Genetic transformation of Spizellomyces punctatus, a resource for studying chytrid biology and evolutionary cell biology.

Chytrids are early-diverging fungi that share features with animals that have been lost in most other fungi. They hold promise as a system to study fungal and animal evolution, but we lack genetic tools for hypothesis testing. Here, we generated transgenic lines of the chytrid Spizellomyces punctatus, and used fluorescence microscopy to explore chytrid cell biology and development during its life cycle. We show that the chytrid undergoes multiple rounds of synchronous nuclear division, followed by cellularization, to create and release many daughter 'zoospores'. The zoospores, akin to animal cells, crawl using actin-mediated cell migration. After forming a cell wall, polymerized actin reorganizes into fungal-like cortical patches and cables that extend into hyphal-like structures. Actin perinuclear shells form each cell cycle and polygonal territories emerge during cellularization. This work makes Spizellomyces a genetically tractable model for comparative cell biology and understanding the evolution of fungi and early eukaryotes.

Introduction

Zoosporic fungi, commonly referred to as ‘chytrids’, span some of the deepest fungal Phyla and comprise much of the undescribed environmental fungal DNA diversity in aquatic ecosystems (James et al., 2006; Richards et al., 2012; Powell and Letcher, 2014; Grossart et al., 2016). Many chytrids are saprophytes or parasites of photosynthetic organisms and actively shuttle carbon to higher trophic levels (Kagami et al., 2014; Grossart et al., 2016). Other chytrids are animal parasites, including the Batrachochytrium genus that includes the frog-killing B. dendrobatidis (Longcore et al., 1999) and salamander-killing B. salamandrivorans that are devastating global amphibian populations (Martel et al., 2013).

Chytrids are unique in that they have retained ancestral cellular features, shared by animal cells and amoebae, while also having fungal features. For example, chytrids begin their life as motile zoospores that lack a cell wall, swim with a single posterior cilium nucleated from a centriole, and crawl across surfaces (Fuller, 1976; Sparrow, 1960; Deacon and Saxena, 1997; Held, 1975; Fritz-Laylin et al., 2017b). Later life cycle stages exhibit fungal characteristics including the formation of chitinous cell walls, the growth of hyphal-like structures, and the development of a sporangium (sporangiogenesis); see Figure 1. Chytrid zoosporogenesis involves multiple rounds of mitosis without cytokinesis to create a multi-nuclear coenocyte, followed by cellularization to form zoospores with a single nucleus. The formation of a multinuclear compartment followed by cellularization is reminiscent of development in flies (e.g. Drosophila), amoeba (e.g. Physarum), and protozoa (e.g. Plasmodium). Although there are important differences from fly embryos (particularly the need for the chytrid sporangium to extract nutrients from the environment and coordinate growth with the cell cycle), determining the mechanisms controlling chytrid cellularization provides a comparative framework for understanding cellularization in animals and other eukaryotic lineages.

Figure 1.

Life cycle of the chytrid Spizellomyces punctatus.

Timeline and events as measured in this work. The chytrid produces globular zoospores (3–5 µm) that swim with a motile cilium (20–24 µm). (A) The uninucleate zoospore (nucleus in blue) has a cilium associated with a basal body. Swimming zoospores can also crawl on surfaces using amoeboid-like motion (polymerized actin in red). (B) The start of encystment (before 1 hr) occurs when the cilium retracts by a lash-around mechanism, followed by formation of cell wall (Koch, 1968). (C) The cyst then germinates and forms a single germ tube (at 1–3 hr) that later expands and branches into a rhizoidal system. The nucleus remains in the cyst during germ tube expansion as the cyst develops into a single reproductive structure called the sporangium. The first mitotic event (at 8–12 hr) usually correlates with the ramification of rhizoids from the germ tube. (D) Mitosis in the sporangium is coordinated with growth, as nuclei replicate and divide in a shared compartment. There can be a total of five to eight synchronous mitotic cycles as each sporangium develops a branched rhizoid system with subsporangial swelling in the main rhizoid. (E) Mitosis halts and zoospore formation begins in the sporangium. Ciliogenesis likely occurs before cellularization as in other chytrids (Renaud and Swift, 1964). (F) The nuclei cellularize and develop into zoospores while the sporangium develops discharge papillae. Once cellularization is complete and environmental conditions are favorable, the zoospores will escape the sporangium through the discharge papillae (at 20–30 hr). Diagram not drawn to scale. Times are relative to the start of microscopy after zoospore harvest.

The major bottleneck to studying chytrids in molecular detail has been the lack of a model organism with tools for genetic transformation. Here, we describe the successful adaptation of Agrobacterium-mediated transformation to generate reliable and stable genetic transformation of the soil chytrid Spizellomyces punctatus. We expressed fluorescent proteins fused to histone and actin-binding proteins to characterize the development and cell biology of Spizellomyces throughout its life cycle using live-cell imaging. Below, we will show that Spizellomyces is a well-suited model system for uncovering molecular mechanisms of cell cycle regulation, cell motility, and development because it is fast-growing, displays both crawling and swimming motility, and possesses a characteristic chytrid developmental life cycle. These tools allow, for the first time, direct molecular probing to test new hypotheses about the evolution and regulation of the cell cycle (Medina et al., 2016), cell motility (Fritz-Laylin et al., 2017b), and development in chytrid fungi.

Results

Developing tools for genetic transformation

The plant pathogen Agrobacterium tumefaciens normally induces plant tumors by injecting and integrating a segment of transfer DNA (T-DNA) from a tumor-inducing plasmid (Ti-plasmid) into the plant genome. Researchers have exploited this feature to integrate foreign genes in plants by cloning them into the T-DNA region of the Ti-plasmid, inducing virulence genes for processing/transport of T-DNA, and co-culturing induced Agrobacterium with the desired plant strain. Because Agrobacterium-mediated transformation has been adapted for transformation of diverse animals and fungi (Bundock et al., 1995; Kunik et al., 2001; Covert et al., 2001; Ianiri et al., 2017; Vieira and Camilo, 2011), we chose to use this system in Spizellomyces punctatus. To this end, we modified an Agrobacterium plasmid to integrate and express a selectable marker (e.g. drug resistance) in Spizellomyces.

To determine a suitable selection marker for Spizellomyces, we tested the effects of drugs on the growth of the chytrid on agar plates. We spread zoospores on plates with various concentrations of drugs and assessed the cultures for cell growth, colony formation, and zoospore release using light microscopy. Although Geneticin (G418), Puromycin, and Phleomycin D10 (Zeocin) did not inhibit growth up to 800 mg/L, we determined that 200 mg/L Hygromycin and 800 mg/L Nourseothricin (CloNAT) resulted in complete absence of growth after 6 days of incubation at 30°C. All remaining experiments were performed using Hygromycin (200 mg/L).

Next, we identified Spizellomyces promoters that can drive gene expression at sufficient levels to provide resistance to Hygromycin and measurable protein fluorescence. In the absence of a chytrid system to perform these tests, we reasoned that Spizellomyces promoters that express at high levels in yeast (Saccharomyces cerevisiae) would likely also work in chytrids. Therefore, we used an Agrobacterium plasmid (Ianiri et al., 2017) that propagates in yeast to first screen Spizellomyces promoters that successfully express a fusion of Hygromycin resistance (hph) and green fluorescent protein (GFP); see Materials and methods. We confirmed that Spizellomyces HSP70 and H2B promoters resulted in resistance to Hygromycin as well as measurable GFP fluorescence in yeast via flow cytometry (Figure 2—figure supplement 1A) and microscopy. All remaining experiments were performed using the stronger H2B promoter.

Figure 2—figure supplement 1.

Spizellomyces promoters successfully express fluorescent protein and drug-resistance genes.

(A) Fluorescence distribution of yeast strains transformed with plasmids pGI3EM09 (Hsp70pr-hph-GFP) and pGI3EM11 (H2Bpr-hph-GFP) relative to untransformed wild-type (WT). Flow cytometry data were collected on a MacsQuant VYB with a 488 nm excitation laser and FITC emission filter (525/50 nm). Data were collected from two independent transformants. The large width of the fluorescence distribution arises from copy number fluctuations of the 2µ plasmid in Saccharomyces cerevisiae. (B) Typical time scale and transformation efficiency using Agrobacterium-mediated transformation of Spizellomyces. Data is shown for pGI3EM11 (H2Bpr-hph-GFP) and pGI3EM18 (H2Bpr-hph-tdTomato) in the absence and presence of selection (200 mg/L Hygromycin). Ampicillin and tetracycline (50 mg/L) are included to kill any Agrobacterium transferred from the co-culture plate; see Materials and methods. Small red arrows indicate three examples of tiny colonies that appeared on pGI3EM11 plate on Day 4. Photos are shown with inverted contrast to better highlight colonies.

