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A different shade of hypha : cytological and molecular phylogenetic evidence for the independent rise… Dee, Jaclyn Marie 2011

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A DIFFERENT SHADE OF HYPHA: CYTOLOGICAL AND MOLECULAR PHYLOGENETIC EVIDENCE FOR THE INDEPENDENT RISE OF THE HYPHAL HABIT IN THE CLASS MONOBLEPHARIDOMYCETES (CHYTRIDIOMYCOTA) by Jaclyn Marie Dee B.Sc., The University of British Columbia, 2007  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in The Faculty of Graduate Studies (Botany)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) October 2011  ! Jaclyn Marie Dee, 2011  !  Abstract Once the ancestors of fungi stopped moving and instead started reaching out with hyphae, their filamentous growth made possible new variety in form and habitat. Hyphae mediated substrate colonization, absorptive nutrition, mating and reproduction. Although shared across most familiar terrestrial fungal lineages, little was known about where hyphae evolved in early fungi. In chapter one, I review the phylogenetic origins of hyphae and current understanding of the cytology of hyphal tips. Better understanding of fungal phylogeny and hyphal growth near the base of the fungal tree was needed. In Chapter 2, I investigated the phylogeny and cytology in the Class Monoblepharidomycetes (Chytridiomycota), a group of deeply diverging, zoosporic fungi, encompassing a range of body types. Species can be either crescent or rod-shaped unicells or sprawling hyphal growths. I inferred a phylogeny of the fungi based on 28S ribosomal DNA sequence data using maximum likelihood (ML) and Bayesian inference methods. I recovered the monophyly of modern fungal phyla and the topology was comparable to the most taxonomically diverse and gene–rich phylogeny of the fungi to date. I used likelihood methods to trace the origins of hyphae on my likelihood tree, concluding that hyphae arose independently in the Monoblepharidomycetes and at least three other times in the fungi. Next, I searched for evidence of convergent evolution in the cellular organization of hyphal Monoblepharidomycetes using fluorescence and transmission electron microscopy. I showed that the hyphae of Monoblepharidomycetes have a novel form with an unusual microtubule cytoskeleton and without a typical fungal Spitzenkörper. This constitutes the first report on the cytology of hyphae from the Chytridiomycota. In Chapter 3, I discuss the significance of my research and possible future directions including cytological experiments on the Monoblepharidomycetes cytoskeleton.  !  ""!  Preface The work described in Chapter 2 is the result of my collaboration with my supervisor, Mary Berbee as well as researchers at Arizona State University (ASU) and the University of Maine (U. Maine). I co-conceived of this study with Berbee and captured all of the images of Monoblepharidomycetes with the exception of Fig. 2.1a, which was taken by Marilyn Mollicone (U. Maine) who also isolated some of the monoblepharidalean cultures. Following chemical fixation, I performed all of the immunolocalization and epifluorescence microscopy preparations. I maintained 40 Monoblepharidomycetes cultures, extracted DNA from them, preserved them on fresh slants and cryopreserved them and sent them to the American Type Culture Collection. Following DNA extraction, I carried out DNA amplifications, purifications and sequencing for the vast majority of the isolates, however, Siobhan Wong, an undergraduate research assistant at UBC helped me obtain sequence for 11 DNA fragments. I conducted all the phylogenetic analyses and ancestral state reconstructions, although Berbee provided advice for these analyses. I processed all of the transmission electron microscopic images while Robert W. Roberson (ASU) processed most of the immunofluorescence and epifluorescence light micrographs. Roberson and David Lowry (ASU) also provided training and performed all chemical fixation of fungal materials for microscopic observation. Lowry was responsible for some of the post-fixation procedures for TEM imaging and he did all of the serial sectioning. Joyce Longcore (U. Maine) collected some of the fungal isolates, maintained them in culture and sent them to me in early September 2009. Berbee aided in editing and providing some intellectual input and feedback on this text. ! ! ! ! ! ! ! ! !  """!  Table of Contents Abstract !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!""! Preface!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! """! Table of Contents!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!"#! List of Tables!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! #! List of Figures !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!#"! Acknowledgements !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! #"""! Dedication!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!"$! Chapter 1. Introduction !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!%! 1.1! On the Evolution of Hyphae !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! %! 1.2! Literature Review !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! &! 1.2.1 The Distribution of Hyphae Through the Evolutionary Tree of Fungi $$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$ %! 1.2.2 Hyphae, the Defining Growth Form of Fungi $$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$$ &! 1.3! Research Questions and Hypotheses!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! %'! Chapter 2. A Different Shade of Hypha: Cytological and Molecular Phylogenetic Evidence for the Independent Rise of the Hyphal Habit in the Monoblepharidomycetes (Chytridiomycota)!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! %(! 2.1 Introduction!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! %(! 2.2 Materials and Methods!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! %)! 2.3 Results!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! &&! 2.4 Discussion!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! &)! 2.5 Tables !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! '(! 2.6 Figures!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! '*! Chapter 3. Concluding Chapter!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! *+! 3.1 Outcomes of this Project in Light of Current Research !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! *+! 3.2 Strengths and Limitations of this Research !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! ,'! 3.3 Applications of These Findings and Future Research Directions !!!!!!!!!!!!!!!!!!!!!!!!!!! ,(! References !!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!! ,*!  ! ! ! ! ! ! ! ! ! ! ! ! ! ! !  "#!  !  List of Tables  !  Table 2.1  List of Monoblepharidomycetes specimens sampled in this study…...34  Table 2.2  List of species from AFTOL database used in this study…………….35  #!  List of Figures ! Figure 2.1  Three species in the Class Monoblepharidomycetes possess a range of body forms……………………………………………………………………………..37  Figure 2.2  Hyphal tips of phylogenetically diverse fungi with and without Spitzenkörper viewed under phase contrast or Nomarksi differential interference optics (DIC). …………………………………………………………………………..……......38  Figure 2.3  Phylogenetic distribution of Spitzenkörper in the Kingdom Fungi………....….....39  Figure 2.4  Typical appearance of vegetative hyphae of Gonapodya prolifera under the light microscope. ……………………………………………….……………….….40-41  Figure 2.5  Ultrastructural features of a typical, vegetative Monoblepharis macrandra hypha. …………………………………………………………………………………42-43  Figure 2.6  Nuclei and surrounding organelles in subapical regions of hyphae in Monoblepharis macrandra…………………………………………………………………...…….44  Figure 2.7  Nuclei and surrounding organelles in subapical regions of hyphae in Gonapodya prolifera.……………………………………………………………………..…45-46  Figure 2.8  Mesh-like “reteasomes” surrounded vacuoles near nuclei in Monoblepharis macrandra…………………..…………………………………………….……47-48  Figure 2.9  Gonapodya prolifera hyphae contained eukaryote-typical organelles as well as uncharacterized dense granular bodies. ………………………………….…..49-50  Figure 2.10  Dense granular bodies were the most prominent structure in the cytoplasm of Gonapodya prolifera.………………………………………………………....…..…51  Figure 2.11  Ultrastructure of Gonapodya prolifera pseudosepta. ……………………….…52-53  Figure 2.12  Barbell-shaped nuclei demonstrated that nuclei often squeezed through pores in pseudosepta.……………………...…………………………………………...…..54  Figure 2.13  Ultrastructure at the hyphal apex of Monoblepharis macrandra.…….………..55-56  Figure 2.14  Ultrastructure at the hyphal apex of Gonapodya prolifera..……………..