About

Nick Buchler is an Associate Professor at NC State University within the Department of Molecular Biomedical Sciences. His research focuses on molecular biomedical sciences, contributing to the understanding of biological barriers, immunology, infectious diseases, and regenerative medicine. As part of his academic role, he is involved in advancing knowledge in these areas through research and teaching, supporting the development of veterinary and biomedical sciences.

Research topics

• Biology
• Evolutionary biology
• Genetics
• Ecology
• Botany
• Cell biology

Selected publications

• Deep learning–driven imaging of cell division and cell growth across an entire eukaryotic life cycle

  Molecular Biology of the Cell · 2025-05-06 · 2 citations

  article

  The life cycle of eukaryotic microorganisms involves complex transitions between states such as dormancy, mating, meiosis, and cell division, which are often studied independently from each other. Therefore, most microbial life cycles are theoretical reconstructions from partial observations of cellular states. Here we show that complete microbial life cycles can be directly and continuously studied by combining microfluidic culturing, life cycle stage-specific segmentation of micrographs, and a novel cell tracking algorithm, FIEST, based on deep learning video frame interpolation. As proof of principle, we quantitatively imaged and compared cell growth and the activity state of the cell division kinase, Cdk1, across the life cycle of Saccharomyces cerevisiae for up to three sexually reproducing generations. Our analysis of S. cerevisiae's life cycle provided the following new insights: 1) the accumulation of cell cycle regulators, such as Whi5, is tailored to each life cycle stage; 2) cell growth always preceded exit from nonproliferative states in our conditions; 3) the temporal coordination of meiotic events is the same across sexually reproducing populations when each generation is exposed to same conditions; 4) information such as cell size and morphology resets after each sexual reproduction cycle. Image processing and tracking algorithms are available as the Python package Yeastvision, which could be used study pathogens such as Candida glabrata, Cryptococcus neoformans, Colletotrichum acutatum, and other unicellular systems.

  PublisherDOI

• Deep learning-driven imaging of cell division and cell growth across an entire eukaryotic life cycle

  bioRxiv (Cold Spring Harbor Laboratory) · 2024-04-27 · 4 citations

  preprintOpen access

  Abstract The life cycle of biomedical and agriculturally relevant eukaryotic microorganisms involves complex transitions between proliferative and non-proliferative states such as dormancy, mating, meiosis, and cell division. New drugs, pesticides, and vaccines can be created by targeting specific life cycle stages of parasites and pathogens. However, defining the structure of a microbial life cycle often relies on partial observations that are theoretically assembled in an ideal life cycle path. To create a more quantitative approach to studying complete eukaryotic life cycles, we generated a deep learning-driven imaging framework to track microorganisms across sexually reproducing generations. Our approach combines microfluidic culturing, life cycle stage-specific segmentation of microscopy images using convolutional neural networks, and a novel cell tracking algorithm, FIEST, based on enhancing the overlap of single cell masks in consecutive images through deep learning video frame interpolation. As proof of principle, we used this approach to quantitatively image and compare cell growth and cell cycle regulation across the sexual life cycle of Saccharomyces cerevisiae . We developed a fluorescent reporter system based on a fluorescently labeled Whi5 protein, the yeast analog of mammalian Rb, and a new High-Cdk1 activity sensor, LiCHI, designed to report during DNA replication, mitosis, meiotic homologous recombination, meiosis I, and meiosis II. We found that cell growth preceded the exit from non-proliferative states such as mitotic G1, pre-meiotic G1, and the G0 spore state during germination. A decrease in the total cell concentration of Whi5 characterized the exit from non-proliferative states, which is consistent with a Whi5 dilution model. The nuclear accumulation of Whi5 was developmentally regulated, being at its highest during meiotic exit and spore formation. The temporal coordination of cell division and growth was not significantly different across three sexually reproducing generations. Our framework could be used to quantitatively characterize other single-cell eukaryotic life cycles that remain incompletely described. An off-the-shelf user interface Yeastvision provides free access to our image processing and single-cell tracking algorithms.

  PublisherOA PDFDOI

• Gene expression noise accelerates the evolution of a biological oscillator

  arXiv (Cornell University) · 2022-03-21

  preprintOpen accessSenior author

  Gene expression is a biochemical process, where stochastic binding and un-binding events naturally generate fluctuations and cell-to-cell variability in gene dynamics. These fluctuations typically have destructive consequences for proper biological dynamics and function (e.g., loss of timing and synchrony in biological oscillators). Here, we show that gene expression noise counter-intuitively accelerates the evolution of a biological oscillator and, thus, can impart a benefit to living organisms. We used computer simulations to evolve two mechanistic models of a biological oscillator at different levels of gene expression noise. We first show that gene expression noise induces oscillatory-like dynamics in regions of parameter space that cannot oscillate in the absence of noise. We then demonstrate that these noise-induced oscillations generate a fitness landscape whose gradient robustly and quickly guides evolution by mutation towards robust and self-sustaining oscillation. These results suggest that noise can help dynamical systems evolve or learn new behavior by revealing cryptic dynamic phenotypes outside the bifurcation point.

