The history of Salpingoeca rosetta as a model for reconstructing animal origins

: Choanoflagellates, the closest living relatives of animals, have the potential to reveal the genetic and cell biological foundations of complex multicellular development in animals. Here we describe the history of research on the choanoflagellate Salpingoeca rosetta. From its original isolation in 2000 to the establishment of CRISPR-mediated genome editing in 2020, S. rosetta provides an instructive case study in the establishment of a new model organism. mechanistic investigation of the functions of conserved genes. Moreover, understanding the transition between unicellular and colonial (i.e. multicellular) life history stages in choanoflagellates was poorly understood because of the absence of multicellular development in laboratory cultures. Here we describe the history of molecular genetic approaches in choanoflagellates and the establishment of the colony-forming choanoflagellate Salpingoeca rosetta as a simple model for investigating the ancestry of animal gene function and cell biology. We focus on rosette colony development to show how discoveries from the study of this process not only revealed the biology of S. rosetta but also provided crucial insights that led to the establishment of genetic tools. We emphasize that a deep appreciation of choanoflagellate biology gained through the application of (relatively) inexpensive and general tools provided foundational knowledge required to establish methods for gene delivery, transgene expression, and genome editing.


Introduction
By virtue of their close relationship with animals ( Figure 1A) and the many animal-like features of their cell physiology and genomes, choanoflagellates have great potential as experimental models for reconstructing animal origins and core mechanisms underlying animal cell biology. Interest in choanoflagellates began following their first observations in the mid-tolate nineteenth century (Leadbeater, 2015). Early microscopists documented the fascinating collar complex, a diagnostic feature of choanoflagellates in which a picket fence of actin-filled microvilli encircles an apical flagellum (Figure 1B and C;Karpov and Leadbeater, 1998). Cells in the feeding chambers of sponges (choanocytes; Figure 1D) also contain a collar complex and these similarities in cellular architecture led Elie Metchnikoff in 1886 to hypothesize that these features were homologous and that animals and choanoflagellates shared a recent common ancestor (Metchnikoff, 1886;Brunet and King, 2020). The collar complex is now recognized as a widespread feature of animal cell biology, consistent with the results from multiple independent phylogenetic and phylogenomic comparisons that firmly established choanoflagellates as the sister group of animals ( Figure 1A; Wainright et al., 1993;Lang et al., 2002;Burger et al., 2003;Philippe et al., 2004;Ruiz-Trillo et al., 2008). Subsequent forays into comparative genomics and transcriptomics revealed that many mediators of intercellular signaling, adhesion, cell differentiation, extracellular matrix and immunity in animals are also encoded in and transcribed from choanoflagellate genomes (King and Carroll, 2001;King et al., 2003;Snell et al., 2006;King et al., 2008;Fairclough et al., 2013;Richter et al., 2018;López-Escardó et al., 2019). These shared cellular and genetic traits suggested that mechanistic studies in choanoflagellates could prove valuable for core features of animal cell and molecular biology.
Nonetheless, the value of choanoflagellates to molecular and cell biologists was historically limited by the absence of key molecular genetic approaches that would allow mechanistic investigation of the functions of conserved genes. Moreover, understanding the transition between unicellular and colonial (i.e. multicellular) life history stages in choanoflagellates was poorly understood because of the absence of multicellular development in laboratory cultures.
Here we describe the history of molecular genetic approaches in choanoflagellates and the establishment of the colony-forming choanoflagellate Salpingoeca rosetta as a simple model for investigating the ancestry of animal gene function and cell biology. We focus on rosette colony development to show how discoveries from the study of this process not only revealed the biology of S. rosetta but also provided crucial insights that led to the establishment of genetic tools. We emphasize that a deep appreciation of choanoflagellate biology gained through the application of (relatively) inexpensive and general tools provided foundational knowledge required to establish methods for gene delivery, transgene expression, and genome editing.

