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# Clonal-aggregative multicellularity entrained by salinity in one of the closest relatives of animals

Núria Ros-Rocher<fnref n="1" />*, Josean Reyes-Rivera<fnref n="2" />*, Uzuki Horo<fnref n="1" />, Chantal Combredet<fnref n="1" />,

Yeganeh Foroughijabbari<fnref n="1" />, Ben T. Larson<fnref n="2" />,<fnref n="3" />, Maxwell C. Coyle<fnref n="2" />, Erik A.T.

Houtepen<fnref n="4" />, Mark J. A. Vermeij<fnref n="4" />,<fnref n="5" />, Jacob L. Steenwyk<fnref n="2" />, Thibaut Brunet<fnref n="1" />+

<footnote n="1">¹Institut Pasteur, Université Paris-Cité, CNRS UMR3691, Evolutionary Cell Biology and Evolution of Morphogenesis Unit, 25-28 Rue du Dr. Roux, 75015 Paris, France</footnote>

<footnote n="2">²Howard Hughes Medical Institute and the Department of Molecular and Cell Biology, University of California, Berkeley, Berkeley, CA, USA</footnote>

<footnote n="3">³Present address: Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA, USA</footnote>

<footnote n="4">⁴Caribbean Research and Management of Biodiversity foundation (CARMABI), Piscaderabaai z/n Willemstad, Curaçao</footnote>

<footnote n="5">⁵Department of Freshwater and Marine Ecology, Institute for Biodiversity and Ecosystem Dynamics, University of Amsterdam, Amsterdam, 1012 WP, The Netherlands</footnote>

<footnote>*Contributed equally</footnote>

<footnote>*Lead contact</footnote>

<footnote>Correspondence: N. R.-R.: nuria.ros@pasteur.fr J. R.-R.: josean.reyes@berkeley.edu T.B.: thibaut.brunet@pasteur.fr</footnote>
Multicellularity evolved multiple times independently during eukaryotic diversification¹⁻⁴. Two distinct mechanisms underpin multicellularity⁵: clonal development (serial cell division of a single precursor cell) and aggregation (in which independent cells assemble into a multicellular entity). Clonal and aggregative development are traditionally considered mutually exclusive¹,⁶⁻⁹, and evolutionary hypotheses have addressed why multicellular development might diverge toward one or the other extreme³,⁴. Both animals and their sister group, the choanoflagellates, are currently thought to only develop clonally¹⁰,¹¹, apparently supporting an exclusively clonal pre-history for animal multicellularity⁴,¹². Here, we show that the choanoflagellate *Choanoeca flexa*, one of the closest living relatives of animals¹³, develops into motile and contractile monolayers of cells (or “sheets”) through an unexpectedly mixed, plastic mechanism: *C. flexa* sheets can form by purely clonal processes, purely aggregative processes, or a combination of both. We characterize the life history of *C. flexa* in its natural environment, ephemeral splash pools on the island of Curaçao, and show that multicellular development is controlled by salinity during natural cycles of splash pool evaporation and refilling. Different splash pools house genetically distinct strains of *C. flexa* between which aggregation is constrained by kin recognition, a hallmark of aggregative multicellularity¹⁴⁻¹⁷. We propose that clonal-aggregative development allows fast and reversible transitions between unicellular and multicellular lifestyles in this rapidly fluctuating environment. Our findings challenge former generalizations about the choanoflagellate-animal lineage and expand the option space for the development and evolution of multicellularity.

Multicellularity has evolved independently over 45 times across eukaryotes¹⁸, with multiple independent origins of both clonal and aggregative multicellularity⁷,¹⁹. Efforts to reconstruct the origin of animal multicellularity have benefited from the study of their closest living relatives: the choanoflagellates (Figure 1A)²⁰⁻²³. Choanoflagellates are bacterivorous aquatic microeukaryotes bearing an apical flagellum surrounded by a collar of interconnected actin-filled microvilli (Figure 1B-C)²⁴. Moreover, many choanoflagellate species display facultative multicellular development²⁴⁻²⁸. As in animals, multicellular development is clonal in all choanoflagellate species investigated thus far²⁵,²⁹⁻³¹, which contrasts with the occurrence of aggregative multicellularity in more distantly related lineages, such as filastereans³²⁻³⁵ and dictyostelid amoebae³⁶,³⁷. These observations, together with the clonal nature of animal embryogenesis, have inspired the hypothesis that animals evolved from organisms with a
simple form of clonal multicellularity, akin to that observed in certain modern choanoflagellates and in other close relatives of animals such as ichthyosporeans³⁸⁻⁴⁰.

Here, we describe an unusual type of multicellular development in the colonial choanoflagellate *Choanoeca flexa*⁴¹, that challenges previous generalities about choanoflagellates. We report that *C. flexa* colonies develop by a combination of clonal and aggregative development, which we refer to as ‘clonal-aggregative multicellularity’. We propose that this unusual mode of development represents an adaptation to the dynamic natural environment of *C. flexa*: splash pools that undergo extreme salinity fluctuations during natural cycles of evaporation and refilling.

### C. flexa sheets can develop clonally

*C. flexa* was discovered in 2018 in the form of concave monolayers of polarized cells (or “sheet colonies”; **Figure 1A-D**) held together through direct collar-collar adhesions⁴¹,⁴². *C. flexa* sheets are capable of inverting their curvature under the control of environmental cues, thus switching from a feeding state (relaxed form, flagella-in; **Figure 1B**) to a swimming state (contracted form, flagella-out; **Figure 1D**)⁴¹,⁴³. In an earlier study, we established stable cultures of *C. flexa* sheets from a single isolated cell, indicating that sheets can arise from individual cells⁴¹. Nevertheless, the mechanisms of *C. flexa* multicellular development remained unexplored.

To understand *C. flexa* colony development, we first isolated and monitored single cells from mechanically disassembled sheets by time-lapse microscopy. In this context, we observed clonal development of sheets by serial cell division (**Figure 1E-F; movie S1**): cells divided asynchronously every ~8-10 hours and sister cells remained attached to each other by intermicrovillar contacts. This resulted in a monolayer of polarized cells with the signature curved morphology of *C. flexa* sheets (**Figure 1E-F; movie S1**). These observations show that small *C. flexa* sheets (up to 7 cells) can develop clonally from a single cell. To test whether clonal development can contribute to the further growth of *C. flexa* colonies, we monitored small and medium-sized colonies (from 6 to 46 cells) by time-lapse microscopy and regularly observed clonal expansion by cell division both at the core and periphery of the sheets (**Figures 1G and S1; movies S2-S3**). However, unexpectedly, we also captured instances of free-swimming single cells and doublets meeting colonies, attaching to their periphery, re-orienting to align their apico-basal axis with neighboring cells, and seemingly integrating into
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the sheet (Figures 1G and S1; movies S2-S3). Taken together, these results show that C. flexa colonies can form clonally but might also be able to expand by aggregation. This motivated us to test whether C. flexa could also acquire multicellularity purely by aggregation (Figure 1H).