With active promoters and effective selection, we performed Agrobacterium-mediated transformation by co-culturing Spizellomyces zoospores with Agrobacterium carrying H2Bpr-hph-GFP; see Materials and methods. Although Hygromycin-resistant, none of the GFP transformants (Figure 2—figure supplement 1B) exhibited green fluorescence above background. This has been seen in other emerging model systems (i.e. choanoflagellate Salpingoeca Booth et al., 2018) and is likely due to GFP misfolding. When GFP was replaced by tdTomato, we obtained transformants that exhibited both Hygromycin resistance (Figure 2—figure supplement 1B) and cytoplasmic fluorescence (Figure 2—figure supplement 2). Further tests with other fluorescent protein fusions showed that mClover3, mCitrine, and mCerulean3 are functional in Spizellomyces (Figure 2—figure supplement 2). We then designed a construct with greater applicability, in which the selectable marker and fluorescent protein are expressed independently and where the fluorescent protein (tdTomato) is fused in-frame to the C-terminus of a protein of interest (POI). This design exploits the compact and divergent architecture of the Spizellomyces H2A/H2B promoters to express (POI)-tdTomato in an upstream direction (H2B promoter) while expressing hph in a downstream direction (H2A promoter). As a proof of concept and because we were interested in following nuclear dynamics to measure the timing and synchrony of mitotic events (e.g. DNA segregation, see next section), our first protein of interest was histone H2B; see Figure 2A.

Figure 2—figure supplement 2.

Diverse fluorescent proteins are functional in Spizellomyces punctatus.

DIC and fluorescence images were taken with a Deltavision Elite microscope using POL Transmission 32%, exposure time 0.5 s; Filter Transmission 32%, exposure time 0.3 s. TRITC filter set excitation (542/27 nm) and emission (597/45 nm); GFP filter set excitation (475/28 nm) and emission (525/48 nm); YFP filter set excitation (513/17 nm) and emission (548/22 nm); CFP filter set excitation (438/24 nm) and emission (475/24 nm). Paired wild-type and fluorescent protein strains are plotted with the same intensity range.

Figure 2.

Genomic integration of H2B-tdTomato using Agrobacterium-mediated transformation.

(A) Plasmid GI3EM20C takes advantage of the divergent architecture of H2A/H2B to express an H2B-tdTomato fusion in an upstream direction (H2B promoter) while expressing hph in a downstream direction (H2A promoter). (B) Representative images from wild type (left), and transformants expressing cytoplasmic hph-tdTomato (plasmid GI3EM18) (center) and nuclear-localized H2B-tdTomato (right). Top row shows DIC and the middle row shows fluorescence microscopy at 561 nm with overlaid images on the bottom row. For comparable results, all strains are presented at the same intensity levels used for H2B-tdTomato fluorescence image. Scale bar indicates ten microns. Image acquisition conditions: POL: transmittance 32%, exposure 0.15 s; TRITC filter, maximal projection, transmittance 32%, Exposure 0.2 s, 0.3 micrometers slice thickness. (C) Southern blot of four transformants, in which genomic DNA was digested either with XbaI or KpnI and probed using the Hygromycin resistance gene (hph). We used plasmid GI3EM20C as a positive control (+). (D) Four independent transformants were transferred, every two days, in both selective and non-selective medium at 30°C for a total of 27 days (minimum of 23 life cycles or 116–185 mitotic cycles), followed by a challenge on selective medium. These strains were spotted in a twofold dilution series on non-selective and selective (+Hygromycin) plates, and incubated for 2 days at 30°C.

(A) Fluorescence distribution of yeast strains transformed with plasmids pGI3EM09 (Hsp70pr-hph-GFP) and pGI3EM11 (H2Bpr-hph-GFP) relative to untransformed wild-type (WT). Flow cytometry data were collected on a MacsQuant VYB with a 488 nm excitation laser and FITC emission filter (525/50 nm). Data were collected from two independent transformants. The large width of the fluorescence distribution arises from copy number fluctuations of the 2µ plasmid in Saccharomyces cerevisiae. (B) Typical time scale and transformation efficiency using Agrobacterium-mediated transformation of Spizellomyces. Data is shown for pGI3EM11 (H2Bpr-hph-GFP) and pGI3EM18 (H2Bpr-hph-tdTomato) in the absence and presence of selection (200 mg/L Hygromycin). Ampicillin and tetracycline (50 mg/L) are included to kill any Agrobacterium transferred from the co-culture plate; see Materials and methods. Small red arrows indicate three examples of tiny colonies that appeared on pGI3EM11 plate on Day 4. Photos are shown with inverted contrast to better highlight colonies.

DIC and fluorescence images were taken with a Deltavision Elite microscope using POL Transmission 32%, exposure time 0.5 s; Filter Transmission 32%, exposure time 0.3 s. TRITC filter set excitation (542/27 nm) and emission (597/45 nm); GFP filter set excitation (475/28 nm) and emission (525/48 nm); YFP filter set excitation (513/17 nm) and emission (548/22 nm); CFP filter set excitation (438/24 nm) and emission (475/24 nm). Paired wild-type and fluorescent protein strains are plotted with the same intensity range.

(A) Primer locations and amplicon sizes of different pGI3EM20C primer pairs. RB and LB correspond to the ‘right border’ and ‘left border’ of the T-DNA plasmid. These sequences define the T-DNA fragment that is excised and transferred into the host cell by the Agrobacterium virulence machinery. The total size of the T-DNA (LB to RB) is 4280 bp. (B) Gel electrophoresis of different PCR reactions using genomic DNA of untransformed (WT) and four independent Spizellomyces transformants. (+) control used pGI3EM20C plasmid DNA as template.

(A) Diagram of T-DNA integration and the location of PCR/sequencing primers and restriction sites used for inverse PCR (invPCR). We only show primers adjacent to the left border (LB) because they consistently amplified for all transformants, unlike the primers adjacent to the right border. (B) Example of amplification by invPCR of the LB-genome border after EcoRI genomic digestion and ligation for an untransformed strain (WT), four independent transformants and non-template control. (C) Amplification by invPCR of the LB-genome border after HindIII genomic digestion and ligation. T-DNA location for all transformants was confirmed by two independent biological replicates (i.e. independent genomic extractions, ligation and invPCR). (D) T-DNA insertion sites in four independent transformants of Spizellomyces. In strain EM20C-3, invPCR for EcoRI indicated LB is located toward SPPG_02523, while invPCR for HindIII shows same insertion site but with an inverted direction. The divergent invPCR results might represent an insertion of a tandem inverted T-DNA. (E) Three of the four strains (EM20C-2,3,4) have similar tdTomato fluorescence levels as determined by flow cytometry.

Timing of germ tube emergence (), mitosis (), and bursting of the sporangium () (i.e. zoospore release) was determined through brightfield and fluorescence microscopy. Cell cycle period is the interval of time between mitotic events ( - ). (A) Timing of mitotic events and B) Cell cycle period in transformant EM20C-1. (C) Timing of mitotic events and D) Cell cycle period in transformant EM20C-2. Two biological replicates (yellow and purple) per transformant. Number of measured events per replicate are shown in the top panels. For a given strain, the variability between replicates is mostly due to a 1–2 hr shift in germ tube formation and early mitotic events. Both EM20C-1 and EM20C-2 form germ tubes at 1–3 hr, the first mitosis occurs between 8 and 12 hr, the cell cycle periods are ∼150 min before shortening in late development. EM20C-2 tends to have a shorter developmental cycle than EM20C-1 (bursting at 20–25 hr versus 25–30 hr) through a combination of shorter cell cycle periods in late development and fewer cycles before bursting. Not all germ tubes were visible and some sporangia had fewer mitotic cycles before bursting; thus, , , , have smaller sample sizes. Asterix above each plot reflects the p-value of a two-sample Kolmogorov-Smirnov test of equality of distributions between replicates (*, p<0.05; **, p<0.01; ***, p<0.001).

The left panel shows DIC microscopy, while right panel shows 561 nm fluorescence microscopy of EM20C-1. Time-stamp is hr:min. Scale bar indicates ten microns.

Mitotic cycles are fast and highly synchronous during sporangiogenesis

Chytrid transformation with H2B-tdTomato resulted in bright nuclear localization of fluorescence when compared to cytoplasmic tdTomato (Figure 2B). The presence of the T-DNA (total size of 4280 bp) in the transformants was confirmed using PCR for hph and H2B-tdTomato (Figure 2—figure supplement 3) and through Southern blot analysis of hph (Figure 2C). The results were consistent with random, single T-DNA genomic integration events in each transformant. To determine the location of the T-DNA integrations, we identified the genomic region adjacent to the left border (LB) of the T-DNA by inverse PCR (Figure 2—figure supplement 4). In three of the four transformants (EM20C-1, 2, 3), the T-DNA LB was located within 200 bp upstream of the transcription start site (TSS) of a gene (SPPG_04375 an M48 peptidase, SPPG_03425 an adenine nucleotide hydrolase, and SPPG_02523 a PHO-like cyclin, respectively). For strain EM20C-3, two different DNA:genome junctions were detected in the 5’UTR of the gene SPPG_02523, suggesting an irregular T-DNA insertion. Last, for strain EM20C-4, the LB T-DNA was inserted 844 bp from the closest TSS (SPPG_08788 a hypothetical protein). As observed in Arabidopsis, we might expect variation in gene expression based on the genomic locus of integration of the T-DNA fragment. To quantify this variation, we measured H2B-tdTomato expression in our EM20C transformants using flow cytometry (Figure 2—figure supplement 4). The data show that strains EM20C-2, 3, 4 showed similar and unimodal levels of tdTomato fluorescence at the population level, despite the different sites of genomic integration. The exception is EM20C-1, which exhibits bimodal gene expression: the top mode is identical to the other transformants, but the bottom mode is half the intensity. Last, we established that the transformants had transgenerational stability by passaging them in non-selective medium for several weeks, followed by a challenge in selective medium (Figure 2D).

