unfavourable conditions. The 375 authors also demonstrated that AtFLOE2 and AtFL OE3, in addition to representatives from 376 algae and bryophytes, undergo LLPS in a transient expression system (Dorone et al., 2021). 377 AtFLOE1 is a member of the FLOE1-like (FLOE1L) clade, which contains only those 378 representatives from the monilophytes and seed plants. On the other hand, AtFLOE2 and 379 AtFLOE3 reside within the FLOE2-like (FLOE2L) clade, which contains representatives from 380 algal species as well as all land plant lineages. This includes Pp3c8_19720 (PpFLOE2L-1), 381 in addition to three additional homologues in P. patens. We were able to complement the 382 snog1a mutant phenotype with a full-length vers ion of the wild-type coding sequence and 383 recapitulate the phenotype by generating two independent double disruptant mutants (Fig. 384 6). The confirmation of gene identity enabled us to amend the name of Pp3c8_19720 to 385 PpFLOE2L-1. The presence of three additional FLOE2L homologues in the P. patens 386 genome suggests that these genes may function redundantly. Thus, we hypothesise that 387 higher order mutants of these genes will exhibit a progressive restoration of 3D growth, and 388 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 12 indeed cuticle formation. FLOE2L genes were notably absent from hornworts, which 389 suggests that the genetic toolkit underpinning 3D growth processes in hornworts is 390 somewhat distinct. 391 Liquid-liquid phase separation (LLPS) is a phenomenon that has been increasingly linked to 392 important developmental processes in plants (Fang et al., 2019; Huang et al., 2021; Cao et 393 al., 2023). Furthermore, there have been reports that LLPS enables ubiquitin-binding shuttle 394 proteins (i.e., those containing UBAs ) to degrade ubiquitinated substrates (Dao and 395 Castaneda, 2020). Similarly, to the three other FLOE2-like genes identified in P. patens , 396 PpFLOE2L-1 encodes a protein that contains a diso rdered domain enriched in glutamine, 397 proline, and serine (QPS-rich), previously show n to serve as a prerequisite for LLPS; coiled-398 coil (CC) and nucleation domains; a disorder ed domain enriched in aspartic acid and serine 399 (DS-rich); and a UBA. The presence of the UBA suggests that PpFLOE2L-1 plays a role in 400 protein degradation, which is curiously remini scent of the PpNOG1 protein. Due to the 401 presence of two in-frame premature stop codons in the PpFLOE2L-1 transcript, the snog1a 402 mutant lacks most of the QPS-rich disordered domain as well as the UBA (Fig. 5). Thus, we 403 hypothesise that PpFLOE2L-1 undergoes cell-type dependent LLPS to compartmentalize 404 the cellular components required to degrade a ubiquitinated repressor of the 2D to 3D 405 growth transition. In support of this notion, it has been reported that E3 ligases involved in 406 protein degradation processes are able to func tion within condensates. Thus, we propose 407 LLPS as a mechanism by which the induction of 3D growth can be rapidly triggered in 408 response to both intrinsic and extrinsic cues, at the correct stage of development. 409 410 SUMMARY 411 Since the loss of PpFLOE2L-1 function can reverse the 3D-defective phenotype of the 412 nog1dis mutant phenotype, we have demonstrated that PpFLOE2L-1 acts as a negative 413 regulator of 3D growth. Thus, we propose that PpNOG1 acts upstream of, and represses the 414 activity of PpFLOE2L-1, which in turn induc es the degradation of protein(s) that repress 415 PpAPB gene transcription. We speculate that the decis ion to induce 2D versus 3D growth is 416 dependent on the biophysical state of PpFLOE2L-1, and that these changes dynamically 417 alter in a spatial and temporal manner; c ondensed when a side branch acquires 2D fate and 418 dispersed when a side branch acquires 3D fate . We also propose that the transcriptional 419 targets of the PpAPBs trigger the cytokinin response required to initiate the 3D growth 420 transition. In the presence of cytokinin, cuticle-related genes are induced, which we propose 421 subsequently triggers the induction of a CLAVATA-dependent auxin res ponse to diminish 422 the cytokinin response and prevent the inappropriate induction of 3D growth (Fig. 7). 423 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 13 424