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C. flexa sheets can develop by aggregation

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To test whether C. flexa colonies can form by aggregation, we mechanically disassembled colonies into free-swimming single cells and performed live imaging of the dissociated cells (Figure 2A-B; movie S4). We found that C. flexa single cells aggregated within minutes into irregular masses of cells that continued incorporating additional cells and underwent repeated fusion into larger groups (i.e. agglomeration⁴⁴; Figure 2B; movie S4). To determine the fate of these aggregates, we labeled two populations of dissociated single cells with different fluorophores. After mixing both populations, cells of both colors aggregated within minutes (Figure S2A; movie S5). After 24 hours, we observed chimeric colonies displaying the polarized cup-shaped morphology typical of canonical C. flexa sheets and comprising cells of both colors (Figures 2C and S2B-C; movies S6-S8), indicating that they had developed by aggregation. To better understand the dynamics of aggregative development, we fixed aggregates at different stages and imaged them by Airyscan confocal microscopy. We found that early aggregates (up to 6 hours post-dissociation (hpd)) displayed a variable and irregular morphology with cells frequently showing unaligned apico-basal polarity, likely resulting from collision and attachment of cells in diverse random orientations (Figures 2D and S3; movies S9-S12). Colonies observed 24 hpd, on the other hand, were regular polarized monolayers of cells (Figures 2D and S3; movie S13), suggesting that early irregular aggregates had matured into canonical sheets. At that stage, colonies comprised about 50 cells on average and as many as 120 cells (Figure 2E). This size further supported the idea that these sheets could not have formed exclusively by clonal development within 24 hours, given the cell cycle duration in C. flexa (at least 8 hours (Figure 1F), which could result in sheets comprising at most 16 cells assuming maximal and synchronized proliferation). Indeed, treatment with the cell cycle inhibitor aphidicolin abolished cell proliferation (Figure S4A) but did not affect colony formation from dissociated cells (Figure 2F and S4B), confirming that aggregation was sufficient for C. flexa multicellular development.

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Our observation of aggregative development in *C. flexa* contrasts with earlier observations in other species of multicellular choanoflagellates, which have so far been found to develop strictly clonally<sup>24-26</sup>. Given that other protists often deploy aggregative development as an "emergency response" to fluctuating environments<sup>3,45</sup>, while clonal development is thought to be more frequent in environments sufficiently stable to sustain cell proliferation, we were curious about the natural conditions of *C. flexa* mixed clonal-aggregative development. Thus, we set out to investigate the life history of *C. flexa* in its natural environment.

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137 Occurrence of multicellular *C. flexa* in the field is limited by salinity

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139 *C. flexa* was originally discovered in its multicellular form on the tropical island of 140 Curaçao<sup>41</sup> (**Figure 3A**). Unlike other multicellular choanoflagellate species<sup>21,46</sup>, *C. flexa* has 141 been repeatedly re-isolated in the field since its discovery in 2018 and can thus be studied in 142 its native biotope. *C. flexa* sheets are found on the windward northern part of the island in 143 splash pools (**Figures 3A-B and S5A-B**) that undergo natural cycles of desiccation and 144 refilling<sup>47,48</sup>. Water-filled splash pools experience gradual evaporation, leading to increasing 145 salinity and, occasionally, complete desiccation (**Figure 3C**). Splash pools refill from crashing 146 waves, splash, or rain, restoring lower salinity levels (**Figures 3C and S5B; movie S14**). As 147 a consequence, splash pools are ephemeral habitats in which organisms experience extreme 148 and recurrent hypersaline and hyperosmotic stress<sup>48-51</sup>. We set out to investigate how this 149 environment might influence the life history and multicellular development of *C. flexa*. 150

150 We focused our studies on Shete Boka National Park, where *C. flexa* is reliably found 151 (**Figure 3A**). We surveyed the distribution of *C. flexa* in 150 splash pools and measured splash 152 pool salinity in two different field expeditions: Exped-A and Exped-B (**Figure 3D**). During 153 Exped-A, we sampled 79 splash pools (numbered Sp1 to Sp79) along about 2 kilometres of 154 coastline (**Figure 3D**; see methods). While seawater collected from the neighbouring inlets 155 (or *bokas*) had a stable salinity of ~40 parts per thousand (ppt), splash pool salinity ranged 156 from below average seawater salinity (15 ppt) to saturation (≥ 280 ppt) (**Figures 3E and S5D**). 157

157 In 10 out of 79 splash pools, we found choanoflagellate sheets that exhibited the 158 stereotypical *C. flexa* cup-shaped morphology and inversion behaviour in response to light- 159 to-dark transitions<sup>41</sup> (**Figures 3F and S6A-C; movie S15 and Table S1**). 160 161 sequencing of manually isolated sheets confirmed their identification as *C. flexa* (**Figure S6D; 162 supplementary files S1-S4**). Interestingly, although the salinity of surveyed splash pools 163 ranged from 15 to 280 ppt, *C. flexa* sheets were only found in splash pools with <94 ppt salinity

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(Figures 3E and S5D; Table S1). To independently test whether the presence of multicellular C. flexa was constrained by an upper salinity limit, we exhaustively sampled all splash pools (n=71, numbered M1 to M71; Figure 3D' and S5C) within a 4-meter by 10-meter quadrant in a second expedition (Exped-B). We found sheets in 14 out of 71 splash pools, with a 128 ppt salinity upper bound for sheet occurrence (Figures 3E and S5D). Across both expeditions, C. flexa sheets were found in splash pools with an average salinity of 61.5±23.7 ppt, 1.56-fold that of natural seawater (Figures 3E and S4C) and significantly lower than splash pools that did not contain sheets (155.1±107.2 ppt; p=3.1e-04 by the Mann-Whitney U test). We never observed sheets in splash pools with a salinity above 128 ppt (3.2-fold seawater; Figures 3E and S5D; Table S1).

## Multicellular development is entrained by evaporation-refilling cycles in nature

We then set out to study how the natural evaporation-refilling cycle of splash pools impacts the presence/absence of sheets. We monitored ten splash pools in which C. flexa sheets were found on Day 1 and five more randomly selected splash pools daily over eight days (Figures 3G-H and S7). We measured salinity and maximum depth and screened samples for sheets once per day (Figures 3G-H and S7). We observed a gradual decrease in depth and increase in salinity in all 15 splash pools (presumably due to evaporation; Figure S7A-B), six events of complete desiccation (Figure S7C-D), and four events of refilling (Figure S7E-F; see three examples in Figure 3H). In all cases, C. flexa sheets were no longer observed after salinity crossed a ~120 ppt threshold during gradual evaporation (Figures 3H and S7), consistent with results from Exped-A and Exped-B (Figures 3E-H and S7). Moreover, evaporation in the laboratory of a natural splash pool sample containing sheets led to sheet disappearance (Figure S8). These observations confirmed that the multicellular form of C. flexa does not tolerate high salinity.