Figure 2—figure supplement 3.

PCR validation of H2B-tdTomato transformants.

(A) Primer locations and amplicon sizes of different pGI3EM20C primer pairs. RB and LB correspond to the ‘right border’ and ‘left border’ of the T-DNA plasmid. These sequences define the T-DNA fragment that is excised and transferred into the host cell by the Agrobacterium virulence machinery. The total size of the T-DNA (LB to RB) is 4280 bp. (B) Gel electrophoresis of different PCR reactions using genomic DNA of untransformed (WT) and four independent Spizellomyces transformants. (+) control used pGI3EM20C plasmid DNA as template.

Figure 2—figure supplement 4.

Mapping T-DNA genomic insertion sites with inverse PCR.

(A) Diagram of T-DNA integration and the location of PCR/sequencing primers and restriction sites used for inverse PCR (invPCR). We only show primers adjacent to the left border (LB) because they consistently amplified for all transformants, unlike the primers adjacent to the right border. (B) Example of amplification by invPCR of the LB-genome border after EcoRI genomic digestion and ligation for an untransformed strain (WT), four independent transformants and non-template control. (C) Amplification by invPCR of the LB-genome border after HindIII genomic digestion and ligation. T-DNA location for all transformants was confirmed by two independent biological replicates (i.e. independent genomic extractions, ligation and invPCR). (D) T-DNA insertion sites in four independent transformants of Spizellomyces. In strain EM20C-3, invPCR for EcoRI indicated LB is located toward SPPG_02523, while invPCR for HindIII shows same insertion site but with an inverted direction. The divergent invPCR results might represent an insertion of a tandem inverted T-DNA. (E) Three of the four strains (EM20C-2,3,4) have similar tdTomato fluorescence levels as determined by flow cytometry.

To quantify the timing and synchrony of the Spizellomyces cell cycle, we used live cell epi-fluorescence imaging of H2B-tdTomato strains EM20C-1 and EM20C-2 at 2 min intervals (Figure 2—figure supplement 5). Our results show that zoospores have a single nucleus, they retract their flagellum and encyst in less than 1 hr, the germ tube emerges at ∼1−3 hr, the first mitotic event (i.e. one nucleus to two nuclei) occurs at ∼8−12 hr, and sporangia develop and undergo 5–8 mitotic cycles in less than 30 hr before completing their life cycles and releasing 32–256 zoospores; see Figure 2—video 1. To measure all nuclei within a sporangium with better temporal and z-resolution, we followed nuclear dynamics of EM20C-1 at 1 min time intervals using live-cell confocal microscopy of a H2B-tdTomato strain (Figure 3—video 1). We measured the number of nuclei over time per sporangium (Figure 3A) to estimate the synchrony of nuclear division waves and the period of time between waves of nuclear division. Wave time () is the time for a wave of synchronous nuclear divisions to propagate across the sporangium. The cell cycle period () is the interval of time between nuclear division waves. We found that the average cell cycle period was ∼150 min and that each wave of nuclear division was completed within 1 min (Figure 3B). In addition, by following the compaction and localization of H2B-tdTomato we show that all measurable mitotic events occurred within 5 min, or less than 3.3% of the cell cycle period (Figure 3B and C). Altogether, these results show that Spizellomyces is mitotically inactive in its early life cycle (zoospore and germination stages) but, once committed, the cell cycle is fast and nuclear divisions within each sporangium are highly synchronous; see Table 1.

Figure 2—figure supplement 5.

Comparison of development and cell cycle timing between EM20C-1 and EM20C-2.

Timing of germ tube emergence (), mitosis (), and bursting of the sporangium () (i.e. zoospore release) was determined through brightfield and fluorescence microscopy. Cell cycle period is the interval of time between mitotic events ( - ). (A) Timing of mitotic events and B) Cell cycle period in transformant EM20C-1. (C) Timing of mitotic events and D) Cell cycle period in transformant EM20C-2. Two biological replicates (yellow and purple) per transformant. Number of measured events per replicate are shown in the top panels. For a given strain, the variability between replicates is mostly due to a 1–2 hr shift in germ tube formation and early mitotic events. Both EM20C-1 and EM20C-2 form germ tubes at 1–3 hr, the first mitosis occurs between 8 and 12 hr, the cell cycle periods are ∼150 min before shortening in late development. EM20C-2 tends to have a shorter developmental cycle than EM20C-1 (bursting at 20–25 hr versus 25–30 hr) through a combination of shorter cell cycle periods in late development and fewer cycles before bursting. Not all germ tubes were visible and some sporangia had fewer mitotic cycles before bursting; thus, , , , have smaller sample sizes. Asterix above each plot reflects the p-value of a two-sample Kolmogorov-Smirnov test of equality of distributions between replicates (*, p<0.05; **, p<0.01; ***, p<0.001).

Figure 2—video 1.

Time-lapse microscopy in developing H2B-tdTomato sporangia taken over the course of 32 hr with images captured every 2 min.

The left panel shows DIC microscopy, while right panel shows 561 nm fluorescence microscopy of EM20C-1. Time-stamp is hr:min. Scale bar indicates ten microns.

Figure 3—video 1.

Time-lapse microscopy of nuclear divisions in developing H2B-tdTomato sporangia taken over the course of 24 hr with images captured every 1 min.

Time-stamp is hr:min. Scale bar indicates five microns.

Figure 3.

H2B-tdTomato reveals the timing and synchrony of mitotic events during sporangiogenesis.

(A) Number of nuclei as a function of time during the development of a sporangium, along with H2B-tdTomato fluorescence images from select time points. Each colored line corresponds to a different sporangium. The cell cycle period () is the interval of time between waves of nuclear division (i.e. metaphase to anaphase transition). The wave time () is the interval of time for a synchronous wave of nuclear division to sweep across the sporangium. (B) Distribution of cell cycle period (), wave time () and duration of mitosis () across multiple cell cycles. (C) Timing of mitotic events. H2B-tdTomato permits observation of (1) leakage from the nucleus likely due to fenestration of nuclear envelope by the mitotic spindle (Heath, 1980; Fuller, 1976), followed by chromosome condensation, and (2) chromosome separation during anaphase. Dotted line highlights the cell wall of the sporangium. This particular example shows a mitosis duration of 4 min (=time from nuclear leakage to anaphase). Time in hr:min. Scale 2.5 micrometers. Distributions are from one time-lapse movie of EM20C-1 (6 cells).

Data was used to create Figure 3A and analyzed to create Figure 3B.

Time-stamp is hr:min. Scale bar indicates five microns.

Table 1.

Comparison of nuclear division synchrony for different coenocytic organisms.

Wave time () is defined as the average interval of time for a wave of synchronized nuclear divisions to propagate across the coenocytic nuclei. The nuclear division period () is the average interval of time between waves. Organisms are listed from highest to lowest synchrony index.

Coenocytic organism	Length scale	Wave time	Wave speed	Nuclear division	Synchrony index	References
	(µm)	Δt (min)	(µm min-1)	Period τ (min)	100%⋅(1-Δ⁢t/τ)
Physarum polycephalum (amoeba)	1000	2	500	840	99.8%	Halvorsrud et al., 1995
Spizellomyces punctatus (fungi)	5–10	1	5–10	150	99.3%	This work
Creolimax fragrantissima (holozoa)		20		300	93.3%	Suga and Ruiz-Trillo, 2013
Drosophila melanogaster (metazoa)	500	1.5	360	10	85.5%	Deneke et al., 2016
Aspergillus nidulans (fungi)	700	20	35	60	66.7%	Clutterbuck, 1970; Momany and Taylor, 2000

Number of nuclei per cell as a function of time (min).

Actin polymerization drives zoospore motility

Like their pre-fungal ancestors, chytrids swim with a motile cilium. Some chytrids can also crawl across and between solid substrates, much like amoeba and animal immune cells (Fuller, 1976; Sparrow, 1960; Whisler et al., 1975; Couch, 1945; Deacon and Saxena, 1997; Held, 1973; Dorward and Powell, 1983; Fritz-Laylin et al., 2017b). Eukaryotes employ multiple strategies to crawl (filopodia, pseudopodia and blebs) that depend on distinct molecular mechanisms (Fritz-Laylin et al., 2017a). One form of crawling, the pseudopod-based -motility, relies on the expansion of branched-actin filament networks that are assembled by the Arp2/3 complex and allow cells to navigate complex environments at speeds exceeding 20 μm/min. The activators of branched-actin assembly WASP and SCAR/WAVE have been described as a molecular signature of the capacity for -motility (Fritz-Laylin et al., 2017b). Spizellomyces has homologs of WASP (SPPG_00537), SCAR/WAVE (SPPG_02302), and its zoospores are proficient crawlers.

To test whether Spizellomyces zoospores crawl using -motility, we expressed a LifeAct-tdTomato fusion. LifeAct is a 17 amino acid peptide that binds specifically to polymerized actin in a wide variety of cell types, such as actin patches and cables in yeast and actin-filled protrusions of crawling animal cells (Riedl et al., 2008; Belin et al., 2014). To confirm that our LifeAct-tdTomato fusion binds specifically to polymerized actin in Spizellomyces, we first fixed and stained actin in zoospores (Figure 4) and sporangia (Figure 5) with fluorescent phalloidin. We then compared the fluorescent images of cells expressing LifeAct-tdTomato and those expressing hph-tdTomato (negative control) relative to phalloidin-staining.