425 Physcomitrium patens growth conditions 426 To encourage bud development, P. patens was grown on cellophane-overlaid BCD medium 427 (1 mM MgSO4, 1.84 mM KH2PO4 (pH 6.5), 10 mM KNO 3, 45 µM FeSO 4; 1 mM CaCl2; 0.1% 428 Trace Elements Solution: 116 µM AlK(SO 4)2, 220 µM CuSO 4, 10 mM H 3BO4, 235 µM KBr, 429 660 µM LiCl, 230 µM CoCl 2, 190 µM ZnSO 4, 2 mM MnCl 2, 170 µM KI, 124 µM SnCl 2) 430 containing 0.8% agar. For routine propagation, and to stimulate filamentous growth, P. 431 patens was grown on BCD medium supplemented with 5 mM ammonium tartrate (BCDAT). 432 To enable propagation, tissues were harvested and homogenized in sterile water using an 433 IKA T-25 digital ULTRA-TURRAX®. Tissues were then pipetted onto cellophane-overlaid 434 BCDAT plates in a laminar flow hood. Plates were then placed in a growth cabinet at 24°C 435 with a 16 h:8 h light (300 µmol m -2 s-1): dark cycle. Protoplasts were regenerated on 436 cellophane-overlaid Protoplast Regeneration Medium (PRMB; BCDAT supplemented with 10 437 mM CaCl2, 0.5% glucose and 6% (w/v) D-mannitol) (Cove et al., 2009). 438 439 Generation of the nog1dis mutant 440 A genomic DNA fragment from -827bp upstream of the start codon, up to but excluding the 441 start codon of the PpNOG1 sequence, was PCR-amplified using NOG1.5FKpnI and 442 NOG1.5RXhoI primers (Table 1) and ligated into KpnI/XhoI cut pAHG1 (a kind gift from 443 Yasuko Kamisugi and Andrew Cuming) to create pAHG1- NOG1-5’. A PpNOG1 genomic 444 DNA fragment including the stop codon up to 1420bp downstream of the PpNOG1 stop 445 codon was PCR-amplified using NOG1.3FNotI and NOG1.3RNotI primers and ligated into 446 NotI cut pAHG1- NOG1-5’ to create pNOG1delH. pNOG1delH was linearized using KpnI 447 prior to transformation into protoplasts isolated from the Villersexel wild type strain. Stable 448 transformants were selected using 15 mgmL-1 Hygromycin B (Sigma-Aldrich cat. no. H9773). 449 450 RT-PCR to detect absence of NOG1 in the nog1dis mutant 451 Total RNA was isolated from two-week-old Villersexel wild type and the nog1dis mutant 452 using the RNeasy kit (Qiagen) according to the manufacturer’s instructions. DNase 453 treatment was carried out using TURBO DNase (Ambion) and cDNA synthesis performed 454 using Superscript TM III (ThermoFisher Scientific), as specified by the manufacturers. The 455 tubulin transcript was amplified using PptubF and PptubR, and the PpNOG1 transcript was 456 amplified using 32970008(exon)_GSP.F and 32970008(exon)_GSP.R (Table 1). 457 458 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 14 UV mutagenesis and screening 459 One-week old protonemata from the nog1dis mutant was digested for 1 h in 1% Driselase 460 dissolved in 8% mannitol. The resulting cell suspension was passed through a 40 µm cell 461 strainer and centrifuged for 3 min at 120 xg without braking. Protoplasts were subsequently 462 washed twice in 8% mannitol, with repeated centrifugation steps in between washes. Cells 463 were resuspended in 6 ml 8% mannitol, counted using a haemocytometer, and then plated 464 at a density of 50,000 cells per plate onto cellophane overlaid PRMB. Protoplasts were 465 immediately exposed to a 75,000 mJ dose of UV light using a Stratalinker UV Crosslinker 466 and this was performed with the petri dish lids removed. Following irradiation, the lids were 467 quickly replaced, and the plates were wrapped with micropore tape. Plates were incubated 468 at 24°C in the dark for 24 h, to prevent photoactivatable DNA damage repair, before being 469 transferred to standard growth conditions for a further 2-3 weeks. When visible, regenerating 470 protoplasts were transferred into individual wells of 24 well plates containing BCD medium. 