Interestingly, in two dry splash pools that underwent refilling during our study (restoring salinity down to ~50 ppt), sheets were observed 48 hours after refilling (Sp69 and Sp70; Figures 3H and S7E-F). The newly observed sheets may have arrived from the ocean or from neighbouring splash pools. Alternatively, C. flexa might have persisted in the soil of desiccated splash pools in a cryptic resistant form, such as unicellular cysts (described in other choanoflagellates⁵²⁻⁵⁷). Under this hypothesis, colonies would disassemble and differentiate into resistant single cells under high salinity, and those cells would then develop back into sheets once salinity decreased. To explore this possibility in situ, we collected soil
samples from seven desiccated splash pools in which *C. flexa* had been previously observed, rehydrated them in the lab (down to ~65 ppt salinity) and monitored the rehydrated samples for several days (**Figure 3I-K; movie S16 and Table S1**). Sheets were recovered from various soil samples collected from two splash pools after two to seven days of desiccation in the field (**Figure 3I-K and Table S1**). Sheets were consistently observed 48-72 hours after rehydration in the laboratory, suggesting that a resistant form of *C. flexa* can survive complete desiccation (**Figure 3I-K and Table S1**). Interestingly, earlier metagenomic analyses of diverse environments had suggested that choanoflagellates might often be present in soil⁵⁸,⁵⁹⁶.

Given that salinity fluctuations in splash pools seemed to dramatically impact *C. flexa* multicellularity in nature, we next set out to investigate the phenotypic response of *C. flexa* to comparable evaporation-refilling cycles in a laboratory context.

**Multicellular development is entrained by evaporation-refilling cycles in the laboratory**

We designed an experimental setup mimicking the natural evaporation-refilling cycles of Shete Boka splash pools (**Figures 4A-C and S9A**). We subjected *C. flexa* cultures to evaporation in an incubator (**Figure 4A**), approximating the temperature (30°C) and evaporation rate of natural splash pools (see methods; **Figure 4B-C and S9A**). Starting from the standard salinity of artificial seawater (ASW; 35 ppt, hereafter referred to as 1X salinity), this setup resulted in complete desiccation after four days of evaporation. Under these conditions, *C. flexa* sheets gradually dissociated into non-motile single cells (**Figures 4D-E and S9B; movie S17*), with more than 50% solitary cells when salinity crossed the natural limit of sheet occurrence (128 ppt; **Figures 3E and 4AD**. Almost all sheets had dissociated into single cells when salinity reached saturation (**Figures 4D-E and S9B; movie S17**). By contrast, control cultures without evaporation remained multicellular throughout the experiment (**Figure S9C**). Finally, to directly test whether loss of multicellularity was due to increased salinity, we added seawater salt to *C. flexa* cultures (without evaporation) and similarly observed dissociation of multicellular sheets into single cells (**Figure S10A-C**).

We then tested whether single cells resulting from sheet dissociation were viable and could survive desiccation. We rehydrated samples by adding artificial seawater three hours after desiccation, thus mimicking refilling by waves. This brought salinity back down to ~50 ppt and was followed by the reappearance of sheets as soon as 24 hours post-rehydration as
well as by a continuous increase in the number of colonies over the following days (Figure 4D-E).

We assessed the mechanism of colony re-formation by time-lapse microscopy of desiccated cultures after rehydration and captured instances of unicellular flagellates engaging in both clonal and aggregative multicellular development (Figure S9D; movie S18). Taken together, our results show that C. flexa sheets dissociate into non-motile single cells during gradual evaporation, that these solitary cells can survive complete desiccation, and that sheets reform after rehydration through both clonal division and aggregation. These findings are consistent with our field observations, where we recovered sheets after rehydrating soil samples from a dry splash pool (Figure 3I-K and Table S1), and further support the existence of a desiccation-resistant form of C. flexa.

## C. flexa sheets dissociate and differentiate into unicellular cysts at high salinity

We set out to further investigate the phenotype of desiccation-induced single cells. In diverse protists, resistance to desiccation is often achieved by differentiation into cysts<sup>61-63</sup>, i.e. dormant cells with reduced metabolic activity and minimal or arrested proliferation. Encystation often entails significant morphological changes such as rounding of the cell body, changes in cell volume, flagellar loss, and formation of a protective cell wall<sup>55,61,64-70</sup>. We monitored cellular morphology by DIC microscopy during gradual evaporation and observed structural changes that occurred asynchronously, resulting in a heterogeneous cell population (Figure 4F-I). At 3X salinity, most cells had dissociated from their colonies and bore a round cell body without microvilli and often without flagellum (Figure 4G-I). Additionally, some cells exhibited filopodia-like protrusions (Figure 4G-H). Membrane and F-actin staining of desiccation-resistant cells (hereafter ‘cysts’) confirmed that they lacked a collar complex (Figures 4J-L and S11) but also revealed that a transient F-actin cortex, detectable at 3X salinity but lost above 6X (close to saturation; Figures 4K-L and S11). This cortex might contribute to protecting the plasma membrane against osmotic stress during early differentiation<sup>71-74</sup>.

Next, we determined whether C. flexa cysts actively proliferate by quantifying cell growth during gradual evaporation in the laboratory. We found that growth was arrested above 2X salinity, with net cell loss above 3X salinity (Figures 4M and S12). These effects may be attributable to a halt in the cell cycle and cell death due to hypersaline stress, similar to changes observed during encystation in other protists<sup>36,61,64,70,75</sup>.
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Finally, morphometric analysis of cell body and nucleus volume showed that cysts had a larger nucleus-to-cytoplasm ratio compared to flagellates in 1X salinity (p=3.4e-05; Figures 4N and S13). An increase in the nucleus-to-cytoplasm ratio frequently correlates with cell quiescence<sup>76</sup>, consistent with the growth arrest in C. flexa cysts (Figures 4M and S12). To confirm that these changes were induced by hypersaline stress, we directly increased the salinity of C. flexa cultures by addition of seawater salt and observed similar morphological changes and arrest in cell growth (Figure S10).

The suite of drastic and reversible cellular changes induced by hypersalinity in C. flexa (Figure 4O) resembles encystation in diverse other protists<sup>61,64</sup>. During the natural evaporation-refilling cycle of a splash pool, C. flexa thus likely alternates between a multicellular flagellated form (at low salinity) and a unicellular cyst-like form (at high salinity), suggesting these two phenotypes each confer an adaptive advantage in their respective environment. We therefore directly tested these putative advantages in laboratory conditions.

**Multicellular sheets and unicellular cysts are respectively advantaged at low and high salinity**
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We further wondered whether multicellularity, by contrast, was advantageous in the other phase of the cycle, marked by low salinity. It has been speculated that multicellularity in choanoflagellates might enhance feeding via cooperative hydrodynamic interactions between flagella<sup>77-79</sup>, although attempts to test this hypothesis in the model choanoflagellate *Salpingoeca rosetta* have yielded conflicting results<sup>77,80,81</sup>. To test for a feeding advantage in multicellular *C. flexa* sheets, we quantified the capture of fluorescent bacteria in sheets and dissociated single flagellates (**Figure 4Q-R**). We found that sheets captured more than twice as many fluorescent bacteria per cell as single cells (**Figure 4Q-R**). This suggests that multicellularity confers a prey capture advantage at the low salinity level compatible with maintenance of a functional collar complex, and therefore with feeding.