Figure 4.

Localization of LifeAct-tdTomato in zoospores highlights cortical and pseudopod actin networks.

(A) Zoospores from wild type (left), and transformants expressing hph-tdTomato (center) and LifeAct-tdTomato (right) were fixed and stained with fluorescent phalloidin (green). Top row shows DIC and second row shows DNA stain (DAPI). The bottom row shows the phalloidin stain and 561 nm images overlaid. Scale bar indicates two microns. (B and C) Line scan of fixed and stained hph-tdTomato (B) and LifeAct-tdTomato fusion (C). The plot shows line scans of normalized fluorescence intensity of the respective fusion protein (magenta) and fluorescent phalloidin (green). The location for generating the line scans is shown by a yellow dotted line in the image above each plot. Scale bars indicate 1 µm. (D) Stills taken at 20-s intervals from timelapse microscopy of crawling zoospores from the indicated strains at the given timepoints. Images were taken using DIC microscopy (top) and 561 nm fluorescence microscopy (middle), also shown with images merged (bottom). Scale bar indicates 5 µm. E+F) Line scan of fixed and stained hph-tdTomato (E) and LifeAct-tdTomato fusion (F). The plot shows line scans of normalized fluorescence intensity of the respective fusion protein (magenta) and fluorescent phalloidin (green). The location for generating the line scans is shown by a yellow dotted line in the image above each plot. Scale bars indicate 2 µm.

The left panel shows DIC microscopy, the center panel shows 561 nm fluorescence microscopy, and the right panel shows a merge of fluorescence and DIC microscopy. Time stamp is min:s. Scale bar indicates 10 µm.

Figure 5.

Localization of LifeAct-tdTomato in sporangia highlights actin patches, cables, and perinuclear shells.

(A) Sporangia from wild type (left), and transformants expressing hph-tdTomato (center) and LifeAct-tdTomato (right) were fixed and stained with fluorescent phalloidin (green). The third row shows DNA stain (DAPI). The bottom row shows the phalloidin stain and 561 nm images overlaid. Scale bar indicates 5 µm. Arrows point to examples of actin patches present in sporangia and rhizoids (‘p’), cables (‘c’) and perinuclear actin shells (‘ns’). (B) Selected stills taken from timelapse microscopy of developing sporangia from the LifeAct-tdTomato transformant strain at times indicated (hr: min). Formation of polygonal territories precedes cellularization. Images taken using DIC (top) and 561 nm (middle), also shown with images merged (bottom). Scale bar indicates ten microns. (C) Multiple planes of a single sporangium show how polygonal territories formed during later stages of cellularization encompass the entire cytoplasm.

The left panel shows DIC microscopy, the center panel shows 561 nm fluorescence microscopy, and the right shows a merge of fluorescence and DIC microscopy. Time stamp is hr:min. Scale bar indicates 10 µm.

In contrast to the relatively homogeneous distribution of fluorescence in hph-tdTomato zoospores, both fixed (Figure 4A–C) and living zoospores (Figure 4D–E, and Figure 4—video 1) of the LifeAct-tdTomato strain showed a thin layer of fluorescence at the cell cortex, and high levels of fluorescence in the pseudopods at the leading edge. Because the LifeAct fusion localization nearly perfectly correlated with phalloidin intensities in fixed cells, but the hph fusion did not, we presume that this fluorescence represents polymerized actin. The minor deviations between Lifeact and phalloidin can be explained by intrinsic biases of the probes. All live-cell probes of filamentous actin (e.g. Lifeact, F-tractin, Utrophin actin-binding domain) have different biases in their patterns of F-actin localization and dynamics (Belin et al., 2014). None of them can fully recapitulate the patterns seen with phalloidin, but they do provide new insights into the live cell dynamics of actin. The distribution and dynamics of actin that we see in zoospores is in agreement with -motility and actin localization observed for fixed cells during zoospore crawling of Neocallimastix (Li and Heath, 1994) and Batrachochytrium (Fritz-Laylin et al., 2017b).

Figure 4—video 1.

Time-lapse microscopy of crawling wild type, cytoplasmic tdTomato and LifeAct-tdTomato zoospores taken over the course of 2 min with images captured every second, playback in real time.

The left panel shows DIC microscopy, the center panel shows 561 nm fluorescence microscopy, and the right panel shows a merge of fluorescence and DIC microscopy. Time stamp is min:s. Scale bar indicates 10 µm.

Actin polymerization during sporangiogenesis

Once the chytrid zoospore encysts, it builds a sporangium by growing both radially and in polarized fashion during germ tube extension and rhizoid formation (Figure 1). During early sporangiogenesis, the nuclei are very dynamic while replicating and dividing but then slow down in late sporangiogenesis, presumably during cellularization and zoospore formation (Figure 2—video1). Actin has been reported to play fundamental roles during cellularization in another chytrid, Allomyces macrogynus (Lowry et al., 1998; Lowry et al., 2004). Thus, we expect polymerized actin to play a role in the nuclear dynamics and cellularization during sporangiogenesis in Spizellomyces.

Our experiments revealed an actin cytoskeleton distributed primarily between cortical patches and linear structures that resemble the actin cables of other fungal species (Figure 5A). Each of these structures was visible in both fixed phalloidin stained cells and living cells expressing the LifeAct-tdTomato gene. We also detected transient perinuclear polymerized actin shells with a period similar to mitotic events (Figure 5B and Figure 5—video 1), which suggests a role for polymerized actin with nuclei during the cell cycle. In late sporangiogenesis, polymerized actin delineated polygonal zoospore territories (Figure 5B–C). This is reminiscent of epithelial cellularization seen during Drosophila embryogenesis when syncytial nuclei are encapsulated by cell membrane.

Figure 5—video 1.

Time-lapse microscopy of developing LifeAct-tdTomato sporangia taken over the course of 19 hr with images captured every 5 min.

The left panel shows DIC microscopy, the center panel shows 561 nm fluorescence microscopy, and the right shows a merge of fluorescence and DIC microscopy. Time stamp is hr:min. Scale bar indicates 10 µm.

Discussion

Here, we report stable and robust genetic transformation of the chytrid Spizellomyces punctatus. We identified and tested native Spizellomyces promoters, including a divergent H2A/H2B promoter that can simultaneously express Hygromycin resistance (hph) and a gene of interest throughout the chytrid life cycle. This design was used to express fluorescent proteins and resistance genes in Dikaryotic marine fungi (data not shown), which suggests they should be useful for transforming other chytrids, such as the pathogenic Batrachochytrium. Agrobacterium was previously used to transform the aquatic chytrid Blastocladiella emersonii (Vieira and Camilo, 2011). Unfortunately, genetic transformation of Blastocladiella displayed a narrow window of gene expression and could only be detected at the zoosporic stage. The advances described in this study should be useful for random and possibly targeted gene integration in other chytrids. For example, Agrobacterium T-DNA is often randomly integrated into the host chromosome as a single copy and, if inserted within a gene, it will disrupt gene function and act as a mutagen (Gelvin, 2017). Random insertional mutagenesis by Agrobacterium has been exploited for forward genetics in plants and fungi (Alonso et al., 2003; Michielse et al., 2005; Idnurm et al., 2017). In Fungi, Agrobacterium-mediated transformation has also been used for targeted gene knock-outs by including 1–1.5 kb long, flanking region of homology in the T-DNA to the desired target gene (Frandsen et al., 2012). The feasibility of this approach in a chytrid relies on the rate of homologous recombination, which is in general high in the majority of Fungi.

As a first step to characterize the chytrid cell cycle, we used an H2B-tdTomato fusion and live-cell microscopy to measure the timing of nuclear division. The Spizellomyces developmental program allocates a narrow window (∼5 min) of time to the mitotic process during each cell cycle with a highly synchronous wave of nuclear divisions. The level of synchrony is similar to plasmodial nuclei in the amoeba Physarum polycephalum or syncytial nuclei during Drosophila development, where all nuclei divide within 2 min (Deneke et al., 2016) (see Table 1). These mitotic dynamics make Spizellomyces an interesting comparative model for exploring the physical and molecular determinants of cell division synchrony and its evolution in fungi, animals, and amoeba.

By visualizing actin dynamics throughout the chytrid life cycle, we found that Spizellomyces zoospores assemble thick actin cortices and build actin-filled protrusions during motility, similar to other chytrid species (Fritz-Laylin et al., 2017b), animal cells, and amoebae. Once the zoospores encyst, there is a drastic shift in actin cytoskeleton organization, in which the cortical shell of polymerized actin was replaced by dynamic puncta (actin patches), some of which are associated with actin cables that often extended into the germ tube or rhizoids. This architecture is typical of fungi, where actin patches are associated with endocytosis and cell wall deposition, whereas actin cables are pathways for targeted delivery of exocytic vesicles (Lichius et al., 2011). This biphasic actin distribution – an actin cortex and actin-filled protrusions in zoospores, and actin patches and cables in sporangia – indicates that, like its cell cycle regulatory network (Medina et al., 2016), the actin cytoskeleton of Spizellomyces displays features that resemble those of both animal and fungal cells.