S. rosetta growth and life history in the laboratory
When one of us (NK) began to study choanoflagellates in 2000, choanoflagellates were known to exist in diverse unicellular and colonial forms in nature (Leadbeater, 1983(Leadbeater, , 2015, but there were no readily available colony-forming species in culture. Fortunately, a visit with Dr. Tom Nerad at the ATCC inspired him to isolate a colony-forming choanoflagellate from brackish water samples collected near Hog Island, Virginia. This species, initially named Proterospongia sp. 50818 (King et al., 2003;Ruiz-Trillo et al., 2007;Carr et al., 2008) and later renamed Salpingoeca rosetta (Fairclough et al., 2010), was isolated as a single rosette colony ( Figure  2A) that was then propagated in cereal grass media with environmental bacteria (as food) to establish a stable culture.
Within a small number of generations in the laboratory, S. rosetta cultures transitioned into a stably unicellular state ( Figure 2B) and the rosette colonies that were apparent in the first environmental samples were rarely detected, preventing their study. Diverse environmental perturbations, including changes in media content, temperature, pH, and salinity, failed to restimulate rosette formation. However, a serendipitous observation made during the preparation of S. rosetta for genome sequencing revealed how rosette development could be induced (Fairclough et al., 2013).
Like all other choanoflagellates, S. rosetta must be co-cultured with bacteria as a food source. The diversity and abundance of bacteria in choanoflagellate cultures can interfere with eukaryotic genome sequencing and assembly, so then-undergraduate Rick Zuzow treated the cultures with different cocktails of antibiotics (Fairclough et al., 2010). One antibiotic cocktail blocked all rosette development while another unexpectedly yielded a culture in which rosettes developed readily and reproducibly. Experiments with these cultures performed by then-postdoc Rosie Alegado and members of the laboratory of Prof. Jon Clardy (Harvard Medical School) revealed that a key factor in rosette induction was the presence of a previously-uncharacterized bacterial species, which we named Algoriphagus machipongonensis (Alegado et al., 2012(Alegado et al., , 2013. Addition of A. machipongonensis to a culture of single cells was sufficient to induce rosette development through serial rounds of cell division. Subsequent improvements to S. rosetta culture conditions, including co-culture with the feeder bacterium Echinicola pacifica and the development of new semi-defined media formulations (Levin and King, 2013;Woznica et al., 2016), increased the efficiency of rosette induction and later aided the establishment of transfection and genome editing methods Booth and King, 2020). These advances provided the first evidence of bacterial regulation of choanoflagellate cell state transitions and provided a means by which to control rosette development in the laboratory.
Multicellular development was not the only S. rosetta life history transition regulated by environmental cues. In a separate line of research, then-graduate student Tera Levin found that prolonged starvation (i.e. over two weeks) induced mating in S. rosetta (Levin and King, 2013). Four years later, then-graduate student Arielle Woznica found that treatment of S. rosetta cultures with the bacterium Vibrio fischeri induced S. rosetta to swarm and then mate, in this case within an hour after treatment (Woznica et al., 2017). These findings demonstrated the essentiality of bacterial cues to the regulation of choanoflagellate biology and provided two independent methods for inducing mating, which later proved critical for the establishment of genetic screens. This period of S. rosetta research demonstrated that simple alterations to growth conditions, including altering nutrient content or changing the bacterial composition, could reveal novel biology and provide levers for controlling life history transitions in the laboratory.

Forward genetics and the hunt for rosette defective mutants
Although rosette development in S. rosetta could now be readily induced by treatment with A. machipongonensis, nothing was known about the genes that regulated the process. The gold standard for discovering the genes underlying a biological process is to perform an unbiased mutant screen, but such screens faced multiple barriers in choanoflagellates. Methods for mutagenesis had not been established (and there were reasons to think that it might be challenging), no genetic markers had yet been identified and the cells are grown in liquid media, meaning mutants could not be isolated, propagated, and phenotyped on plates.