## C. flexa cell density in natural splash pools is sufficient for aggregation

We next challenged our model by testing whether aggregative development can plausibly occur in the natural context of Curaçao splash pools. A pre-requirement for aggregation is sufficient cell density, as cells need to be at above a minimum concentration to encounter each other and aggregate. We quantified *C. flexa* cell density in 12 splash pools and found that cell density ranged from 30 to 4.4*10<sup>4</sup> cells/mL. 5/12 splash pools had a higher estimated cell density than the minimum density used in laboratory aggregation experiments (from 1.0*10<sup>4</sup> to 6.7*10<sup>5</sup> cells/mL). Thus, *C. flexa* can be present in nature in sufficient density to allow aggregation.

## Aggregation is constrained by kin recognition

To further assess the relevance of aggregation to the life history of *C. flexa*, we decided to test for additional biological signatures of aggregation. A frequent feature of aggregative multicellularity is kin recognition – i.e. the ability to discriminate kin versus non-kin, thereby restricting aggregation to closely related strains. This is classically interpreted as an adaptation to restrict the spread of cheater mutants<sup>15</sup>. We set out to test kin recognition between three *C. flexa* strains: the one used in laboratory experiments above (Strain 1, originally isolated in 2019) and two additional strains isolated in 2023 during Exped-B, Strains 2 and 3 (**Figure 5B-C**). Each strain was independently isolated from a different splash pool
by manual isolation of a single sheet colony followed by amplification in laboratory cultures (resulting in "single-sheet-bottlenecked" cultures). Several clonal lines were established from each strain by single-cell bottlenecking.

To test for genetic divergence between these strains, we first sequenced, assembled and annotated the reference C. flexa genome by a combination of long-read and Omni-C sequencing. The final assembly comprised 56 MB and 14,084 genes across 528 scaffolds, with a BUSCO completeness score of 82.8%. This represents the third high-quality choanoflagellate genome after M. brevicollis and S. rosetta⁸²,⁸³. We then performed short-read sequencing of Strains 1, 2 and 3, as well as of three clonal (single-cell-bottlenecked) descendants for each strain. 193,564 SNPs were detected with high confidence across all samples, allowing us to reconstruct a phylogenomic tree in which all three parent strains were clearly delineated from each other (Figure 5D and supplementary files S5-S6). All single-cell-bottlenecked clones clustered with their strain of origin with high support, indicating significant genetic divergence between strains but little to no diversity within strains. Single-sheet-bottlenecked cultures contained a similar amount of genetic diversity as single-cell-bottlenecked clones (Figure S16). The latter suggests either clonal development of the originally isolated sheets (as in Figure 1E-G) or loss of genetic diversity during sheet isolation and establishment of laboratory cultures (see Methods).

We then tested whether aggregation between strains was constrained by kin recognition. We dissociated sheets of all three strains, stained them with different fluorophores, and assessed aggregation between all pairwise combinations of strains (seeded at equal initial cell density, as in Figure 2C). All strains aggregated into sheets within hours, confirming aggregation as a widespread behavior in newly isolated strains. Interestingly, cells of Strains 1 and 2 preferentially aggregated with their own strain when combined, indicating kin discrimination (Figure 5E-H and S15). By contrast, other pairwise combinations of strains showed no quantifiable preference for cells of the same strain and readily aggregated into chimeric sheets (as did cells of the same strain stained with different fluorophores, consistent with earlier results; Figure 2C).

These results suggested the existence of a kin recognition system in C. flexa, similar to those described in other aggregative multicellular organisms such as dictyostelid social amoebae. In the latter, kin recognition is mediated by polymorphic surface receptors rich in immunoglobulin domains and other adhesion domains¹⁷,⁸⁴. In Dictyostelium, kin recognition receptors were originally identified by their high ratio of non-synonymous to synonymous substitutions (dN/dS), a signature of diversifying selection¹⁷. We reasoned that, if C. flexa
365 indeed possesses a kin recognition mechanism, candidate kin recognition loci might be 366 revealed by screening the genomes of Strain 1 and Strain 2 for high dN/dS loci. We identified 367 466 genes with high dN/dS ratio regions, that encoded predicted proteins with a significant 368 overrepresentation of domains consistent with an adhesion (immunoglobulin, cadherin, EGF, 369 fibronectin, HYR) or a signaling (protein tyrosine phosphatase, SH2 domain) function (Figure 370 5I-K). Many of these genes additionally encoded predicted transmembrane domains, with 371 adhesion domains on the extracellular side and signaling domains on the cytoplasmic side 372 (Figure 5J-K). This shows that multiple polymorphic receptors potentially mediating kin 373 recognition can readily be identified between diverse C. flexa strains, although some of these 374 proteins might perform other functions (such as capture of bacterial prey). Determining their 375 function will require functional genetic tools, that currently do not exist for C. flexa. 376

377 Taken together, our findings support a model of C. flexa life history that correlates with the 378 evaporation-refilling cycle of splash pools characteristic of their natural habitat (Figure 6). C. 379 flexa sheets are found in water-filled splash pools with salinities below ~3-fold that of natural 380 seawater. As gradual evaporation proceeds, salinity increases, and sheets dissociate and 381 differentiate into solitary, non-motile, and non-proliferative cysts (some of which undergo cell 382 death). These cysts can survive complete desiccation and persist in the soil of the splash 383 pools. After splash pool refilling, cysts regrow a collar complex and transition back into free- 384 swimming flagellates that develop into multicellular sheets by aggregation and/or clonal 385 division, a phenotype that more efficiently captures bacteria and presumably supports faster 386 cell proliferation. 387

388 Discussion 389

390 Our results show that C. flexa can form multicellular sheets through a combination of 391 clonal division and aggregation following the rehydration of desiccated cysts. This mixed 392 developmental strategy might represent an adaptation to the ephemeral nature of splash 393 pools, allowing faster multicellular development by simultaneous action of both mechanisms. 394 Indeed, one key advantage of aggregation is speed: across eukaryotes, 395 aggregativemulticellular forms often complete multicellular development from a unicellular 396 state in only a few hours¹⁹,¹⁹,⁸⁵. By contrast, estimates of the duration of the cell cycle in nature 397 (that constrains the speed of clonal development) for diverse micro-organisms range from 398 one to eleven days (Table S3). The clonal-aggregative multicellularity of C. flexa might thus
serve as a versatile strategy for robust establishment of multicellularity across a broad range of environmental conditions: indeed, aggregative and clonal multicellularity are subjected to different constraints, as they respectively depend on cell density and on the possibility of cell division (the latter, in turn, being constrained by salinity and availability of bacterial prey). In our laboratory experiments, *C. flexa* appeared capable of both purely clonal development (at low cell density and under stable permissive conditions for proliferation) and purely aggregative development (at sufficient cell density, even if proliferation is compromised), as well as intermediate modes.