Our results revealed further actin dynamics and organization during the chytrid development. This includes the formation of perinuclear actin shells that are fleetingly detected by live imaging. Previous observations suggested that perinuclear actin shells of anaerobic chytrids were a fixation artifact (Li and Heath, 1994); however, our live cell data indicate that these are real, dynamic cellular structures that likely occur in many chytrids. Similar perinuclear actin shells are associated with nuclear lamina in animal cells, where they play a role in changing nuclear shape before and after mitosis (Clubb and Locke, 1998; Clubb and Locke, 1996 or when squeezing through narrow channels (Thiam et al., 2016). Although chytrids and other fungi have lost nuclear lamins, it seems likely that Spizellomyces perinuclear actin shells are associated with changes in nuclear shape during the cell cycle.

Finally, we showed that polymerized actin is likely involved in the formation of zoospore polygonal territories (Figure 5B) before the formation of the cleavage planes during cellularization. In contrast to cellularization in Drosophila, which occurs at the surface of the embryo (i.e. two-dimensional cellularization), cellularization in Spizellomyces happens in a three-dimensional context within 3 hr. The chytrid cellularization process is reminiscent of the polarized epithelium of the social amoeba Dictyostelium discoideum (Dickinson et al., 2011; Dickinson et al., 2012) and the membrane invagination dynamics during cellularization in the ichthyosporean Sphaeroforma arctica, an unicellular relative of animals (Dudin et al., 2019). To what extent is the emergence of multicellularity, which appears to have evolved independently in amoeba, animals, and fungi, dependent upon a shared ancestral toolkit? Establishing shared and unique traits between chytrid fungi and other key lineages will provide a powerful cross-lineage experimental system to test core hypotheses on the evolution of multicellularity and derived fungal features. Based on our tools and findings, we expect that Spizellomyces will be a useful model system to study the evolution of key animal and fungal traits.

Materials and methods

Strains and growth conditions

We used Spizellomyces punctatus (Koch type isolate NG-3) Barr (ATCC 48900) for all chytrid experiments. Unless otherwise stated, Spizellomyces were grown at 30°C in Koch’s K1 medium (1L; 0.6 g peptone, 0.4 g yeast extract, 1.2 g glucose, 15 g agar if plates; Koch, 1957). Two days prior to harvesting zoospores, we aliquoted and spread 1 mL of active, liquid culture pregrown in K1 medium onto K1 plates and incubated them to allow zoospores to encyst, mature, and colonize the agar surface. We flooded each active Spizellomyces plate with 1 mL of dilute salt (DS) solution (Machlis, 1953) and incubated at room temperature. After 1 hr, released zoospores were retrieved by harvesting the DS medium and purified by slowly filtering the harvest in Luer-Lok syringe through an autoclaved syringe filter holder (Advantec REF:43303010) preloaded with a 25 mm Whatman Grade one filter paper (CAT No. 1001–325). Strains listed in Table 3 are available from the Buchler lab upon request.

Plasmids

We initially tried to transform Spizellomyces via zoospore electroporation using a protocol developed in zoosporic protists, such as Phytophthora (Ah-Fong et al., 2018). This was unsuccessful and we turned to Agrobacterium-mediated transformation because it has worked in other fungi. We used the pGI3 plasmid backbone for Agrobacterium-mediated transformation (Ianiri et al., 2017), which contains the Saccharomyces cerevisiae 2µ replication origin and the URA3 selectable marker. This allows pGI3 and its derivatives to replicate in E. coli, A. tumefaciens and S. cerevisiae. Complete details of primers and plasmid construction are in Table 2 and the Appendix. All plasmids are available from Addgene and their RRIDs are listed in the Appendix.

Table 2.

Primers used in this study.

Capital letters in SLIC and Gibson primers indicate template binding regions.

Plasmid creation	Target amplicon (Source DNA)	Primer name	Primer sequence (5’→ 3’)
pEM01	CMV promoter (pAB1T7)	CMV_F	ccctcactaaagggaacaaaagctggagctgagctcGACATTGATTATTGACTAGTTATTAATAG
pEM01	CMV promoter (pAB1T7)	CMV_R	ctttttacccatgttaattaaAGCTCTGCTTATATAGACCTCC
pEM01	hph (pRS306H)	Hyg_F	tatataagcagagctttaattaacATGGGTAAAAAGCCTGAACTC
pEM01	hph (pRS306H)	Hyg_R	tagaagtggcgcgccaTTATTCCTTTGCCCTCGGAC
pEM01	ADH1 terminator (pNB780)	ADH1t_F	agggcaaaggaataatggcgcgccACTTCTAAATAAGCGAATTTCTTATG
pEM01	ADH1 terminator (pNB780)	ADH1t_R	tgacccggcggggacgaggcaagctaaacaATATTACCCTGTTATCCCTAGC
pEM03	Hsp70 promoter (Spun gDNA)	HSP70_F	actaaagggaacaaaagctggagctgagctcTTTTAAAATCTTGTCTTTGTGC
pEM03	Hsp70 promoter (Spun gDNA)	HSP70_R	ggtgagttcaggctttttacccatgttaattaaATTGTGCTGATCTTTGGTCC
pEM09	hph (pFA6-GFP(S65T)-hph)	HygR_F	aagatcagcacaatttaattaATGGGTAAAAAGCCTGAACTCAC
pEM09	hph (pFA6-GFP(S65T)-hph)	HygR_R	tcctcctcctcctccTTCCTTTGCCCTCGGACG
pEM09	GFP (pFA6-GFP(S65T)-hph)	GFP_F	gcaaaggaaggaggaggaggaggaggaAGTAAAGGAGAAGAACTTTTCAC
pEM09	GFP (pFA6-GFP(S65T)-hph)	GFP_R	cgcttatttagaagtggcgcgccTATTTGTATAGTTCATCCATGC
	Sequencing	pRS-up	aacataggagccggaagcataaagtg
	Sequencing	ADHt-dn	ctgccggtagaggtgtggtcaataag
pGI3EM9	Hsp70pr-hph-GFP (pEM09)	GI3EM9IIF	cgttgtaaaacgacggccagtgccaagcttttttaaaatcttgtctttgtgcac
pGI3EM9	Hsp70pr-hph-GFP (pEM09)	GI3EM9IIR	aggaaacagctatgacatgattacgaattcccggtagaggtgtggtcaataag
pGI3EM11	H2B promoter (Spun gDNA)	prH2B_F	cgttgtaaaacgacggccagtgccaagCTTTTATGCTCCAAGCGGAG
pGI3EM11	H2B promoter (Spun gDNA)	prH2B_R	gagttcaggctttttacccattaattaaTTTGTGTGTGTGATGGATGAG
pGI3EM11	HygR_up	GI3EM11up	ggatcctcctcctccTTCCTTTGCCCTCGGACG
pGI3EM18	tdTomato (pKT356)	HygtdTom_F	cagcactcgtccgagggcaaaggaaggaggaggaggatccATGGTGAGCAAGGGCGAG
pGI3EM18	tdTomato (pKT356)	AdhtdTom_R	tcgcttatttagaagtggcgcgcctTTACTTGTACAGCTCGTCCATG
pGI3EM20C	H2A/H2B promoter (Spun gDNA)	H2B2D_F	gctttttacccatttaattaatgctgtgtaaggtgtgcg
pGI3EM20B/C	H2A/H2B promoter (Spun gDNA)	H2B2D_R	gggtgccatgtcgacttgtgtgtgtgatggatgag
pGI3EM20B/C	H2B CDS (Spun cDNA)	H2Bgen_F	atcacacacacaagtcgacatggcacccaaggaagctc
pGI3EM20B/C	H2B CDS (Spun cDNA)	H2Bgen_R	gacctcctcgcccttgctcaccatggatccggaggaggatttagcagactggtacttcgtcac
pGI3EM20B/C	HygR (pEM03)	AdhHygF	cgttgtaaaacgacggccagtgccaagcttccggtagaggtgtggtcaataag
pGI3EM20B/C	HygR (pEM03)	AdhHygR	ccttacacagcattaattaaatgggtaaaaagcctgaactc
pGI3EM20B/C	invPCR T-DNA (LB)	4_LB_R	tgtggaattgtgagcggata
pGI3EM20B/C	invPCR/sequencing T-DNA (LB)	ai077_F	agaggcggtttgcgtattgg
pGI3EM22C	LifeAct (Synthesized DNA)	TwistLifeact_F	ctataaaaggcgggcgtgt
pGI3EM22C	LifeAct (Synthesized DNA)	TwistLifeact_R	gcgcatgaactctttgatga
pGI3EM29	mCitrine (mCitrine-PCNA-19-SV40NLS-4)	EM18Citrine_F	ccgagggcaaaggaaggaggaggaggatccATGGTGAGCAAGGGCGAG
pGI3EM29	mCitrine (mCitrine-PCNA-19-SV40NLS-4)	EM18Citrine_R	aagaaattcgcttatttagaagtggcgcgcCTTGTACAGCTCGTCCATGC
pGI3EM30	mClover3 (pKK-mClover3-TEV)	EM18Clover3_F	gagggcaaaggaaggaggaggaggatccATGGTGAGCAAGGGCGAG
pGI3EM30	mClover3 (pKK-mClover3-TEV)	EM18Clover3_R	gaaattcgcttatttagaagtggcgcgcCTTGTACAGCTCGTCCATGC
pGI3EM31	mCerulean3 (mCerulean3-N1)	EM18Cerulean3_F	ccgagggcaaaggaaggaggaggaggatccATGGTGAGCAAGGGCGAG
pGI3EM31	mCerulean3 (mCerulean3-N1)	EM18Cerulean3_R	aagaaattcgcttatttagaagtggcgcgccTTACTTGTACAGCTCGTCCATG
	mCitrine/Clover/Cerulean Sequencing	mClover_Down	gtccaagctgagcaaagacc
	PCR screen (Set C)/H2Bpr/Sequencing	H2BprF1	tttatgctccaagcggagac
	PCR screen (Set C)/Sequencing	TomR1	cttgtacagctcgtccatgc
	PCR screen (Set D)/H2Bpr/Sequencing	H2BprF2	cgttaaatgacctgctcgaa
	PCR screen (Set D)/H2Bpr/Sequencing	TomR2	ccatgccgtacaggaacag
	PCR screen (Set A)/Southern Blot/Sequencing	HygF1	gatgtaggagggcgtggata
	PCR screen (Set A)/Southern Blot/Sequencing	HygR1	atttgtgtacgcccgacagt
	PCR screen (Set B)/Sequencing	HygF2	gtcctgcgggtaaatagctg
	PCR screen (Set B)/Sequencing	HygR2	cgtctgctgctccatacaag