471 After approximately one month of growth in standard conditions, the mutagenized plants 472 were screened for reversion of the nog1dis phenotype (i.e., restoration of gametophore 473 development). 474 475 Imaging 476 To stain cell walls, tissues were submerged in propidium iodide (10 µgmL -1) for around 10 477 min and then mounted on slides in water. Images were acquired using a Leica SP5 scanning 478 confocal microscope with a 40x water immersive lens. A 488 nm laser was used to excite the 479 propidium iodide with 30% laser power and fluorescence was detected at 600-630 nm. 480 481 Plant phenotyping 482 To assay the number of gametophores produced by different P. patens lines, tissues were 483 grown on cellophane-overlaid BCDAT plates for one week and then the tissues were 484 harvested into separate 50 ml tubes containing 10 ml sterile water. The tissues were each 485 homogenized with an IKA disperser for 20 s, and then 1 ml of homogenate was removed to 486 measure optical density (OD600) in a spectrophotometer. The density of the homogenate 487 was normalized by adding the required amount of water to each tube. 50 µl of homogenate 488 was pipetted onto BCD medium in small petri dishes, with 5 repeats for each line. These 489 plates were grown for two months under sta ndard conditions, so that mature gametophores 490 had sufficient time to develop. The number of gametophores formed on each plate were 491 counted. To measure gametophore height, a representative and intact sample from each line 492 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 15 was extracted from the tissue, placed in a petri dish and photographed. The heights were 493 measured in ImageJ, by drawing a line from the apex to the base. To determine the cytokinin 494 responsiveness of each line, tissues were grown on BCD medium for one week and then the 495 cellophanes were cut and transferred to BCD medium containing 1 µM BAP for 3 days (or 496 control medium) before being examined and images captured using a Leica M165C 497 stereomicroscope equipped with a QImaging Micropublishing 5.1 RTV camera. 498 Plant crossing and bulk segregant analysis 499 Both the snog1a mutant and the Reute::mCherry line (Perroud et al., 2019) were grown in 500 magenta pots on BCD medium in standard growth conditions for 2-3 months. Pots were 501 subsequently transferred to sporophyte induction conditions (16 h dark: 8 h light, 16oC). After 502 approximately three weeks, gametophores were examined for the presence of gametangia 503 (archegonia and antheridia). When gametangia were present, the Reute::mCherry line was 504 submerged under water to make a sperm suspension, which was subsequently added to the 505 Ppsnog1a mutant to allow outcrossing. After approximately 4-6 weeks, the resulting 506 sporangia were removed carefully using forceps and examined under a Leica M165C 507 fluorescence stereomicroscope to detect mCherry expression. This demonstrated a 508 successful outcrossing event. Sporangia were sterilized in 70% ethanol for 4 min, washed 3 509 times with sterile water and then incubated at 4 oC in the dark for one week. In a laminar flow 510 hood, the sporangia were transferred to a 15 ml tube containing 10 ml of sterile water and 511 then the sporangia ruptured with a pipette tip. 1 ml of spore suspension was plated onto 512 each of ten cellophane-overlaid BCDAT plates and then grown in standard conditions to 513 permit germination and subsequent growth. Indi vidual