The same Tera Levin who previously uncovered a method for inducing mating (Levin and King, 2013; see above) systematically overcame each of these barriers, allowing the establishment of forward genetics in S. rosetta (Levin et al., 2014). She first developed methods for mutagenesis of S. rosetta with both EMS and X-rays and identified effective doses using "death curves." Next, she established a phenotypically wild type mapping strain containing over 39,000 genetic variants distributed throughout the genome. The mapping strain was clonally propagated from an isolated cell that had been exposed to high doses of mutagen, found to have no obvious mutant phenotypes (e.g. growth defects or loss of rosette development), and then sequenced to identify genetic variants across the genome that could be used for mapping mutations through crosses.
These tools set up the stage for the first genetic screen in a choanoflagellate. Over 37,000 clones were isolated by serial dilution and screened by eye to identify 16 mutants that fell into seven phenotypic classes. One mutant, Rosetteless, was of particular interest; while it lacked the ability to form rosettes, all other aspects of its life history and cell biology seemed to be wild type. By crossing the Rosetteless mutant to the mapping strain, Levin identified a genetic lesion that co-segregated with the loss of rosette development. The lesion fell in a Ctype lectin gene that we named "rosetteless" after its phenotype. Rosetteless protein is secreted into the extracellular matrix (ECM) at the basal ends of cells in rosettes ( Figure 2A, C), and the mutant allele abrogates Rosetteless protein expression, resulting in cells that grow normally except for their inability to form rosettes ( Figure 2D, C; (Levin et al., 2014)).
In two other rosette-defect mutants, Jumble and Couscous, single cells aggregated into large amorphous clumps and true rosettes were never observed to develop (Wetzel et al., 2018). Then-graduate student Laura Wetzel used crosses between these mutants and the mapping strain, coupled with bulk segregant analysis, to identify genetic lesions in two different glycosyltransferases that co-segregated with the clumping phenotype. Using new transgenic approaches (described in more detail below), Wetzel corroborated the mapping cross results and demonstrated that Jumble localizes to the Golgi, whereas Couscous localizes to puncta that may represent the endoplasmic reticulum.
Taken together, these studies demonstrated the efficacy of forward genetics for identifying genes required for S. rosetta rosette development. Moreover, the unbiased genetics approach emphasized the importance of the extracellular matrix during the development of rosettes. The criticality of the ECM for rosette development was shown independently through a recent study of biophysical constraints on rosette morphogenesis (Larson et al., 2020). Finally, the connection of genotype to phenotype in the context of the Rosetteless mutant provided a toehold against which different genome editing approaches could later be tested.

Gene delivery and genome editing in S. rosetta
While genomic, biochemical, and forward genetic approaches provided key early insights into rosette development, the absence of transgenic approaches in S. rosetta limited its experimental tractability and potential relevance as a laboratory model. From its isolation in 2000 until 2018, repeated efforts to establish transgenics failed, in large part because the field lacked methods for robustly delivering macromolecules into S. rosetta cells. Choanoflagellate cells are too small (~ 5 m) for microinjection and standard electroporation methods had thus far failed, both because of the necessity of culturing S. rosetta in sea water and because we lacked a positive control for transfection. Key advances in our understanding of S. rosetta's genetics and cell physiology that arose through our basic research into rosette development proved to be critical for overcoming these barriers.