  PublisherOA PDFDOI

• Gene expression noise accelerates the evolution of a biological oscillator

  bioRxiv (Cold Spring Harbor Laboratory) · 2022-03-23

  preprintOpen accessSenior author

  Abstract Gene expression is a biochemical process, where stochastic binding and unbinding events naturally generate fluctuations and cell-to-cell variability in gene dynamics. These fluctuations typically have destructive consequences for proper biological dynamics and function (e.g., loss of timing and synchrony in biological oscillators). Here, we show that gene expression noise counter-intuitively accelerates the evolution of a biological oscillator and, thus, can impart a benefit to living organisms. We used computer simulations to evolve two mechanistic models of a biological oscillator at different levels of gene expression noise. We first show that gene expression noise induces oscillatory-like dynamics in regions of parameter space that cannot oscillate in the absence of noise. We then demonstrate that these noise-induced oscillations generate a fitness landscape whose gradient robustly and quickly guides evolution by mutation towards robust and self-sustaining oscillation. These results suggest that noise can help dynamical systems evolve or learn new behavior by revealing cryptic dynamic phenotypes outside the bifurcation point. Graphical Abstract

  PublisherOA PDFDOI

• Diploid-dominant life cycles characterize the early evolution of Fungi

  Proceedings of the National Academy of Sciences · 2022 · 80 citations

  • Biology
  • Evolutionary biology
  • Genetics

  Most of the described species in kingdom Fungi are contained in two phyla, the Ascomycota and the Basidiomycota (subkingdom Dikarya). As a result, our understanding of the biology of the kingdom is heavily influenced by traits observed in Dikarya, such as aerial spore dispersal and life cycles dominated by mitosis of haploid nuclei. We now appreciate that Fungi comprises numerous phylum-level lineages in addition to those of Dikarya, but the phylogeny and genetic characteristics of most of these lineages are poorly understood due to limited genome sampling. Here, we addressed major evolutionary trends in the non-Dikarya fungi by phylogenomic analysis of 69 newly generated draft genome sequences of the zoosporic (flagellated) lineages of true fungi. Our phylogeny indicated five lineages of zoosporic fungi and placed Blastocladiomycota, which has an alternation of haploid and diploid generations, as branching closer to the Dikarya than to the Chytridiomyceta. Our estimates of heterozygosity based on genome sequence data indicate that the zoosporic lineages plus the Zoopagomycota are frequently characterized by diploid-dominant life cycles. We mapped additional traits, such as ancestral cell-cycle regulators, cell-membrane- and cell-wall-associated genes, and the use of the amino acid selenocysteine on the phylogeny and found that these ancestral traits that are shared with Metazoa have been subject to extensive parallel loss across zoosporic lineages. Together, our results indicate a gradual transition in the genetics and cell biology of fungi from their ancestor and caution against assuming that traits measured in Dikarya are typical of other fungal lineages.

  PublisherOA PDFDOI

• Competition for DNA binding between paralogous transcription factors determines their genomic occupancy and regulatory functions

  Genome Research · 2021-05-11 · 40 citations

  articleOpen access

  Most eukaryotic transcription factors (TFs) are part of large protein families, with members of the same family (i.e., paralogous TFs) recognizing similar DNA-binding motifs but performing different regulatory functions. Many TF paralogs are coexpressed in the cell and thus can compete for target sites across the genome. However, this competition is rarely taken into account when studying the in vivo binding patterns of eukaryotic TFs. Here, we show that direct competition for DNA binding between TF paralogs is a major determinant of their genomic binding patterns. Using yeast proteins Cbf1 and Pho4 as our model system, we designed a high-throughput quantitative assay to capture the genomic binding profiles of competing TFs in a cell-free system. Our data show that Cbf1 and Pho4 greatly influence each other's occupancy by competing for their common putative genomic binding sites. The competition is different at different genomic sites, as dictated by the TFs' expression levels and their divergence in DNA-binding specificity and affinity. Analyses of ChIP-seq data show that the biophysical rules that dictate the competitive TF binding patterns in vitro are also followed in vivo, in the complex cellular environment. Furthermore, the Cbf1-Pho4 competition for genomic sites, as characterized in vitro using our new assay, plays a critical role in the specific activation of their target genes in the cell. Overall, our study highlights the importance of direct TF-TF competition for genomic binding and gene regulation by TF paralogs, and proposes an approach for studying this competition in a quantitative and high-throughput manner.