sporelings were then transferred to 514 wells of 24-well plates containing BCD medium and grown for a further 4-6 weeks. The 515 progeny were screened for the presence or absence of gametophores, and sorted into two 516 populations; nog1dis-like (wild-type pool, 80 individuals) and wild type or snog1a-like (mutant 517 pool, 98 individuals). Genomic DNA was then extracted from one-week old protonemal 518 tissues prepared from each individual line. 519 520 Isolation of genomic DNA 521 Immediately before use, 0.07% (v/v) 2-merc aptoethanol and 0.1% (w/v) ascorbic acid was 522 added to aliquots of extraction buffer (100 mM Tris-HCl pH 8.0, 1.42 M NaCl, 2 % CTAB, 20 523 mM EDTA, 2% PVP-40). The aliquots of extraction buffer were pre-warmed at 65°C in a 524 water bath. One plate of 1-week-old protonemal tissues, grown on cellophane-overlaid 525 BCDAT plates, was harvested and then blotted on filter paper to remove excess water. 526 Tissues were then placed in a 2 ml microcentrifuge tube and frozen in liquid nitrogen. A 527 miniature pestle was used to grind the tissue into powder, and then 500 µl of pre-warmed 528 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 16 extraction buffer was added to the powder while frozen and further homogenised. An 529 additional 200 µl of extraction buffer was added to the tube, along with 7 µl 10 mgmL -1 530 RNase A, and then incubated at 65°C for 10 min. 600 µl of chloroform-isoamyl alcohol (24:1) 531 was added and the tube, shaken and centrifuged at 13,000 rpm for 10 min. Then the upper 532 aqueous phase of the mixture was transferred to a fresh tube, 0.7 volumes of isopropanol 533 were added, the tube shaken and spun again immediately at 13,000 rpm for 10 min. The 534 pellet was washed in 70% ethanol, allowed to air-dry, and then resuspended in 30 µl 535 nuclease-free water. A NanoDrop™ spectrophotometer was used to assess the quantity and 536 quality of the extracted DNA. 537 538 Preparation of genomic DNA samples for Whole Genome Sequencing 539 In total, four genomic DNA samples were prepared for sequencing; both parental lines 540 (nog1dis and Reute::mCherry) and two pooled samples (a snog1a-like ‘mutant pool’ and a 541 nog1-like ‘wild-type pool’). The pooled samples were prepared by pooling 1 µg genomic 542 DNA extracted from all individuals in that population (80 individuals in the ‘WT pool’ and 98 543 individuals in the ‘mutant pool’). 544 545 Candidate identification through bulk segregant analysis 546 Whole genome was performed using a NovoSeq X Plus sequencing platform (150 bp PE 547 read lengths, 44X coverage) at Novogene. Bioinformatic analysis was performed as 548 described previously (Moody et al., 2018; Moody et al., 2021). The presence of the 549 premature termination codons in Pp3c8_19720 was confirmed by sequencing PCR products, 550 amplified from both wild type and snog1a derived genomic DNA, using the primers 551 Pp3c8_19720_int_F and Pp3c8_19720_int_R (Table 1). 552 553 Physcomitrium patens transformation 554 Before the transformation, large quantities of plasmid DNA were acquired by midiprep using 555 a QIAGEN plasmid midi kit. Approximately 20 µg of the plasmid was linearized by restriction 556 digest overnight, treated with CIAP and precipitated with sodium acetate. Before starting the 557 transformation, all solutions required were made fresh and filter sterilized using [filter]. The 558 transformation was carried out in a biological safety cabinet. 2 g of polyethylene glycol 559 (PEG) 6000 in a flat-bottomed vial, that had been sterilised in the autoclave, was melted in 560 the microwave for approximately 1 min. 5 ml mannitol/Ca(NO 3)2 solution (0.8% mannitol, 0.1 561 M Ca(NO3)2, 10 mM Tris pH 8.0) was added to the molten PEG 6000, shaken and allowed to 562 cool for 2-3 h. 563 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 17 For each P. patens line that was to be transformed, 1-2 plates of tissue were harvested. 