In many single-celled organisms, electroporation with nucleic acids or proteins is physically blocked by an organic layer (e.g. a cell wall or extracellular matrix) surrounding the cell (Meilhoc et al., 1990;Shimogawara et al., 1998;Thompson et al., 1998;Benatuil et al., 2010;Kawai et al., 2010). However, because choanoflagellates lack a cell wall and prior electron microscopy studies of unicellular life stages suggested that they might be "naked," we admittedly paid little heed to the extracellular matrix in our early forays into establishing transgenics. This changed with the discovery of rosetteless (Levin et al., 2014) and improvements in electron microscopy that revealed a layer of extracellular matrix encapsulating S. rosetta cells ( Figure 3A; Dayel et al., 2011). Following lessons learned in other microeukaryotes, we therefore developed a method ( Figure 3B) to prime S. rosetta for transfection by bathing single cells in a cocktail of papain (a protease) and specific salts that gently removes extracellular proteins (dx.doi.org/10.17504/protocols.io.h68b9hw), leaving the cell membrane vulnerable to pore-forming electrical pulses delivered through electroporation, but healthy enough for the pores to seal and the cells to recover. The next step was to identify the best buffering conditions and electrical pulses to create pores in the S. rosetta membrane. We chose to use nucleofection, a specialized form of electroporation, as it has proven effective for delivering DNA and protein into difficult-to-transfect cells (Gresch et al., 2004;Hamm et al., 2002). Although the exact details of nucleofection remain proprietary, an excellent review of the biophysical principles that underlie a variety of gene delivery methods can be found in (Stewart et al., 2018). A major strength of nucleofection, particularly when used in conjunction with a plate-based platform, is that the combination of pulses and diverse transfection buffers creates a large parameter space to explore for testing a wide range of nucleofection conditions. By first removing the extracellular matrix and then applying the most effective nucleofection conditions, we achieved transfection efficiencies of 1%. (An important downside for nucleofection is the expense of the machine itself and the proprietary buffers. Therefore, we have recently established a second transfection protocol for S. rosetta using an inexpensive, square-pulse electroporator [ Figure 3C; dx.doi.org/10.17504/protocols.io.bx5vpq66]. We hope that these initial parameters will make transfection accessible to a broader community of choanoflagellate researchers and serve as the start of further optimization.) Our second big challenge was the detection of successful gene delivery events. Until everything worked -gene delivery, gene expression, and detection -we would not know if any part of the transfection protocol was working and therefore were operating in the dark. To improve our chances of success, we cloned promoters and regulatory elements from highlyexpressed S. rosetta genes that lacked introns and replaced their open reading frames with genes, e.g. luciferase or fluorescent proteins, that would report successful transfection. Although genes that confer resistance to a toxic drug can aid in selection for rare drug-resistant transformants, we were discouraged by the variable and unpredictable responses of S. rosetta cultures to drug treatments. Fluorescent protein expression in S. rosetta also presented problems because, as we later learned, green fluorescent protein (GFP) and its derivatives did not fold properly in S. rosetta.
Therefore, we settled on a reporter gene that encodes a small, sensitive, and stable protein, Nanoluc (Hall et al., 2012), to detect expression from transfected plasmids. In other experimental models (Vinayak et al., 2015), the use of nanoluc provided exceptional signal to noise, allowing clear distinction between successful and unsuccessful transfection experiments ( Figure 3C). Moreover, the small size of nanoluc (a 519 bp open reading frame that encodes a ~19 kD protein), helped reduce the overall size of our transgenic constructs. After testing a broad array of transfection parameters with nanoluc constructs, we identified a small combination of nucleofection buffers and pulse programs that yielded luciferase activity. We subsequently optimized around these parameters to develop a robust and reproducible transfection protocol (dx.doi.org/10.17504/protocols.io.h68b9hw).