  PublisherOA PDFDOI

• Author response: Genetic transformation of Spizellomyces punctatus, a resource for studying chytrid biology and evolutionary cell biology

  2020-04-23

  peer-reviewOpen accessSenior author

  Article Figures and data Abstract Introduction Results Discussion Materials and methods Appendix 1 Data availability References Decision letter Author response Article and author information Metrics Abstract 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 Download asset Open asset 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. 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 with 6 supplements see all Download asset Open asset 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. 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). 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 (Δt) is the time for a wave of synchronous nuclear divisions to propagate across the sporangium. The cell cycle period (τ) 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 3 with 1 supplement see all Download asset Open asset 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 (τ) is the interval of time between waves of nuclear division (i.e. metaphase to anaphase transition). The wave time (Δ⁢t) is the interval of time for a synchronous wave of nuclear division to sweep across the sporangium. (B) Distribution of cell cycle period (τ), wave time (Δ⁢t) and duration of mitosis (τM) 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 (τM=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). Figure 3—source data 1 Number of nuclei per cell as a function of time (min). Data was used to create Figure 3A and analyzed to create Figure 3B. https://cdn.elifesciences.org/articles/52741/elife-52741-fig3-data1-v1.xlsx Download elife-52741-fig3-data1-v1.xlsx Table 1 Comparison of nuclear division synchrony for different coenocytic organisms. Wave time (Δt) 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 (τ) is the average interval of time between waves. Organisms are listed from highest to lowest synchrony index. Coenocytic organismLength scaleWave timeWave speedNuclear divisionSynchrony indexReferences(µm)Δt (min)(µm min-1)Period τ (min)100%⋅(1-Δ⁢t/τ)Physarum polycephalum (amoeba)1000250084099.8%Halvorsrud et al., 1995Spizellomyces punctatus (fungi)5–1015–1015099.3%This workCreolimax fragrantissima (holozoa)2030093.3%Suga and Ruiz-Trillo, 2013Drosophila melanogaster (metazoa)5001.53601085.5%Deneke et al., 2016Aspergillus nidulans (fungi)70020356066.7%Clutterbuck, 1970; Momany and Taylor, 2000 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 α-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 (Fritz-Laylin et al., 2017b). Spizellomyces has of WASP SCAR/WAVE and its zoospores are To test Spizellomyces zoospores crawl using α-motility, we expressed a is a that to polymerized actin in a of cell as actin patches and cables in yeast and of crawling animal cells et al., et al., To that our fusion to polymerized actin in Spizellomyces, we first and actin in zoospores (Figure and sporangia (Figure with fluorescent We then compared the fluorescent images of cells expressing and expressing hph-tdTomato relative to Figure 4 with 1 supplement see all Download asset Open asset of in zoospores highlights cortical and actin (A) from wild type (left), and transformants expressing hph-tdTomato (center) and were and with fluorescent Top row shows DIC and row shows DNA The bottom row shows the and 561 nm images Scale bar indicates two microns. and of and hph-tdTomato (B) and fusion The shows line of fluorescence intensity of the fusion protein and fluorescent The location for the line is by a line in the above each Scale 1 (D) at intervals from microscopy of crawling zoospores from the strains at the were using DIC microscopy and 561 nm fluorescence microscopy also with images Scale bar indicates 5 of and hph-tdTomato (E) and fusion The shows line of fluorescence intensity of the fusion protein and fluorescent The location for the line is by a line in the above each Scale 2 Figure 5 with 1 supplement see all Download asset Open asset of in sporangia highlights actin and perinuclear (A) from wild type (left), and transformants expressing hph-tdTomato (center) and were and with fluorescent The row shows DNA The bottom row shows the and 561 nm images Scale bar indicates 5 to of actin patches in sporangia and rhizoids cables and perinuclear actin shells (B) from microscopy of sporangia from the strain at of polygonal territories cellularization. using DIC and 561 nm also with images Scale bar indicates ten microns. (C) of a single sporangium show polygonal territories during later stages of cellularization the In to the of fluorescence in hph-tdTomato zoospores, both (Figure and zoospores (Figure and Figure of the strain showed a of fluorescence at the cell and high levels of fluorescence in the at the Because the fusion localization with in cells, but the hph fusion did we that this fluorescence polymerized The between and can be by of the All live-cell of actin (e.g. actin-binding have different in their of localization and dynamics et al., of them can the seen with but they provide new into the live cell dynamics of The and dynamics of actin that we see in zoospores is in with and actin localization observed for cells during zoospore crawling of and and Batrachochytrium (Fritz-Laylin et al., 2017b). Actin polymerization during sporangiogenesis Once the chytrid zoospore it a sporangium by both and in during germ tube and rhizoid formation (Figure 1). early the nuclei are while and but then in during cellularization and zoospore formation (Figure Actin has been to during cellularization in et al., et al., we expect polymerized actin to a in the nuclear dynamics and cellularization during sporangiogenesis in Spizellomyces. Our experiments an actin between cortical patches and that the actin cables of other fungal (Figure Each of these was in both cells and cells expressing the We also detected perinuclear polymerized actin shells with a period similar to mitotic events (Figure and Figure which a for polymerized actin with nuclei during the cell cycle. In polymerized actin polygonal zoospore territories (Figure This is reminiscent of cellularization seen during when nuclei are by cell Discussion Here, we stable and genetic transformation of the chytrid Spizellomyces punctatus. We identified and tested Spizellomyces including a divergent H2A/H2B that can 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 fungi not which they be for other as the Agrobacterium was used to the aquatic chytrid and Camilo, genetic transformation of a of gene expression and be detected at the The described in this study be for and gene integration in other chytrids. For example, Agrobacterium T-DNA is into the chromosome as a single inserted within a it will gene function and as a by Agrobacterium has been exploited for in plants and fungi et al., et al., et al., In Agrobacterium-mediated transformation has also been used for gene by including region of in the T-DNA to the desired gene et al., The of this in a chytrid relies on the of which is in high in the of As a first 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 a of time to the mitotic during each cell cycle with a highly synchronous wave of nuclear The of synchrony is similar to nuclei in the amoeba polycephalum or nuclei during where all nuclei divide within 2 min et al., Table 1). These mitotic dynamics Spizellomyces an comparative model for the and molecular of cell division synchrony and its evolution in fungi, and actin dynamics throughout the chytrid life cycle, we found that Spizellomyces zoospores actin and during motility, similar to other chytrid (Fritz-Laylin et al., 2017b), animal cells, and Once the zoospores there is a in actin in which the cortical of polymerized actin was replaced by some of which are associated with actin cables that into the germ tube or This architecture is of fungi, where actin patches are associated with and cell wall actin cables are for of et al., This actin an actin and in zoospores, and actin patches and cables in sporangia indicates like its cell cycle (Medina et al., 2016), the actin of Spizellomyces displays features that of both animal and fungal Our results actin dynamics and during the chytrid This includes the formation of perinuclear actin shells that are detected by live imaging. that perinuclear actin shells of chytrids were a and our live cell data that these are cellular that likely in many chytrids. perinuclear actin shells are associated with nuclear in animal cells, where they a in nuclear before and after mitosis and and or when through et al., 2016). Although chytrids and other fungi have lost nuclear it likely that Spizellomyces perinuclear actin shells are associated with in nuclear during the cell cycle. we showed that polymerized actin is likely in the formation of zoospore polygonal territories (Figure before the formation of the during cellularization. In