0.1 564 g Driselase enzyme was dissolved in 10 ml 8% mannitol for each digest. The tube containing 565 the enzyme solution was wrapped in foil and rocked for approximately 10 min at room 566 temperature. The tube was spun for 3 min in a centrifuge, and the supernatant was filter 567 sterilised. P. patens tissue was placed in the tube with the Driselase solution, wrapped in foil 568 and rocked very gently until the tissue appears well digested (at least 40 min). The digested 569 tissue was put through a 70 µm cell strainer to separate protoplasts from remaining debris. 570 The protoplasts were spun at no more than 120 xg, the supernatant was removed and the 571 protoplasts were resuspended in 6 ml mannit ol. This wash was repeated twice more. The 572 cell density of the protoplasts was determined using a haemocytometer. The protoplasts 573 were spun down again and resuspended in MMM (0.5 M m annitol, 0.15 M MgCl2, 0.1% MES 574 pH5.6), to achieve a cell density of 1.5x10 6 cells mL -1. The next steps were carried out 575 without delay as the protoplasts cannot tolerate the MMM solution for very l ong. At least 10 576 µg of the linearised construct was pipetted into the bottom of a round-bottomed Falcon tube. 577 To this was added 300 µl of protoplast suspension. 300 µl of the PEG solution was added in 578 drops and the tube was swirled gently after the addition of each drop. The tube was heat-579 shocked at 45 °C in a water bath for 5 min and then incubated at room temperature for a 580 further 5 min. Next, 300 µl 8% mannitol was added to the tube, 5 times at 3 min intervals and 581 the mixture was gently swirled after each additi on. Then, 1 ml 8% mannitol was added to the 582 tube 5 times at 3 min intervals, gently tilting the tube after each addition to mix it. The tube 583 was centrifuged at 120 xg, supernatant removed, and the cells were resuspended in 3 ml 8% 584 mannitol. This suspension was gently pipetted onto three PRMB plates overlaid with 585 cellophanes (1 ml for each plate). These plates were sealed with micropore tape and 586 wrapped in foil for 24 h. Subsequently, they were unwrapped and placed in normal growing 587 conditions for 5-7 days. Then the cellophanes, with the regenerating protoplasts, were 588 transferred to selective BCDAT plates containing the appropriate selective agent. After 589 another week on selective plates, the cellophanes are transferred to BCDAT plates with no 590 selective agent to allow recovery. After another 1-2 weeks, the surviving P. patens colonies 591 were individually transferred to selective BC D plates, to select for stably transformed 592 colonies. Following another 1 – 2 weeks on selection, colonies that survived both rounds of 593 selection were transferred to non-selective BCD plates and allowed to grow and generate 594 tissue for further study. 595 596 Generation of snog1a complementation lines 597 RNA was extracted form two-week-old wild-type tissue using an RNeasy kit (Qiagen) and 598 treated with Turbo DNase (Ambion), according to the manufacturer’s specifications. cDNA 599 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 18 was then synthesised using Superscript TM III Reverse Transcriptase (ThermoFisher 600 Scientific). Subsequently the Pp3c8_19720 transcript was PCR-amplified (excluding the stop 601 codon) using Pp3c8_19720.FSalI and Pp3c8_19720.R_NOSTOP_HindIII and ligated into 602 pZAG1 to create Act1p::Pp3c8_19720-GFP. This construct was linearised with SacII for 603 subsequent transformation into the snog1a mutant. Stable transformants were selected 604 using 100 µgml -1 Zeocin (Invitrogen; R25001). Genotyping was performed using the primers 605 putativeSNOG1A_genomic2_F and putativeSNOG1A_genomic2_R (Table 1, Fig. S6). 