The establishment of this protocol almost immediately transformed the study of choanoflagellates. The expression of fluorescent markers brought the cellular architecture of S. rosetta into living color in a way not possible previously ( Figure 3D). The ability to express and perform live imaging of fluorescently tagged proteins also helped to validate the mapping of the jumble and couscous glycosyltransferases (Wetzel et al., 2018), revealed that septins localize to the basal pole of S. rosetta cells , and revealed the cytoskeletal dynamics that accompany the transition from flagellated swimming cells to crawling amoeboid cells upon confinement (Brunet et al., 2021). In addition, this protocol served as the foundation for transfection of the choanoflagellate Monosiga brevicollis (Woznica et al., 2021). Finally, and most importantly for the development of S. rosetta as an experimental model, it opened the possibility for delivering the machinery for genome editing. CRISPR/Cas9-mediated genome editing has become a robust method for performing reverse genetics in established and emerging model systems. An easily designed synthetic guide RNA (gRNA) associates with the Cas9 endonuclease, and together they drive cleavage at specific targeted sequences in the genome (Jinek et al., 2013(Jinek et al., , 2012Cong et al., 2013). After the Cas9 endonuclease introduces double-stranded breaks in the genome, the incorporation of mutations ultimately relies on errors introduced during DNA repair (Rouet et al., 1994;Choulika et al., 1995;Bibikova et al., 2001;Jinek et al., 2013;Cong et al., 2013). The nature of those errors depends on the type of DNA repair mechanism implemented by an organism -either homology-directed repair (HDR) in which repair of the double stranded break incorporates mutations from a DNA template that shares homology with the region surrounding the cut site or non-homologous end-joining (NHEJ) in which repair of the double stranded break can lead to untemplated insertions or deletions as the DNA ends rejoin. Thus, a snag for implementing Cas9-mediated genome editing in emerging model systems can be the paucity of information on the DNA repair mechanisms, which can widely vary between organisms and cell types (Yeh et al., 2019). For example, S. rosetta preferentially incorporates mutations from HDR templates at a frequency ten-fold higher than untemplated insertions and deletions . In contrast, M. brevicollis seems to prefer the incorporation of untemplated mutations, as evidenced by the incorporation of insertions and deletions in a targeted gene despite the codelivery of HDR templates (Woznica et al., 2021).
To establish genome editing in S. rosetta, we first focused on converting the ribosomal protein gene rpl36a to rpl36a P56Q , a cycloheximide resistance allele (Kawai et al., 1992;Dehoux et al., 1993;Kondo et al., 1995;Kim et al., 1998;Stevens et al., 2001). This relatively simple edit allowed us to select edited cells due to their cycloheximide resistance, thereby simplifying one stage of the optimization process. By iterating around the editing of rpl36a, we identified key parameters that promoted genome editing success, including the need to use least 20 pmol of Cas9 ribonucleoprotein (RNP) and more than 200 nmol of a single-stranded DNA repair template that had 50 bases of homology flanking the desired mutation .
Afterward, we targeted rosetteless, both because a successful edit should yield a predictable phenotype (loss of rosette development) and, more broadly, to demonstrate the utility of genome editing for investigating gene function in S. rosetta. The original rosetteless allele was likely a hypomorph whose phenotype manifested through aberrant splicing from a transition mutation in a 5' splice site. By introducing an independent knockout of rosetteless by genome editing, we corroborated the necessity of rosetteless for rosette development ( Figure  3E-H). Thus, genome editing in combination with mating crosses are essential tools for the verification of mutations identified in forward genetic screens. Most importantly, genome editing now provides a method to directly test hypotheses about gene function in S. rosetta.
The major barrier that remains for genome engineering in S. rosetta is the accurate and precise introduction of large DNA fragments. Even with small insertions (~20 bases), the efficiency of genome editing (~1%) requires screening through hundreds of clonal strains to find one that bears a genome edit. Engineering larger insertions, which will likely integrate into the genome with even lower frequencies, will require modifications to improve gene delivery, to increase integration frequencies, and/or to select for stable transgenes. A puromycin resistance gene has already been used for the selection of transgenic cassettes (Wetzel et al., 2018;Brunet et al., 2021;Woznica et al., 2021), but puromycin also affects the feeder bacteria that support choanoflagellate growth, leading to variable selection conditions. The development of more selectable markers that confer resistance to drugs that selectively target eukaryotes would help advance genetics in choanoflagellates and other heterotrophic organisms that require bacterial feeding.