Agrobacterium-mediated transformation of Spizellomyces

We prepared competent Agrobacterium EHA105 strains following the protocol of Weigel and Glazebrook (2006). Plasmids were transformed into competent Agrobacterium using 0.2 cm cuvettes in a Gene Pulser electroporator (Bio-Rad, USA) at 25 µF, 200 Ω , 2.5kV. Single colonies were streaked on selective plates (Kanamycin). A colony of transformed Agrobacterium containing pGI3-derived plasmid was grown overnight at 30°C in 5 mL of Luria-Bertani broth supplemented with Kanamycin (50 mg/L). After centrifugation, the cell pellet was resuspended in 5 mL of IM (Bundock et al., 1995), diluted to an OD660 of 0.1 and grown under agitation at 30°C until achieving a final OD660 of 0.6, at which point the culture was ready for co-culturing with the chytrid (300 µL per transformation). IM is composed of MM salts (Hooykaas et al., 1979) and 40M 2-(N-morpholino)ethanesulfonic acid (MES) pH 5.3, 10 mM glucose, 0.5% (w/v) glycerol and 200µM acetosyringone.

In parallel, chytrid zoospores were harvested and pelleted by centrifugation at 800 g for 10 min. Zoospores were then gently resuspended in 300µL of induction medium (IM). We found that one K1 plate provides enough zoospores for one transformation. For every transformation, zoospores and Agrobacterium were combined at four different ratios: 1:1, 1:0.25, 0.25:1, and 0.25:0.25 in a total volume of 200µL. To guarantee tight contact between A. tumefaciens-S. punctatus cells, the surface of the IM plate was rubbed with the bottom of a sterile glass culture tube to generate slightly concave depressions in each quadrant of the plate, wherein each 200µL co-incubation mixture was spotted. Plates were incubated unsealed for 4 days at room temperature. Mock transformations with empty Agrobacterium (no binary plasmid; grown in the absence of plasmid selective medium) were included as a negative control.

After co-incubation, we added 1 mL of DS solution and gently scraped the plate with a razor blade, pooling the different cell ratios into a single 50 mL centrifuge tube, raising the volume to 20 mL with DS solution, and re-suspending clumps by inversion. The mixture was centrifuged at 1000 g for 10 min and the liquid phase was discarded. The remaining pellet was carefully resuspended with DS solution, plated on K1 plates containing Ampicillin (50 mg/L) and Tetracycline (50 mg/L) to select against Agrobacterium and Hygromycin (200 mg/L) to select for transformed Spizellomyces. Spizellomyces survival controls were performed by plating transformations after co-culture in non-selective K1 media (Ampicillin (50 mg/L), Tetracycline (50 mg/L)). Transformation plates and controls were incubated at 30°C until colonies were observed (5–6 days). All plates were sealed with parafilm to prevent desiccation. Single colony isolates were retrieved with a sterile needle, resuspended in DS solution and re-plated on a selective Hygromycin plate. Chytrid strains listed in Table 3 are available from the Buchler lab upon request.

Table 3.

Chytrid strains available from the Buchler lab upon request.

Plasmid column lists Agrobacterium plasmids from Appendix 1 used to create chytrid strains. Integrated gene(s) are described using yeast genetic nomenclature.

Request ID	Strain name	Plasmid	Integrated gene(s)
NBC24	EM11-1	pGI3EM11	H2Bpr-hph-GFP-ScADH1ter
NBC34	EM18-1	pGI3EM18	H2Bpr-hph-tdTomato-ScADH1ter
NBC25	EM20C-1	pGI3EM20C	H2Bpr-H2B-tdTomato-ScADH1ter:H2Apr-hph-ScADH1ter
NBC26	EM20C-2	pGI3EM20C	H2Bpr-H2B-tdTomato-ScADH1ter:H2Apr-hph-ScADH1ter
NBC27	EM20C-3	pGI3EM20C	H2Bpr-H2B-tdTomato-ScADH1ter:H2Apr-hph-ScADH1ter
NBC28	EM20C-4	pGI3EM20C	H2Bpr-H2B-tdTomato-ScADH1ter:H2Apr-hph-ScADH1ter
NBC1	EM22C-1	pGI3EM22C	H2Bpr-Lifeact-tdTomato-ScADH1ter:H2Apr-hph-ScADH1ter
NBC71	EM29-1	pGI3EM29	H2Bpr-hph-mCitrine-ScADH1ter
NBC55	EM30-1	pGI3EM30	H2Bpr-hph-mClover3-ScADH1ter
NBC44	EM31-1	pGI3EM31	H2Bpr-hph-mCerulean3-ScADH1ter

Nucleic acid manipulation

High-molecular-weight genomic DNA extraction was performed using CTAB/Chloroform protocol (CTAB/PVP buffer: 100 mM Tris-HCl pH7.5; 1.4M NaCl; 10 mM EDTA; 1% CTAB; 1% PVP; 1% - Mercaptoethanol (added just before use)). Briefly, 10 plates of the selected strain were grown at 30°C for 2 days and zoospores were harvested and purified as described before. The pellet of zoospores was resuspended directly in 900µL of CTAB/PVP buffer pre-warmed to 65°C and transferred to 2 mL centrifuge tubes. Tubes were centrifuged briefly and incubated at room temperature for 1 hr in a nutating mixer. After 5 min incubation on ice, DNA was extracted with Chloroform (Sigma-Aldrich REF:288306) twice followed by treatment of supernatant with 100 ng RNase A (Biobasic; 60 U/mg Ref:XRB0473) for 30 min at room temperature. DNA was then precipitated by adding 0.2 vol of 10M Ammonium acetate and one volume of absolute Isopropanol and incubated at 4°C overnight. DNA pellet was washed with 70% ethanol thrice and resuspended in TE buffer. DNA quality and concentration were determined by gel electrophoresis.

Detection of transgene integration by Southern blot. 1 µg of high-molecular-weight gDNA from each strain was treated overnight at 37°C with 10U of XbaI or KpnI-HF restriction enzymes, resolved on a 1% Agarose 1X TAE (Tris-Acetate-EDTA) gel and blotted to a GE Healthcare Amersham Hybond-N+ membrane. The membrane was hybridized with a 809 bp fragment of the Hygromycin resistance gene amplified with primers HygF1 and HygR1 and radiolabeled with [-32P]-dCTP using the Prime-It II Random Primer Labelling Kit (Agilent Technologies; REF:300385) following manufacturer’s instructions.

Identification of T-DNA insertion sites by inverse PCR. 2.5 µg of genomic DNA of wild type (WT) and the four transformed strains of Spizellomyces (EM20C-1,2,3,4) was digested in a final volume of 50μL with 100U of EcoRI-HF (NEB R3101S) or HindIII-HF (NEB R3104S) for 24 hr at 37°C. After assessing the quality of the digestion by gel electrophoresis, the reaction was heat inactivated and 48µL of the digested DNA was incubated with 1µL of T4 ligase (400u/µL) for 48 hr at 4°C. The ligation was purified by chloroform extraction twice, followed by DNA precipitation and resuspended in 30µL of nuclease-free water. 100 ng of ligated product was used for touch-down Inverse PCR reactions in a final volume of 50µL using NEB Phusion high-fidelity DNA polymerase and 3% DMSO following manufacturer instructions and primers ai77_F and 4_LB_R. Touch-down PCR amplification protocol included an initial denaturation step at 98°C for 3 min followed by 10 cycles of amplification in which the annealing temperature was decreased 1C/cycle until an annealing temperature of 62°C was achieved, followed by 20 amplification cycles at 62°C. Annealing and elongation time during all these cycles was 30 s and 6 min, respectively. Amplification was assessed by gel electrophoresis and bands were retrieved using a razor blade, purified using the Promega Wizard SV Gel and PCR Clean-Up System, and Sanger-sequenced using the primer ai77_F.