606 607 Generation of a snog1a deletion mutant 608 A deletion construct was designed. The construct consisted of the PpFLOE2L-1 5’ flanking 609 sequence, with a small section of the PpFLOE2L-1 CDS and a PpFLOE2L-1 3’ flanking 610 sequences. The 5’ and 3’ sequences were either side of a G418 resistance cassette. The 611 construct was synthesized by TWIST Biosci ence. The product was verified by Sanger 612 sequencing. Prior to transformation in to protoplasts isolated from the nog1dis mutant, the 613 plasmid was linearized with PvuI. Stable transformants were selected using 40 µgml -1 G418. 614 5’ integration of the deletion constr uct was confirmed using the primers 615 snog1a_del_genotyping_F and snog1a_del_genotyping_R (Table 1, Fig. S7). 616 617 Phylogenetics 618 For each of the species included in the analys is, proteome sequences (primary transcript 619 only) were obtained. The data used were: Physcomitrium patens v3.3 (Lang et al., 2018), 620 Marchantia polymorpha v3.1 (Bowman et al., 2017), Selaginella moellendorffii v1.0 (Banks 621 et al., 2011), Oryza sativa v7.0 (Ouyang et al., 2007), Zea mays PH207 v1.1 (Hirsch et al., 622 2016), Sorghum bicolor v3.1.1 (McCormick et al., 2018), Brachypodium distachyon v3.1 623 (Vogel et al., 2010), Arabidopsis thaliana Araport11 (Cheng et al., 2017), Solanum 624 lycopersicum ITAG2.4 (Sato et al., 2012), Medicago truncatula Mt4.0v1 (Tang et al., 2014), 625 Populus trichocarpa v3.1 (Tuskan et al., 2006), Micromonas pusilla CCMP1545 v3.0 626 (Worden et al., 2009) and Ostreococcus lucimarinus v2.0 (Palenik et al., 2007), Chara 627 braunii S276v1.0 (Nishiyama et al., 2018), Chlamydomonas reinhardtii v5.5 (Merchant et al., 628 2007), Amborella trichopoda v1.0 (Albert et al., 2013), Botryococcus braunii v2.1 (Browne et 629 al., 2017), Anthoceros agrestis Oxford (Li et al ., 2020), Azolla filiculoides v1.1 (Li, et al ., 630 2018), Brassica rapa FPsc v1.3 (DOE-JGI, http://phytozome.jgi.doe.gov/ ), Ceratodon 631 purpureus R40 v1.1 (Carey et al., 2021), Ceratopteris richardii v2.1 (Marchant et al., 2022), 632 Chlorokybus atmophyticus CCAC 0220 v1.1 (Wang et al ., 2020), Klebsormidium nitens 633 NIES-2285 v1.1 (Hori et al ., 2014), Mesostigma viride NIES-296 (Liang et al ., 2020), 634 Spirogloea muscicola CCAC 0214 (Cheng et al., 2019), Mesotaenium endlicherianum SAG 635 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 19 12.97 (Cheng et al., 2019), Coccomyxa subellipsoidea C-169 v2.0 (Blanc et al., 2012), 636 Dunaliella salina v1.0 (Polle, et al ., 2017), Volvox carteri v2.1 (Prochnik eta l ., 2010), 637 Porphyra umbilicalis v1.5 (Brawley et al., 2017), Sphagnum fallax v1.1 (Healey et al., 2023), 638 Diphasiastrum complanatum v3.1 (DOE-JGI, http://phytozome-next.jgi.doe.gov/), Gingko 639 biloba v2021 (Liu et al ., 2021), Glycine max Wm82 ISU-01 v2.1 (DOE-JGI, 640 http://phytozome.jgi.doe.gov), Gossypium raimondii v2.1 (Paterson et al ., 2012), Musa 641 acuminata v1 (D’Hont et al ., 2012), Panicum hallii v3.2 (Lovell et al ., 2018), Salvinia 642 cucullata v1.2 (Li et al., 2018), Spirodela polyrhiza v2 (Wang et al., 2014), Thuja plicata v3.1 643 (Shalev et al., 2022), Vitis vinifera v2.1 (Jaillon et al., 2007), Saccharomyces cerevisiae R64-644 1-1 (Liachko et al., 2013), Drosophila melanogaster BDGP6.32 (Adams et al., 2000) and 645 Homo sapiens GRCh38 (Lander et al., 2001). 646 With this set of proteomes, OrthoFinder was used to identify orthogroups (Emms & Kelly, 647 2019). The orthogroup containing Pp3c8_19720 was selected and the protein sequences 648 were aligned using MAFFT (L-IN-SI method) and a gene tree constructed using IQ-TREE, 649 with automatic model selection (ModelFinder) and ultrafast bootstrapping (UFBoot) with 650 1000 replicates (Hoang et al., 2018; Kalyaanamoorthy et al., 2017; Katoh & Standley, 2013; 651 Nguyen et al., 2015). Tree was rooted and edited in Interactive Tree of Life (iTOL) (Letunic & 652 Bork, 2016). 653 654