The door is now open to study the molecular biology of S. rosetta and other relatives of animals
The lessons learned from S. rosetta have aided efforts to establish genetics in other emerging microeukaryotic models. Of particular note is the recent development of molecular methods for M. brevicollis (Woznica et al., 2021). The establishment of gene delivery and genome editing in M. brevicollis helped to uncover potential ancestral functions of the innate immunity gene STING. Although the exact protocols from S. rosetta were not readily exported to M. brevicollis, variations on the general principles used during the development of S. rosetta transfection facilitated the construction of transgenic lines and the generation of a STING knockout with genome editing. This work clearly demonstrates the maturation of choanoflagellates as model systems to investigate core process conserved in both choanoflagellates and animals; the Woznica et al., 2021 study drew on comparative genomics (Richter et al., 2018), reported the discovery of a bacterial pathogen of M. brevicollis, and developed methods for molecular genetics to show that STING likely functioned as an innate immunity gene in the last common ancestor of choanoflagellates and animals. Similar advances in the development of genetic methods in other protistan relatives of choanoflagellates and animals (Suga and Ruiz-Trillo, 2013;Parra-Acero et al., 2018;Li et al., 2018;Faktorová et al., 2020;Kożyczkowska et al., 2021) now provide a menagerie of microeukaryotes to reconstruct how evolutionary changes prior to the emergence of animals laid the foundations for complex, multicellular development.
We hope that this review serves as an invitation for the 'choano curious' to pursue their research in these fascinating organisms. Rosette development is just one dimension of the rich biology that is now amenable to molecular investigation in S. rosetta; others include cell differentiation, interspecies interactions, and evolutionary cell biology. As we move forward with a new era of molecular genetics in choanoflagellates, we highlight the important roles that classical methods from biochemistry, cytology, and genetics played in developing modern tools for transforming S. rosetta, editing its genome, and illuminating its cell biology (Supplementary  Table).  (Leys and Hill, 2012) showing the collar complex: the apical flagellum (fl) surrounded by microvilli (mv). Also indicated are the nucleus (nu) and food vacuole (fv).  The dynamic life history of S. rosetta includes a multicellular "rosette" stage (A) and a single celled "slow swimmer" stage (B) The phalloidin-stained microvilli are highlighted in magenta, microtubules in white, and the extracellular matrix protein Rosetteless in green. (C) The transition from the slow swimmer stage to the development of the multicellular rosette proceeds through multiple rounds of incomplete cytokinesis (Fairclough et al., 2010). Thus, every rosette is clonal. Rosette development can be induced by secreted cues from Algoriphagus bacteria (Alegado et al., 2012;Woznica et al., 2016) and requires the activity of the C-type lectin protein Rosetteless (green; Levin et al., 2014), that contributes to the extracellular matrix at the basal poles of cells in rosettes. (D) The Rosetteless mutant was uncovered, along with rosettedefective mutants Jumble and Couscous (previously called Branched), as part of a forward genetic screen. Wild type cells typically exist as single cells or simple chains in the absence of inducing factors produced by Algoriphagus; whereas, the presence of Algoriphagus cues induces the development of compact and robust rosettes (Alegado et al., 2012;Levin et al., 2014). In contrast, Rosetteless mutants never develop into rosettes, even in the presence of Algoriphagus conditioned medium (Levin et al., 2014). Jumble and Couscous (previously called Branched in Levin et al., 2014) mutant cells never develop into rosettes, and instead aggregate into large amorphous clumps of cells (Wetzel et al., 2018). (E) The 12-exon rosetteless gene encodes a protein with an N-terminal signal sequence (SS), two C-type lectin domains (CTL1 and 2) and a pair of repeats (RP1 and 2) near the C-terminus. The mutant allele of rosetteless (rtls tl1 ) in the original Rosetteless mutant