Microscopy

H2B-tdTomato-expressing chytrids were harvested from plates, placed in a glass-bottom dish (Mattek), and covered with a 1.5% K1 agarose pad to keep cells healthy and in the plane of focus (Young et al., 2012). We harvested zoospores from plates and re-suspended them in Leu/Lys paralyzing solution (Dill and Fuller, 1971 ) before putting them in glass-bottom dishes, as above. Live-cell epifluorescence was performed on a temperature-controlled Deltavision Elite inverted microscope equipped with 60x/1.42 oil objective and Evolve-512 EMCCD camera using optical axis integration. Optical axis integration (OAI) is a setting on the Deltavision microscope that opens the shutter and continuously excites and measures fluorescence emission while sweeping from top (+20 microns) to bottom of the z-axis (−20 microns) in 0.5 s, integrating directly the intensities onto the CCD chip. OAI has the advantage that total exposure time is reduced relative to a traditional z-stack. Excitation light was 542/27 nm (7 Color InsightSSI) and emission to the camera was filtered by 594/45 nm (TRITC). Epifluorescence live-cell imaging was done at 30°C. Live-cell confocal microscopy was performed on a Nikon Eclipse Ti inverted microscope equipped with a 100x/N.A. 1.49 CFI Apo TIRF oil objective and fitted with a Yokogawa CSU-X1 spinning disk and Andor iXon 897 EMCCD camera. Excitation light was via 488 nm laser. Confocal live-cell imaging was done at 27°C.

LifeAct-expressing zoospores were collected in DS solution and transferred to cover-glass bottom dishes. For phalloidin staining, glass coverslips were plasma cleaned and immediately coated with 0.1% polyethyleneimine for 5 min, washed thrice with water, then overlaid with zoospores or sporangia suspended in DS solution. Cells were allowed to adhere for 5 min before fixation by adding four volumes of 4% paraformaldehyde in 50 mM Cacodylate buffer (pH 7.2). Cells were fixed for 20 min on ice, washed once with PEM buffer (100 mM PIPES, 1 mM EGTA, 0.1 mM MgSO4), permeabilized and stained with 0.1% Triton X in PEM with 1:1000 Alexa Fluor 488 Phalloidin (66 nM in DMSO, Sigma D2660) for 10 min at room temperature, washed with PEM, then mounted onto glass slides using Prolong Gold with DAPI (Invitrogen P36931). Zoospores were imaged on a Nikon Ti2-E inverted microscope equipped with 100x oil PlanApo objective and sCMOS 4mp camera (PCO Panda). Excitation light was via epi fluorescence illuminator at 405 nm, 488 nm, and 561 nm. Sporangia were imaged on a Nikon Ti-E inverted microscope equipped with 100X oil objective and fitted with a Yokogawa X1 spinning disk (CSU-W1) with 50 μm pinholes and Andor xIon EMCCD camera. Excitation light was via 405 nm laser, 488 nm laser, and 561 nm laser. Z-stack fluorescent images were deconvolved with NIS Elements v5.11 using 20 iterations of the Richardson-Lucy algorithm. Image analysis was performed with the ImageJ bundle Fiji (Schindelin et al., 2012). All imagings were done at room temperature.

Flow cytometry

Zoospores from wild-type and EM20C-(1-4) transformants were harvested from plates and put on ice until fluorescence measurement. Flow cytometry was performed on a MACSQuant VYB using the yellow laser (561 nm) to measure forward scatter (FSC), side scatter (SSC), and tdTomato fluorescence (Y2, 615/20 nm filter). The FSC, SSC, Y2 voltage gain settings for each PMT were 350 V, 350 V, and 510 V, respectively. We recorded the pulse height, width, area for a minimum of 100,000 events per strain. Singlets were identified using FSC/SSC width versus height, and a histogram of singlet tdTomato fluorescence (Y2 height) was plotted using the FlowJo software package.

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This manuscript reports the first description of genetic transformation of a species of chytrid fungi, a group that contains the pathogens responsible for the large-scale die-offs of amphibian species. The authors demonstrate the use of Agrobacterium-mediated transformation to isolate stable transformants of Spizellomyces punctatus. This work is likely to be the starting point for work on the molecular biology of chytrids and has potential to establish this chytrid as a comparative model for understanding the evolution of fungal and animal multicellularity and cell biology.

Decision letter after peer review:

Thank you for submitting your article "Genetic transformation of Spizellomyces punctatus sheds light on the evolution of animal and fungal traits" for consideration by eLife. Your article has been reviewed by Ian Baldwin as the Senior Editor, a Reviewing Editor, and two reviewers. The following individuals involved in review of your submission have agreed to reveal their identity: Tim Stearns (Reviewer #2).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This manuscript reports the first description of genetic transformation of a species of chytrid fungi. This is an important technical advance as this fungal group includes B. dendrobatidis and B. salamandrivorans, two species responsible for large-scale die-off of amphibian species. Although chytrid fungi have been known for more than a century, there has been relatively little genetics and molecular biology done with them, aside from genome sequencing and transcriptomics. The authors demonstrate the use of Agrobacterium-mediated transformation to isolate stable transformants of Spizellomyces punctatus (Sp), a species that they have previously used as a tool for understanding the evolution of cell cycle control pathways. This work is likely to be the starting point for much more work on the biology of chytrids and the expansion of the molecular genetic manipulations to other chytrid species. In addition, this work has potential to establish this chytrid as a comparative model for understanding the evolution of fungal and animal multicellularity and cell biology.

Given that this is a "Tools and Resources" article, we encourage the authors to revise the title to one that emphasizes the tool they have developed and its potential for driving discovery.

Essential revisions:

1) Given that this is a "Tools and Resources" article with potential to help the community perform experiments in chytrids, we recommend that the authors make strains and plasmids (with a full sequence and map) generally available. Deposition into Addgene or other comparable databases would fulfil this request.

2) While the reviewers appreciate the use of a good, old-fashioned southern to show integration, we would like to see more commentary and/or more data that indicate where sites of integration are. Where are the different transgenes inserted? Are insertion sites based on homology or randomly integrated? Based on insertion sites what is the prospect for performing reverse genetics, not just transgenesis?

3) Along similar lines, southern blotting is used to show that the stable transformants are likely to be single integration events into different loci in the Sp genome. Since the different clones all have different integration sites, the authors should be able to say something about variability of expression of the transformed fluorescent protein gene, based on fluorescence intensity. This is important information, as it would help to understand the range of expression one might expect in clones from a transformation experiment, based on position effect of the random integration site in the genome.

4) Figure 1C should be better labeled. What is (+)? What is the size of the integrating plasmid? Knowing the size would help the reader better evaluate that the larger sized fragments on the southern are true integration sites. Related to this topic, what are the regions surrounding the expression cassette in Figure 2—figure supplement 3? What does RB and LB stand for?

5) In Figure 5A, what explains the difference in LifeAct versus phalloidin localization in the sporangium? (Right most set of panels, green versus magenta). The authors skip over that explanation.

6) The authors describe the use of Agrobacterium for transformation in the beginning of the Results section. Since the central advance in this work is the transformation of Sp, and there is likely to be strong interest in the community in transforming other chytrids, it would be helpful to know what other methods of transformation were attempted, presumably without success.

7) There is little information about the promoter and gene segments used in the transformation plasmids. The primer sequences used to amplify them are given, and it would be possible to derive the information from them and the Sp genome sequence, but the reader should not be left having to do that. This is particularly important for the bidirectional histone promoter that is cloned and used in most their experiments. How big is the promoter fragment? Where does it start and stop with reSpecies">spect to the ATGs of the flanking coding sequences?

8) Figure 2—figure supplement 1B: Why are the three tiny colonies pointed out when there are many more on the HygR-tdTomato plate? Are all of those actually transformants, too?

9) Figure 5B, C: The merged images that are shown in these two panels detract from the presentation as they seem to be done in a way that makes them useless for seeing what is going on. Perhaps the DIC was mistakenly put in the colored channel in the merge rather than the LifeAct image?

Summary:

This manuscript reports the first description of genetic transformation of a species of chytrid fungi. This is an important technical advance as this fungal group includes B. dendrobatidis and B. salamandrivorans, two species responsible for large-scale die-off of amphibian species. Although chytrid fungi have been known for more than a century, there has been relatively little genetics and molecular biology done with them, aside from genome sequencing and transcriptomics. The authors demonstrate the use of Agrobacterium-mediated transformation to isolate stable transformants of Spizellomyces punctatus (Sp), a species that they have previously used as a tool for understanding the evolution of cell cycle control pathways. This work is likely to be the starting point for much more work on the biology of chytrids and the expansion of the molecular genetic manipulations to other chytrid species. In addition, this work has potential to establish this chytrid as a comparative model for understanding the evolution of fungal and animal multicellularity and cell biology.

Given that this is a "Tools and Resources" article, we encourage the authors to revise the title to one that emphasizes the tool they have developed and its potential for driving discovery.

We revised our title as suggested.

Essential revisions:

1) Given that this is a "Tools and Resources" article with potential to help the community perform experiments in chytrids, we recommend that the authors make strains and plasmids (with a full sequence and map) generally available. Deposition into Addgene or other comparable databases would fulfil this request.

We have deposited plasmids and sequence maps to Addgene. All chytrid strains (listed in new Table 3) will be provided by the Buchler lab upon request. We state this information in Methods and Materials.

2) While the reviewers appreciate the use of a good, old-fashioned southern to show integration, we would like to see more commentary and/or more data that indicate where sites of integration are. Where are the different transgenes inserted?