655 We are grateful to Pierre-François Perroud for proving the Renute::mCherry line, Yuji 656 Hiwatahi for providing the pZAG1 plasmid; Andrew Cuming and Yasuko Kamisugi for 657 providing the pAHG1 plasmid and John Baker for photography. 658 659 FUNDING 660 The work was funded by a Royal Society University Research Fellowship awarded to L.A.M. 661 (URF\R1\191310), a Royal Society University Research Fellowship (URF\R1\201033) and a 662 Wellcome Trust grant (226598/Z/22/Z) awarded to S.K., a BBSRC PhD studentship 663 (BB/M011224/1) awarded to Z.W. and a BBSRC PhD studentship (BB/T008784/1) awarded 664 to G.C. 665 666 AUTHOR CONTRIBUTIONS 667 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 20 Z.W. conducted the experiments, with assistance from G.C., E.D. and L.A.M.; L.A.M. 668 conceived and designed the study; S.K. carried out the bioinformatics; Z.W. constructed the 669 phylogenetic tree; and L.A.M. and Z.W. wrote and edited the manuscript. 670 671 672 673 674 675

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Scale bars, 1 cm. 1085 Figure 2. The snog1a mutant exhibits a partial restoration of 3D growth. A) 1086 Representative images of 6-week-old Villersexel wild type (WT), nog1dis and snog1a plants 1087 showing the presence (WT and snog1a) and absence (nog1dis) of gametophores. B) Mean 1088 number of gametophores per culture (n=5) ± SEM (t test ***p < 0.05). C) Representative 1089 images of a gametophore from wild ty pe (top) and stunted gametophore from the snog1a 1090 mutant (bottom). D) Mean height of gametophores from wild type (n=100) and snog1a 1091 (n=80) ± SEM (t test ***p < 0.05). Scale bars, 1 cm (A and C). 1092 Figure 3. The snog1a mutant is cytokinin responsive. Representative images of wild type 1093 (WT), nog1dis and snog1a plants cultured in the presence or absence of the cytokinin 1094 analogue 6-benzylaminopurine (BAP). 1095 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 34 Figure 4. The snog1a mutant can establish and maintain a tetrahedral apical cell. 1096 Propidium-iodide-stained buds of wild type at the 2-cell (A), 3-cell (B) and 4-cell (C) and late 1097 stage (D); the nog1dis mutant at the 2-cell (E,F), 4-cell (G) and late stage (H); and the 1098 snog1a mutant at the 2-cell (I,J), 4-cell (K) and late stage (L). Red arrows denote the most 1099 recent division in each developing bud, and blue asterisks highlight misoriented division 1100 planes in nog1dis or snog1a mutants. 1101 Figure 5. Identification of the causative mutation in the snog1a mutant. A) Phenotypic 1102 analysis of spore progeny derived from a cross between snog1a and the Reute::mCherry 1103 wild-type strain. B) Gene candidates identified following interrogation of the genomic locus 1104 on chromosome 8. C) Gene structure diagram of Pp3c8_19720 highlighting the presence of 1105 two termination codons in the third exon (exons – blocks, introns – horizontal lines). D) The 1106 wild-type Pp3c8_19720 protein (top) and the truncated Pp3c8_19720 protein in snog1a 1107 (bottom). 1108 Figure 6. Confirmation of the causative mutation in the snog1a mutant. (A,B) 1109 Representative images of 6-week-old snog1a (A) and snog1a complemented with 1110 pAct::PpFLOE2L-1 (B). (C-E) Representative images of 6-week-old nog1dis (C) and 1111 nog1dis/floe2l-1_4 (D) and nog1dis/floe2l-1_6 (E) double disruptants. Scale bars, 1 cm. 1112 Figure 7. Speculative model for 3D growth regulation in P. patens . Top panel shows 1113 possible mechanism for cell-type specific LLPS of PpFLOE2L-1. Bottom panel shows 1114 possible relationship between PpFLOE2L-1 and other known regulators of 3D growth. 