  PublisherDOI

• Chytrid fungi

  Current Biology · 2020 · 38 citations

  Senior authorCorresponding
  • Biology
  • Ecology
  PublisherOA PDFDOI

• Faculty Opinions recommendation of Cell cycle transcriptomics of Capsaspora provides insights into the evolution of cyclin-CDK machinery.

  Faculty Opinions – Post-Publication Peer Review of the Biomedical Literature · 2020-05-26

  dataset1st authorCorresponding
  PublisherDOI

• Faculty Opinions recommendation of Automated continuous evolution of proteins in vivo.

  Faculty Opinions – Post-Publication Peer Review of the Biomedical Literature · 2020-06-05

  dataset1st authorCorresponding
  PublisherDOI

Recent grants

• Measuring and perturbing metabolic rhythms and the cell division cycle in single cells

  NIH · $1.2M · 2018–2024

• Determining mechanisms of rewiring of the eukaryotic cell cycle by a virus without disrupting network function

  NSF · $672k · 2018–2022

• NIH Grant DP2OD00865401

  NIH · $2.4M · 2016

• Determining mechanisms of rewiring of the eukaryotic cell cycle by a virus without disrupting network function

  NSF · $339k · 2018–2019

Frequent coauthors

• Heungwon Park

  Fred Hutch Cancer Center

  24 shared

• Lingchong You

  Duke University

  23 shared

• Shuqiang Huang

  Shenzhen Institutes of Advanced Technology

  23 shared

• Ulrich Gerland

  Technical University of Munich

  21 shared

• Terence Hwa

  21 shared

• Anand Pai

  Chandigarh University

  19 shared

• Yu Tanouchi

  Stanford University

  19 shared

• Yen Ting Lin

  Los Alamos National Laboratory

  11 shared

Education

• Ph.D., Animal Science

  University of California, Davis

  2000

• M.S., Animal Science

  University of California, Davis

  1996

• B.S., Animal Science

  University of California, Davis

  1994

Awards & honors

• NIH Director's New Innovator Award (2011)
• Career Award at the Scientific Interface, Burroughs Wellcome…