The environmentally entrained life history transitions of *C. flexa* are in line with other recently described examples. Unicellular-to-multicellular switches are controlled by fluctuating environmental parameters in other species, such as salinity for cyanobacteria in brackish environments<sup>86</sup> and periodic flooding for cave bacteria attached to surfaces<sup>87</sup>. A selective advantage for regulated life cycles in fluctuating environments has also been supported by laboratory experiments in yeast<sup>88</sup> and by theoretical models<sup>5,89,90</sup>. The phenotypic plasticity of *C. flexa* might thus enable it to reap both the benefits of multicellularity (for feeding and collective behavior) under permissive conditions, and of differentiation into solitary cysts for individual survival under harsh conditions.

The selective advantage of multicellularity has been actively debated<sup>78,91-93</sup> including in choanoflagellates<sup>77,78,94-98</sup>. In *C. flexa*, the prey capture efficiency of cells within sheets surpasses that of solitary flagellated cells, suggesting a feeding (and likely reproductive) benefit of multicellularity. Indeed, cooperative feeding is observed among various protists and has been postulated as a selective factor for the evolution of multicellularity<sup>99</sup>. However, multicellularity might confer other advantages to *C. flexa*. For example, collective contractility underlying whole-colony inversion allows emergent behaviors such as photokinesis (preferential accumulation of sheets under light), which has been shown to require multicellularity<sup>41</sup>. Yet another advantage might be decreased vulnerability to size-selective predators, as described in other protists<sup>19,78,92,100</sup> (but see <sup>97</sup>).

The clonal-aggregative multicellularity of *C. flexa* contrasts with that of previously characterized multicellular choanoflagellates, which appear to develop solely clonally<sup>24,26,101-103</sup> – the best-characterized example being the development of rosette colonies in *S. rosetta*<sup>26</sup>. Beyond choanoflagellates, both clonal and aggregative multicellularity are widely present, with a discontinuous occurrence, across unicellular holozoans<sup>2,25</sup> and eukaryotes as a whole<sup>25,104-106</sup>. The scattered distribution of clonal and aggregative multicellularity is currently interpreted
as reflecting independent evolutionary origins of both types of development³. However, the discovery of clonal-aggregative multicellularity in a choanoflagellate raises the intriguing alternative possibility of evolutionary interconversions between aggregative and clonal multicellular development (via a mixed intermediate). Indeed, clonal-aggregative multicellularity might be more widespread than currently appreciated. Notably, clonality and aggregation have been proposed to cooperate in the formation of diverse structures across prokaryotes and eukaryotes, including bacterial biofilms¹⁰⁷, experimentally evolved clusters of bacteria¹⁰⁸, predator-induced groups of freshwater algae (which can even combine different species)¹⁰⁹, and unicellular, but multinucleated syncytia in reticulopodial amoebae¹⁸. Although these diverse processes might not all represent bona fide multicellular development¹¹⁰ (as the resulting structures seem to lack controlled shape, size, or collective behavior), they show that cell division and aggregation behaviour can coexist.

Aggregative multicellularity comes with a well-known evolutionary challenge: one aggregate can combine cells of different ancestries and potentially different genotypes, thus raising the possibility of conflict¹¹¹⁻¹¹³. Aggregates are notably vulnerable to cheater genotypes, which reap the benefits of multicellularity without contributing their fair share of the cost of multicellular development. Such cheaters have for example been described in *Dictyostelium*¹¹⁴, *Myxococcus*¹¹⁵, and in flocculating yeast¹¹⁶. The difficulty of resolving such conflicts, alongside that of coordinating complex functions in aggregates potentially combining multiple genotypes¹⁹, might limit the evolutionary potential of aggregative multicellularity. Indeed, all five eukaryotic lineages with "complex multicellularity" (i.e. controlled three-dimensional morphogenesis and spatial cell differentiation), including animals, fungi, and land plants, develop clonally rather than by aggregation²⁵,¹¹⁷, and aggregative multicellular forms instead tend to be transient and facultative¹¹⁶⁻¹²⁰. Aggregative development often occurs in conditions where the benefit of fast development outweighs the cost of potential genetic conflict – either because adverse conditions make cell proliferation (and therefore clonal development) infeasible¹⁹,³⁶,¹²¹ or because fast multicellular development is required as an "emergency response" to acute stress¹⁹,¹¹⁶,¹²². Indeed, aggregative multicellularity is frequently induced by stressors such as nutrient depletion³⁶,⁴⁵ or presence of predators¹⁰⁰,¹²¹,¹²³. In such situations, aggregation allows for rapid establishment of a multicellular structure that confers a selective advantage (e.g. increased size, shielding of inner cells, or elevation from the substrate by formation of a stalk). Often, the endpoint of stress-induced aggregative multicellularity is the formation of a fruiting body: a mass of non-
proliferative cells that disperses into single cells before resuming metabolic activity and proliferation (as described in dictyostelid amoebae<sup>3,45,124</sup>, multiple other eukaryotes<sup>125,126</sup>, and myxobacteria<sup>127</sup>). Interestingly, and by contrast, *C. flexa* sheets, although they similarly develop under time pressure, are proliferatively and behaviorally active. Thus, the ecological niche of mixed clonal-aggregative multicellularity might differ from that of typical aggregative forms.

Nonetheless, the risk posed by cheaters can be addressed by strategies that limit aggregation to fellow co-operators, either directly by co-operator recognition (so-called "green beard" mechanisms<sup>128,129</sup>) or indirectly by preferential aggregation with close relatives. Restriction of aggregation to relatives can either be enforced actively by kin recognition (usually mediated by polymorphic loci<sup>130-132</sup>) or be facilitated passively by spatial structuration of the environment that can limit dispersal<sup>133,134</sup>. The latter mechanism might also be relevant to *C. flexa*, as splash pools are collections of physically disconnected environments that might lead to geographic divergence of genotypes (as shown for other organisms in similar environments, such as *Daphnia* metapopulations in tide pools<sup>135</sup>). Splash pools might thus favor the evolution of aggregative multicellularity by initially relaxing the need for strong kin recognition mechanisms (which *C. flexa* nonetheless displays). This is in line with recent models proposing an “ecological scaffolding” function for patchy environments in the emergence of multicellularity<sup>136</sup>. In the future, deeper characterization of molecular kin recognition mechanisms, natural genetic diversity and dispersal in *C. flexa* might shed further light on these questions.