1115 PpNOG1 is proposed to act upstream of, and negatively regulate PpFLOE2-1, which in turn 1116 negatively regulates the expression of the PpAPB genes. Activation of the cytokinin signaling 1117 triggers the expression of cuticle-related genes, which act upstream of CLAVATA signaling 1118 components, which trigger an auxin-mediated repression of ectopic bud formation. 1119 Supplementary Figure 1. Generation of the nog1dis line. A) Schematic of the construct 1120 designed to knockout the endogenous PpNOG1 gene, and schematic of the modified 1121 PpNOG1 locus in nog1dis following recombination. B) RT-PCR reveals the presence of the 1122 PpNOG1 transcript in wild type but not in the nog1dis mutant (tubulin – control). 1123 Supplementary Figure 2. Gametophores formed in the snog1a mutant are stunted 1124 relative to wild type. Representative images of 2-month-old gametophores from wild type 1125 and snog1a plants. Scale bars, 0.5 cm. 1126 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 35 Supplementary Figure 3. Bulk segregant analysis and the identification of the 1127 causative mutation in the snog1a mutant. A) An outcrossing event between snog1a and 1128 the Reute::mCherry line yields a diploid sporophyte that undergoes meiosis to produce 1129 phenotypically segregating progeny (phenotypic outcomes highlighted). (B,C) Expected 1130 snog1a mutant, SNOG1A WT, nog1 mutant and NOG1 WT allele frequencies in the mutant 1131 (B) and wild-type (C) pools respectively. D) Allele frequency plot for segregants on 1132 chromosome 8 of the P. patens genome assembly. 1133 Supplementary Figure 4. Phylogenetic analysis of FLOE-related homologues in the 1134 green lineage. Bootstrap values have been indicated on each branch. Both FLOE1L and 1135 FLOE2L clades have also been indicated. 1136 Supplementary Figure 5. Alignment of AtFLOE1, AtFLOE2, AtFLOE3 and PpFLOE2L-1. 1137 Conserved domains have been highlighted as indicated. 1138 Supplementary Figure 6. Generation of the snog1a complementation line. A) 1139 Schematic of the construct used to complement the snog1a mutant phenotype, and the 1140 resulting targeted locus. B) Genotyping of the complementation line using 1141 putativeSNOG1A_genomic2_F and putativeSNOG1A_genomic2_R primers denoted by 1142 purple and green arrows in (A) respectively . The construct is only detected in the 1143 complemented line and not in wild type (tubulin – control). 1144 Supplementary Figure 7. (A) Schematic of the construct used to disrupt the PpFLOE2L-1 1145 locus in the nog1dis mutant, and the resulting targeted locus. B) Genotyping of the 1146 complementation line using snog1a_del_genotyping_F and snog1a_del_genotyping_R 1147 primers denoted by blue and red arrows in (A) respectively. 1148 1149 Table 1. List of Primers 1150 Generation of Vxnog1 NOG1.5FKpnI AAAGGTACCCCATCCATGCACACAACCAA NOG1.5RXhoI AAACTCGAGCCTCCGCTCCAAACTCCCAC NOG1.3FNotI AAGCGGCCGCTAATCTGTGTATGAGTTCAG NOG1.3RnotI AAGCGGCCGCGAGTTATCTAGTTTTGTGGA .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint 36 Primers for RT-PCR PptubF TGTGCTGTTGGACAATGAG PptubR ACATCAGATCGAACTTGTG 32970008(exon)_GSP.F GTTGTAGGTTTGGAGTGGCG 32970008(exon)_GSP.R GCAAGTTGAAAAGCCCACCT snog1a mutation verification Pp3c8_19720_int_F CAAGGCCTACCGTCTCATCC Pp3c8_19720_int_R GTGGAGGAGGGACCTCTTGA Generation of snog1a complementation lines Pp3c8_19720.FSalI aaagtcgacATGGATCATGTGGGATCC Pp3c8_19720.R_NOSTOP_HindIII aaaaagcttCCGGCCATACCAGC Verification of snog1a complementation lines putativeSNOG1A_genomic2_F CAGCTGACAACTTTGGTGCA putativeSNOG1A_genomic2_R CATCTGCTGCTGAGGAAGTG Verification of PpFLOE2L-1 disruption lines snog1a_del_genotyping_F GTCCACCAAGACCACGAAAC snog1a_del_genotyping_R CATCAGAGCAGCCGATTGTC 1151 1152 .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint .CC-BY-NC-ND 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted April 10, 2024. ; https://doi.org/10.1101/2024.04.08.588603doi: bioRxiv preprint