To directly address this question, we performed inverse PCR on the independent transformants from Figure 2 to map where each T-DNA is inserted into the Spizellomyces genome. Briefly, genomic DNA was digested with a restriction enzyme followed by ligation to create circularized DNA. Primers that anneal inside the T-DNA close to the left border (LB) and amplify outwards were used for PCR. Positive products were sequenced to recover the junction between the LB and genomic DNA, thus identifying the insertion sites in the genome. Our results (Figure 2—figure supplement 4) show that each transformant is a T-DNA integration event in a different genomic locus, as would be expected from Agrobacterium-mediated transformation.

EM20C-1: The T-DNA is integrated in the proximal upstream region (58 bp) of the gene SPPG_04375, a member of the M48 peptidase family.

EM20C-2: The T-DNA is integrated in the proximal upstream region (105 bp) of the gene SPPG_03425, a member of the Usp-like/ adenine nucleotide α-hydrolase family.

EM20C-3: The T-DNA is integrated in the proximal upstream region (189 bp) of the gene SPPG_02523, a PHO-like cyclin. Inverse PCR with EcoRI suggests that LB is closest to SPPG_02523, whereas inverse PCR with HindIII suggests an inverted orientation of the T-DNA in the same locus. One possibility is an irregular integration of T-DNA into the genomic locus. Resolving the architecture of this T-DNA integration through additional inverse PCR or genome sequencing seems beyond the scope of the manuscript because EM20C-3 was not used for downstream experiments.

EM20C-4: The T-DNA is integrated in the distal upstream region (845 bp) of the gene SPPG_08788, a hypothetical protein with no clear conserved protein domains.

Are insertion sites based on homology or randomly integrated? Based on insertion sites what is the prospect for performing reverse genetics, not just transgenesis?

Agrobacterium T-DNA is often randomly integrated into the host chromosome as a single copy and, if inserted within a gene, it will disrupt gene function and act as a mutagen (Gelvin, 2017). Random insertional mutagenesis by Agrobacterium has been exploited for forward genetics in plants and fungi (Alonso et al. 2003; Michielse et al., 2005; Idnurm et al., 2017). In fungi, Agrobacterium-mediated transformation has also been used for targeted gene knock-outs by including 1–1.5 kb long, flanking region of homology in the T-DNA to the desired target gene (Frandsen et al., 2012). The feasibility of this approach relies on the rate of homologous recombination, which is in general high in the majority of Fungi. The low efficiency of homologous recombination in other organisms, such as Arabidopsis, has been improved by creating doublestrand breaks in target genes using CRISPR/Cas9.

Although beyond the scope of the current manuscript, the prospect for reverse genetics in Spizellomyces and other chytrid fungi looks promising. We plan to use T-DNA plasmids with longflanking homology for targeted gene knock-out/knock-in, and concomitantly create T-DNA plasmids that express Cas9 and sgRNA.

We have incorporated parts of this discussion in the manuscript.

3) Along similar lines, southern blotting is used to show that the stable transformants are likely to be single integration events into different loci in the Sp genome. Since the different clones all have different integration sites, the authors should be able to say something about variability of expression of the transformed fluorescent protein gene, based on fluorescence intensity. This is important information, as it would help to understand the range of expression one might expect in clones from a transformation experiment, based on position effect of the random integration site in the genome.

Indeed, as observed in Arabidopsis, we might expect variation in gene expression based on the genomic locus of integration of the T-DNA fragment. To quantify this variation, we measured H2BtdTomato expression in our EM20C transformants in Figure 2 using flow cytometry. Briefly, we grew different transformants and wild type (negative control), harvested zoospores, and measured the tdTomato fluorescence distribution. The data (Figure 2—figure supplement 4) show that H2B-tdTomato expression is identical in 3 out of the 4 transformants, despite the different sites of genomic integration. The exception is EM20C-1, which exhibits bimodal gene expression: the top mode is identical to the other transformants, but the bottom mode is half the intensity. The bimodality is not due to a mixed population with variable T-DNA copy number and/or genomic integrations because Southern blot and inverse PCR data indicate a single copy integration event in EM20C-1. Epigenetic suppression of T-DNA has been observed in Arabidopsis, and bimodal H2B-tdTomato expression in EM20C-1 might arise from epigenetic regulation of the transgene in its specific genomic locus.

Our previous time lapse movies (Figure 2 and Figure 3) were done with transformant EM20C-1. To check whether the location and/or expression level of H2B-tdTomato transgene might affect previous conclusions regarding chytrid development and cell cycle, we made replicate time-lapse fluorescence movies of EM20C-1 and EM20C-2 (Figure 2—figure supplement 5). We scored the timing of germ tube formation, mitosis (i.e. nuclear division), and sporangium bursting in single cells. For either strain, one replicate initiated the developmental program earlier than the other replicate by 1-2 hours. This systematic time shift between replicates, which propagates to downstream events, is likely due to a (currently) unknown and uncontrolled factor that regulates early developmental events. Despite this systematic time shift between replicates, both EM20C1 and EM20C-2 formed germ tubes at 1-3 hours, the first mitosis occurred between 8-12 hours, and the average cell cycle period in early development was ~150 minutes. This cell cycle period is identical to that of EM20C-1, previously measured by confocal fluorescence time lapse (Figure 3). Despite the overall similarity between transformants, there is one significant difference:

EM20C-2 had a shorter developmental cycle than EM20C-1 (bursting at 20 – 25 hours versus 25 – 30 hours). This stemmed from a combination of shorter cell cycle periods in late development and fewer mitotic cycles before bursting.

We have incorporated parts of this discussion in the manuscript. We also modified the timing of developmental events in the caption of Figure 1 to mirror EM20C-1 and EM20C-2 time-lapse data from Figure 2—figure supplement 5.

4) Figure 1C should be better labeled. What is (+)? What is the size of the integrating plasmid? Knowing the size would help the reader better evaluate that the larger sized fragments on the southern are true integration sites. Related to this topic, what are the regions surrounding the expression cassette in Figure 2—figure supplement 3? What does RB and LB stand for?

Figure 2C: The (+) refers to the positive control (GI3EM20C plasmid). We now describe this control in the caption.

Figure 2—figure supplement 3: RB and LB are the "right border" and "left border" of the T-DNA plasmid. These sequences define the T-DNA fragment that is excised and injected into the host cell by the Agrobacterium virulence machinery upon induction. The total size of the T-DNA (left border, LB, to right border, RB) is 4280 bp. We now include this information in the figure legend.

5) In Figure 5A, what explains the difference in LifeAct versus phalloidin localization in the sporangium? (Right most set of panels, green versus magenta). The authors skip over that explanation.

The minor deviations between LifeAct and phalloidin can be explained by intrinsic biases of the probes. All live-cell probes of filamentous actin (e.g. LifeAct, F-tractin, Utrophin actin binding domain) have different biases in their patterns of F-actin localization and dynamics (Belin et al., 2015). None of them can fully recapitulate the patterns observed with phalloidin.

We now clarify this in the text.

6) The authors describe the use of Agrobacterium for transformation in the beginning of the Results section. Since the central advance in this work is the transformation of Sp, and there is likely to be strong interest in the community in transforming other chytrids, it would be helpful to know what other methods of transformation were attempted, presumably without success.

We initially tried zoospore electroporation using a protocol developed in zoosporic protists, such as Phytophthora (Ah-Fong et al., 2018). This was unsuccessful and we turned to Agrobacterium-mediated transformation because it has worked in other fungi and because we had access to the pGI3-based plasmids and expertise of Giuseppe Ianiri (co-author and former postdoc at Duke University in the lab of Joe Heitman). This worked and, thus, we did not continue exploring other methods of transformation (e.g. protoplast lithium acetate, biolistics, etc.).

We have added a brief description in the Materials and methods section.

7) There is little information about the promoter and gene segments used in the transformation plasmids. The primer sequences used to amplify them are given, and it would be possible to derive the information from them and the Sp genome sequence, but the reader should not be left having to do that. This is particularly important for the bidirectional histone promoter that is cloned and used in most their experiments. How big is the promoter fragment? Where does it start and stop with reSpecies">spect to the ATGs of the flanking coding sequences?

The shared/combined promoter region of H2B and H2A is 217 bp. We also included endogenous 5’UTR regions of the H2A (118 bp) and H2B (66bp), resulting in a total of 401 bp. This cloned bidirectional "promoter region" is followed by restriction sites and the start codons of the corresponding proteins of interest (hph or H2B CDS). This is schematically depicted in Figure 2—figure supplement 3A.

We also include this specific information in the Appendix and updated Figure 2—figure supplement 3A.

8) Figure 2—figure supplement 1B: Why are the three tiny colonies pointed out when there are many more on the HygR-tdTomato plate? Are all of those actually transformants, too?

We highlighted these colonies because they are difficult to see without a visual cue, unlike the hph-tdTomato plate. We did not test all hph-tdTomato transformants with PCR and fluorescence microscopy, but we suspect that most of them would be true transformants. The frequency of hygromycin resistant colonies in hph-GFP and hph-tdTomato was consistently higher than the no plasmid control (i.e. no colonies).

We now show photos of the transformation plates using inverted contrast to better highlight the colonies.

9) Figure 5B, C: The merged images that are shown in these two panels detract from the presentation as they seem to be done in a way that makes them useless for seeing what is going on. Perhaps the DIC was mistakenly put in the colored channel in the merge rather than the LifeAct image?

When multiple channels are shown for an image (including DIC) it is best practice to include a merged image of the independent channels (Lee and Kitaoka, 2018). If the journal agrees, we would prefer to keep the merges.