The complex life history of *C. flexa* illustrates the phenotypic and developmental plasticity of close unicellular relatives of animals and thus lends additional support to the emerging concept of a pre-metazoan origin for complex life cycles<sup>20,137</sup>. Beyond this, our study establishes *C. flexa* as a model to study the facultative multicellular development of a close relative of animals in its natural context. This contrasts with other well-characterized facultatively multicellular holozoans, such as *S. rosetta*<sup>21</sup> and *C. owczarzaki*<sup>138</sup>, which could both be isolated only once from their natural environment and in which studies of unicellular-to-multicellular transitions are thus inevitably restricted to laboratory settings. In the future, we expect the dialogue between field and lab studies of *C. flexa* to continue, clarifying questions such as the selective advantage(s) and ecological implications of multicellularity and unicellularity, as well as the mechanism of kin recognition.
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## Data availability

503 The annotated reference C. *flexa* genome has been deposited on Zenodo under doi:

504 10.5281/zenodo.13837466. Raw short reads for Strains 1, 2 and 3 have been deposited on Zenodo

505 under doi: 10.5281/zenodo.13837614.

506

507

## Code availability

508 The codes used for SNP analysis are deposited on Github:

509 https://github.com/uhoro/RRRR_SNP_analysis

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## Acknowledgements

513 We thank Nicole King for support during this project and for providing useful feedback. We thank the

514 personnel of the Shete Boka National Park in Curaçao for access to the field site. Special thanks to

515 Tess Linden for help during preliminary field experiments and feedback on the manuscript, as well as

516 to Geneviève Milon and Noah Whiteman for advice on laboratory and field experiments, respectively.

517 Yashraj Chavhan, Seemay Chou, and Stefania Kapsetaki provided useful feedback on the preprint

518 version of the manuscript. Marvin Albert, Stéphane Rigaud and Jean-Yves Tinevez at the Image

519 Analysis Hub at the Institut Pasteur provided advice on image analysis. Pierre-Henri Commere and

520 Sebastien Megharba from the Flow Cytometry platform at Institut Pasteur provided technical support

521 with single-cell isolation of C. *flexa* by flow cytometry. We acknowledge the help of the HPC Core

522 Facility of the Institut Pasteur for bioinformatic analyses. We thank Thomas Swale (Dovetail

523 Genomics/Cantata) and Tara Rickman (Phase Genomics) for help with genome sequencing and

524 assembly and for drafting the relevant Methods sections. We also thank members of Brunet and King

525 laboratories for feedback and useful discussions and Jaime Ramirez and Kayla Dinshaw for technical

526 support during preliminary experiments. NRR is supported by the European Union's Horizon Europe

527 research and innovation funding program under a Marie Skłodowska-Curie Actions grant (Grant

528 Agreement ID: 101106415). JRR is supported by a National Science Foundation Graduate Research

529 Fellowship Program (grant no. 1752814). UH is supported by the PPU program of the Institut Pasteur.

530 Work in the King lab (TB, JRR, BTL, MCC) is supported by the Howard Hughes Medical Institute. JLS

531 is a Howard Hughes Medical Institute Awardee of the Life Sciences Research Foundation. TB and work

532 in the Brunet lab are supported by the Institut Pasteur (G5 package), the ERC (EvoMorphoCell, grant

533 agreement ID 101040745), the Vallee Foundation Inc, and the CNRS (UMR 3691). Views and opinions

534 expressed are, however, those of the author(s) only and do not necessarily reflect those of the

535 European Union or the European Research Council. Neither the European Union nor the granting

536 authority can be held responsible for them.

537

538

## Author contributions

539 NRR, JRR, TB conceptualized the study and coordinated the work across coauthors. MCC, BTL and

540 TB performed preliminary fieldwork that identified the Shete Boka site and gave first insights in the C.

541 *flexa* salinity range. NRR, JRR and TB collected fieldwork data included in this study. NRR, JRR and

542 TB performed all laboratory experiments, strain isolations and bioinformatic analyses (except those

543 specified below). BTL performed preliminary aggregation assays and aphidicolin treatments. YF

544 optimized live imaging of dual-labelling aggregation experiments and performed growth curves under

545 addition of salts. JLS performed genome assembly and annotation. UH performed SNP analyses with

546 technical guidance from JLS. CC performed single-cell bottleneck isolations and generated the material

547 for genome sequencing of Strains 1, 2 and 3. EATH and MAV provided support for fieldwork studies.
548 EATH performed drone orthomap imagery. JRR wrote the first draft of the manuscript which was
549 subsequently edited by NRR and TB. NRR made the figures with feedback from JRR and TB. TB
550 supervised the study. All authors contributed to the review of the manuscript before submission for
551 publication and approved the final version.

552

**Inclusion & ethics statement**

553

554 Local and regional researchers who supported or performed part of the Curaçao fieldwork are authors
555 of the present manuscript.

556

**Competing interests**

557

558 JLS is an advisor for ForensisGroup Inc.

559

**Materials and Methods**

560

See detailed Materials and Methods in the **Supplementary Information** file.

561

562

**Supplementary Material**

563

564 Supplementary Figures S1-S16, Tables S2-S3 and legends for Table S1, Supplementary Files S1-S6
565 and Supplementary Movies S1-S18 are provided in the **Supplementary Information** file.

566
<figure><img src="image_1.png" /><figcaption>**Figure 1.** *Choanoeca flexa* sheets can develop clonally but display aggregative features.</figcaption></figure>

(A) Choanoflagellates (turquoise) are the sister group to animals (Metazoa). The phylogenetic relationships depicted are based on several recent phylogenomic studies<sup>25,27,32,41,139–142</sup>. Uncertain positions are represented with polytomies. (B) Brightfield image of a *C. flexa* multicellular colony (“sheet”) in its relaxed conformation (B’). B”, white square: zoom-in showing flagella (magenta pseudocolor) and direct cell-cell contacts between collars (green pseudocolor). Scale bar in B”: 10 µm. (C) Diagnostic morphological features of a choanoflagellate cell. *C. flexa* cells within a sheet are linked by their collars (orange arrow). (D) 3D reconstruction of an Airyscan Z-stack of a fixed sheet exhibiting an inverted conformation (D’), with cell bodies stained with a membrane/cytoplasmic dye (FM™ 1-43FX, magenta, which distributes to the membrane and cytoplasm of cells following fixation), and microvilli stained with a filamentous actin (F-actin) dye (phalloidin Alexa 488, green). (E) Stills from a brightfield timelapse movie of clonal *C. flexa* sheet development by serial cell division from a single *C. flexa* swimmer cell (white arrowhead). After each division, the sister cells remain adhered to each other by direct cell-cell contacts between collars (black arrow). Note that cells retract their flagellum during

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583 division. Time scale hh:mm. (F) Cell lineage tracing as a function of time in E shows that cells divide
584 asynchronously during colony formation, taking ~8-10 hours between each round of division. (G) Stills
585 from a brightfield timelapse movie depicting a medium-sized C. flexa sheet (flagella-in conformation)
586 that expands in cell number both by cell division (pseudocoloring in orange, pink, blue and yellow) and
587 by cellular aggregation (white arrow, pseudocoloring in green). Time scale hh:mm. (H) Schematics of
588 C. flexa clonal multicellular development observed under laboratory culture conditions: unicellular
589 flagellate (swimmer) cells can divide clonally to initiate a colony; in turn, colonies can increase in cell
590 number by clonal division, maintaining cell-cell contacts through direct adhesions in their collar (orange
591 arrowhead). The hypothesis that C. flexa sheets might be able to develop purely by aggregation is
592 tested in Figure 2. Figure related to **Figure S1** and **movies S1-S3**.
593
<figure><img src="image_2.png" /><figcaption>Figure 2. C. flexa can develop by aggregation.</figcaption></figure>

(A) Schematics of the experimental assay to test C. flexa aggregation. Colonies are mechanically disassembled into free-swimming single cells and the resulting dissociated single cells are live imaged.