contained a GT to GC transition at the splice junction between exon 7 and intron 7, which disrupted splicing and protein expression. All scale bars are 5 µm. (Panels A and E adapted from Booth and King, 2020, Panel B adapted from Booth et al., 2018, and Panel D from Levin et al., 2014   Also labeled with pseudocolor: nucleus (teal) and representative bacteria (blue). (B) A step-wise procedure to deliver plasmids and other macromolecules into S. rosetta . Cells are prepared for gene delivery by washing away feeder bacteria-through centrifugationfrom S. rosetta that has been grown to its mid-logarithmic phase. Afterwards, S. rosetta is primed for gene delivery by removing a proteinaceous ECM with a buffer comprised of a chaotropic cation (Lithium), a gentle chelator (Citrate), a reducing agent (Cysteine), and a gentle protease (Papain). The primed cells and cargoes (expression plasmids or gene editing materials) are then combined in a nucleofection or electroporation buffer. After applying an electrical pulse to deliver cargoes inside of S. rosetta, a rest buffer, which has low ionic strength and high osmolarity, is added to cells to promote the closure of holes that persist in membranes long after the application of the electrical pulse. Finally, cells are transferred to growth media to recover before being assayed for gene activity or cultured further to establish transgenic strains. An optimized protocol for transfection was developed using a nucleofector and proprietary buffers  and https://dx.doi.org/10.17504/protocols.io.bx58pq9w). (C) More recently we have established a transformation protocol using an inexpensive, square-pulse electroporator (https://dx.doi.org/10.17504/protocols.io.bx5vpq66), which can be used to deliver plasmids for exogenous gene expression in S. rosetta. By electroporating cells with an expression plasmid pEFL-nanoluc (Addgene NK606), containing the EFL promoter from S. rosetta fused in-frame to the nanoluc luciferase gene and varying the field strength (kV/cm), we found that voltages between 1 -1.5 kV/cm transfected cells most effectively and yielded the highest nanoluc activities. Each dot represents one of three independent transfections, and the grey bar indicates the average of those experiments. Because the nanoluc activity in one electroporation reaction uses ten-fold more material to achieve a similar level of activity as one nucleofection reaction, we estimate that these electroporation conditions transfect cells with tenfold lower efficiency than nucleofection. (D) Transfection of S. rosetta with plasmids expressing fluorescent proteins can be used to highlight cellular architecture in live cells. The collar and cell membranes here are visualized through the expression of mCherry fused to an S. rosetta geranyl-geranylation sequence (magenta; Addgene NK644) and the cell body, illuminated by the expression of cytoplasmic mTFP (green; Addgene NK591). (E, F) Genome editing of the rosetteless gene recapitulates the phenotype of the previously isolated rosetteless mutant. (E) Genotypes of S. rosetta strains show two distinct alleles in rosetteless that were either isolated from a forward genetic screen (rtls tl ; Levin et al., 2014) or engineered through CRISPR-Cas9 mediated genome editing (rtls PTS1 rpl36a P56Q ; Booth and King, 2020). The rtls tl1 allele is a T to C transition in the 5'-splice site of intron 7 that abrogates the expression of rosetteless through aberrant splicing (also see Fig. 2). The rtls PTS1 rpl36a P56Q strain was engineered to bear a cycloheximide resistance allele (rpl36 P56Q ) and a rosetteless allele with a palindromic premature termination sequence (PTS: TTTATTTAATTAAATAAA) inserted into the fourth exon that stops the transcription and translation of rosetteless products. (F) Independent mutations show the necessity of rosetteless for multicellular development in S. rosetta. In the absence of inducing cues from Algoriphagus, all strains (F'-H') grew as chains (stars) or single cells. Upon the addition of inducing cues, only the wild-type (F") strain formed rosettes (arrowheads) while strains with rtls tl1 (E") or rtls PTS1 rpl36 P56Q (F") alleles failed to form rosettes. (Panel A courtesy of Kent McDonald and adapted from Booth et al., 2018; Panels E-H adapted from Booth and King, 2020)