(B) Stills from a brightfield timelapse movie showing that dissociated single cells (white arrowheads) quickly re-formed colonies by cellular aggregation. Note that two colonies can fuse together by agglomeration (black arrowheads). (B') Binary image mask of stills in B. Time scale hh:mm. (C) (Upper panel) Dissociated single flagellate cells labelled with Cell Trace CFSE (green) or Cell Trace Far Red (magenta) and mixed in a 1:1 ratio will form chimeric colonies 24 hours post-dissociation (hpd) if aggregation occurs. (Lower panel) 3D reconstructions of Airyscan Z-stacks of a sheet formed by aggregation from dissociated single cells labelled as depicted above and fixed 24 hpd, with additional filamentous actin (F-actin) staining (phalloidin Alexa 405, blue). (D) 3D reconstructions of Airyscan microscopy images of single cells fixed after 10 minutes (n=12), 30 minutes (n=12), 2 hours (n=19), and 24 hours (n=16) post-dissociation stained for membrane/cytoplasm (FM™ 4-64FX, magenta) and F-actin (phalloidin Alexa 488, green). Note that cells frequently show unaligned apico-basal polarity and diverse cell orientations at early timepoints (white arrowheads). Time scale hh:mm. (E) Quantification of cell number per colony during aggregation (from the experiment in D) in two independent biological replicates. Black circles: mean. Error bars: standard deviation. Diamonds: mean

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612 values of independent biological replicates. (F) Quantification of particle area during an aggregation

613 time course of dissociated single cells pre-treated with 17 µg/mL aphidicolin overnight in three

614 independent biological replicates. DMSO: negative control. Figure related to **Figures S2-S4** and

615 **movies S4-S13**.

616
<figure><img src="image_3.png" /><figcaption>Figure 3. Natural evaporation-refilling cycles constrain the occurrence of multicellular *C. flexa* in the wild.</figcaption></figure>

(A) Map of Curaçao and location of Shete Boka National Park (turquoise star) where fieldwork data was collected in Exped-A, Exped-B and Exped-C expeditions (12° 22' 5.718" N, 69° 06' 56.916" W).

(B) Representative photograph of the landscape in Shete Boka, including splash pools where *C. flexa* sheets can be found (white arrowheads).

(C) Schematics of a splash pool natural cycle of seawater evaporation, desiccation, and refilling.

(D) Maps showing the locations of sampled splash pools in Exped-A and in Exped-B expeditions. In Exped-A (D), samples were collected from splash pools along ~2 km of Shete Boka coastline (n=79).

15 of these splash pools were randomly selected for daily monitoring during eight days (stars) (shown are results from the first day of monitoring). Colors indicate whether sheets were found (magenta) or not found (turquoise) upon microscopic inspection.

In Exped-B (D'), a random number generator was used to select the randomized sampling location (purple pin,

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204 m upstream of Boka Wandomi). Samples were collected from splash pools within an area of 10 m
by 4 m (n=71). (E) Distribution of salinity of splash pools surveyed in Exped-A (circles) and Exped-B
(triangles). For daily monitored splash pools, measurements from the first day of monitoring are shown,
except for those where sheets were observed more than one day. In that case, the minimum and
maximum salinity values are reported. Magenta and turquoise colors indicate that sheets were
respectively found and not found in splash pool samples. The observed natural limit of salinity where
sheets were found (128 ppt) is indicated by an orange dotted line, and the average seawater salinity
measured in the *bokas* with a gray dotted line. Gray area: salinity saturation (>280 ppt). (F) Brightfield
image of a sheet observed in a splash pool sample. (G) Representative images of a splash pool near
Boka Kalki (Sp15) followed for eight days, showing recorded salinity in the upper left. This splash pool
was completely desiccated on day 5 (right). Dashed line: splash pool outline. (H) Salinity (upper panel)
and depth (lower panel) measurements in three representative splash pools followed over eighth days.
Shown are one splash pool that experienced evaporation but not complete desiccation (Sp50), one
splash pool that experienced complete desiccation (Sp15), and one splash pool that experienced both
desiccation and refilling (Sp69). Filled and empty circles indicate that sheets were respectively found
and not found in splash pool samples. Gray area: salinity saturation. Dashed lines: dry periods. Orange
circles: refilling events. (I) Recovery of sheets from soil samples collected from a splash pool (Sp12)
during its dry period in Exped-A. Soil samples were collected every day for eight days and were
independently rehydrated in the laboratory with filtered seawater from the *bokas*, reconstituting salinity
back to 42 ppt. Each rehydrated soil sample was monitored over five days for sheet re-appearance.
Filled circle: sheets observed. Empty circle: sheets not observed. Dashed empty circle: sample not
inspected. rh: rehydration. (J) Brightfield image of a soil-recovered sheet (same experiment as in I)
collected after 3 days of desiccation and rehydrated in the laboratory. (K) Brigthfield image of a soil-
recovered sheet independently collected and rehydrated from a splash pool sample (Soil #18) in Exped-
C. Time post-rehydration in J-K is depicted in the lower left. Figure related to **Figures S4-S9, movies
S14-S16, Tables S1-S2 and Supplementary Files S1-S4.**
<figure><img src="image_4.png" /><figcaption>**Figure 4.** Experimental evaporation-refilling cycles cause reversible transitions in and out of multicellularity.</figcaption></figure>

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**Figure 4.** (A) Laboratory experimental evaporation setup in a 30°C incubator. Note that plates were kept with the plate's lid off (gradual evaporation condition) or the plate lid's on (low-evaporation control) over a grid and were covered with a plastic box to avoid contamination and favour air exchange. (B) Salinity monitoring of followed-up splash pools during an 8-day time course from Exped-A expedition. Splash pools during the first 6 days of the time course gradually evaporated, and hence they were used to model splash pool evaporation dynamics. Splash pools with less than 3 points outside saturation range (gray area) were not included in the modelling (i.e. Sp12, Sp42, and Sp70; see **Figure S7**). (C) Normalized inverse of salinity over time during gradual evaporation comparing empirical data from splash pools in B versus the laboratory evaporation setup (gray) in A. (D) Quantification of salinity (upper panel) and fraction of cells in colonies (magenta) or in single cells (turquoise) (lower panel) during a 9-day gradual evaporation time course. Dark gray rectangle: complete desiccation (dry period). Orange dotted line: observed natural salinity limit of sheet occurrence. Gray area: salinity saturation. Lines correspond to mean values and shadowed area to standard deviation. Experiment performed on two independent biological replicates. (E) Stills of C. flexa sheets during the same 9-day gradual evaporation timecourse in D, showing salinity (in ppt, upper left) and time (upper right). Sheets had completely dissociated into single cells (white arrowheads) after 80 hours of gradual evaporation. Multicellular sheets were observed again after complete desiccation and rehydration with 1X ASW. (F-I) Brightfield images of C. flexa showing morphological changes during gradual evaporation, showing salinity (in ppt, upper right). At ~1X seawater salinity (F), C. flexa occurs in the form of multicellular sheets of flagellated cells (white square). Green pseudocolor: collar. Magenta pseudocolor: flagellum. During gradual evaporation (G-I), sheets dissociate into unicellular cysts. Cysts often lack a collar of microvilli (mv), often lack a flagellum (fl), and can exhibit filopodia-like protrusions (magenta arrowheads). (J-L) AiryScan micrographs of C. flexa cells fixed during gradual evaporation stained with a membrane (FMTM 4-64FX, magenta) and F-actin (phalloidin Alexa 488, green) dyes, showing salinity (in ppt, upper right). A flagellate cell (J, low-evaporation control) exhibits distribution of F-actin in the collar of microvilli (mv, white arrowhead) and filopodia (fp, white arrowhead). Gradual evaporation triggers a morphological change from a flagellate to a cyst, showing filopodia-like protrusions (magenta arrowheads) and a transient actin cortex (cx) at early stages of evaporation (K, white arrow) which disappears as salinity approaches saturation (L). (M) Growth rate of cells at different salinities during gradual evaporation in three independent biological replicates. Black circles: mean. Error bars: standard deviation. n=28, p by the Mann-Whitney U test. Diamonds: mean values of independent biological replicates. (N) Quantification of the nucleus-to-cytoplasm ratio in cysts (n=27) and flagellates (n=38) in three independent biological replicates. Black circles: means. Error bars: standard deviations. Diamonds: mean values of independent biological replicates. p by the Mann-Whitney U test. (O) Schematic summarizing phenotypic changes experienced by C. flexa cells during gradual evaporation. At early stages of evaporation (turquoise), flagellate cells differentiate into cysts by losing their collar and flagellum and show a temporal actin cortex. At later stages of evaporation (desiccation, magenta), cysts lack an actin cortex and show a higher nucleus-to-cytoplasm ratio. Rehydration (refilling, purple) induces a cyst-to-flagellate cell transition, where cells regenerate their collar and flagellum and re-form multicellular sheets. (P) Quantification of cell survival after 12 hours of desiccation in multicellular sheets (n=6 technical replicates) to unicellular cysts (n=6 technical replicates). Diamonds: means. Error bars: standard deviations. p by the Mann-Whitney U test. (Q) C. flexa multicellular sheets are more efficient at capturing bacteria than unicellular flagellates. Number of labelled bacteria captured per flagellate cells in multicellular colonies (n=9) or in cultures of unicellular flagellates (n=9) in three independent biological replicates. Black circles: means. Error bars: standard deviations. Diamonds: means of independent biological replicates. p by the Mann-Whitney U test. (R) DIC images of multicellular sheets (upper panel, white arrowheads) capturing fluorescently labelled bacteria using Bactoview dye (upper panel: green; lower panel: white). Figure related to **Figures S9-S14 and movies S17-S18**.
<figure><img src="image_5.png" /><figcaption>Figure 5. Aggregation of *C. flexa* is compatible with cell densities measured in the field and is constrained by kin recognition between genetically different strains.</figcaption></figure>

(A) Cell density in nature is sufficient for aggregation. Cell density measured in 12 splash pools (A to L) compared to the range used in laboratory aggregation experiments (gray area). Diamonds are the average estimate, and bars indicate uncertainty (ranging from minimal to maximal possible estimated values).

(B) Source location of the three distinct strains of *C. flexa* used in kin recognition experiments. (C) Aerial view of two neighboring splash pools in the Exped-B site, M44 and M60, from which strains 2 and 3 were respectively isolated. (D) Phylogenomic tree of strains 1, 2 and 3, including single-sheet-bottleneck cultures (SSB) and three to four single-cell bottlenecked clones from each strain. (E) Cells of the same strain (here strain 1), marked with magenta or green fluorophores, readily aggregate together regardless of color, resulting in dual-colored colonies (white arrowhead; compare Fig. 2).

(F) Cells of strains 1 and 3 (clone 1), marked with distinct fluorophores, readily aggregate into chimeric sheets (white arrowhead). (G) Cells of strains 1 and 2 (clone 1), marked with distinct fluorophores, preferentially aggregate with their own strain, resulting in colonies with little or no chimerism (magenta and green arrowheads). This indicates kin recognition. (H) Quantification of kin recognition in different

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727 pairwise strain combinations (see **Figure S15A-C** and Methods). The kin recognition index is calculated
728 based on the variance of the proportion of green (or red) cells across sheets for a given combination of
729 strains (as in¹⁴³). A unimodal distribution centred around a median peak at 50% indicates no kin
730 discrimination, while a bimodal distribution with two distinct peaks (and thus with higher variance)
731 indicates kin discrimination (see Methods). All experiments were performed in two independent
732 biological replicates. *p* = 4.6e-3 by a one-way ANOVA. (I) Nonredundant top InterProScan domain
733 annotations enriched in candidate kin recognition loci (identified by high Ka/Ks ratio). See **Figure S17**
734 for full list. (J) Four polymorphic candidate kin recognition loci encoding transmembrane proteins with
735 potential roles in cell-cell adhesion and signalling. Upper row: number of coding differences (aminoacid
736 substitutions) between the strain of interest (2 or 3) compared to strain 1. Lower row: domain
737 architecture predicted with InterProScan (see domains legend), highlighting non-cytoplasmic,
738 cytoplasmic (cyto) and transmembrane (tm) regions.
<figure><img src="image_6.png" /><figcaption>Figure 6. The environmentally entrained clonal-aggregative life cycle of C. flexa</figcaption></figure>

741 Summary schematic of C. flexa mixed clonal-aggregative development entrained by natural splash pool evaporation-refilling cycles. Multicellularity is favoured in low salinity, where sheets exhibit enhanced growth and prey capture (turquoise): (1) unicellular flagellate cells can aggregate and/or divide clonally to initiate multicellular sheet development, maintaining direct cell-cell adhesions in their collar (orange arrowheads); (2) sheets expand by clonal cell division, by aggregation of individual cells, and by sheet fusion. Gradual evaporation leads to increase in salinity and favours unicellularity (magenta): (3) C. flexa remains multicellular in up to 3-fold salinity, but hypersaline conditions (>3-fold salinity) trigger sheet dissociation into single cells that are incapable of proliferation and occasionally to cell death (4). Gradual evaporation, instead, results in the differentiation of flagellates into cysts (5) capable of surviving desiccation. (6) Rehydration restores permissive salinity and induces differentiation of cysts into flagellates, which undergo clonal-aggregative multicellular development.

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