……..57-58  Figure 2.15  Immunolocalization of "-tubulin in Monoblepharis macrandra hyphae revealed astral-like arrays of microtubules flanking nuclei….…………………………...…59  Figure 2.16  Immunolocalization of "-tubulin in hyphae of Coelomomyces stegomyiae demonstrated both canonical cortical arrays of microtubules as well as spindle  !  #"!  microtubules……..…………………………………………………………...…....60 Figure 2.17  Immunolocalization of #-tubulin in Monoblepharis macrandra hyphae.…….…..61-62  Figure 2.18  An ultrathin section through a subapical region in a hypha of Monoblepharis macrandra showing a centriole nucleating an astral-like array of microtubules adjacent to a nucleus. ………………………………………………..………..…..63  Figure 2.19  Actin plaques concentrated at hyphal apices in Gonapodya prolifera.……….……64  Figure 2.20  Actin plaques concentrated at hyphal apices in Gonapodya prolifera but also dotted hyphal length.……………………….……………………………………….......…65  Figure 2.21  Series of images through a hypha of Monoblepharis macrandra taken by optically focusing through the hypha demonstrating actin cable distribution…….…….66-67  Figure 2.22  Actin plaques in the periphery of Gonapodya prolifera hyphae in distal parts of the mycelium.…………………………………………………………………...……...68  Figure 2.23  Phylogeny of the fungi based on 28S rRNA sequence using maximum likelihood and Bayesian inference methods. ……………………………………...….…...69-70  Figure 2.24  Portion of phylogenetic tree from Fig. 3.23 showing the phylogeny of the Monoblepharidomycetes. …………...…………………...……………….……71-72  Figure 2.25  Evolutionary history of hyphae in the fungi inferred by a likelihood model with a single backward and forward rate of transition between the hyphal and nonhyphal forms. ……………………………………………………………….…73-74  Figure 2.26  Evolutionary history of hyphae in the fungi under a likelihood model where the rate if hyphal loss was constrained to 13 times the rate of hyphal gain. …….75-76  Figure 2.27  Pseudosepta evolved only once within the Monoblepharidomycetes in the ancestor of Gonapodya based on parsimony reconstruction of ancestral states …………………………………………………………………………………77-78  ! ! ! ! ! ! !  !  #""!  Acknowledgements Graduate school has been the most challenging and one of the most fulfilling experiences of my life thus far – but it would not have been if not for the many individuals who fed, pushed, prodded, supported and humored me all along the way. Of course, most thanks go to my parents who provided for me for many, many years but just as importantly who taught me to work like a dog, ask questions, and love nature. Profuse thanks go to my supervisor, Dr. Mary Berbee who asked me, “So, Jackie, you should think about grad school. I think you’d like it.” Mary, you’ve been right quite often since then. Thank you for understanding my outside commitments while also being an incredible leader, mentor and purveyor of pizza-flavoured calories on long nights in the lab. Thanks for guilting me into doing things that were way out of my comfort zone, I learned a lot and I’ve become more confident. Special thanks also go to Mary for working overtime to help me get this thesis done on time. I’d like to thank my collaborators at Arizona State University: Dr. Robert W. Roberson and David Lowry. Robby, thanks for being a great teacher, for helping me with the images and getting me out of the lab on occasion. Thanks, Dave for being patient with me and doing all that sectioning so quickly! I’m also incredibly grateful to Essie Wilson (ASU) who treated me like family while I was there. I thank my committee members and exterminal examiner, Drs. Naomi Fast, Martin Adamson, Jeannette Whitton and Gary Bradfield who gave up their most precious resource, their time, to discuss my research project development and witness its conclusion. I’d also like to thank Shona Ellis, my personal hero in the field of education who gave me my first TA job and has given me many opportunities in science outreach. I’ve deeply enjoyed teaching over the past few years and look forward to sharing my enthusiasm for biology for a long time. A thousand thanks also go to Dr. Robin Young for teaching me how to process my images. You’re the best, Robin! In addition, I thank Drs. Lacey Samuels, Geoff Wasteneys and Sean Graham who took the time to chat and advise me on my research. Many thanks also go to the Botany office staff for enduring an endless onslaught of financial, program and scholarship application questions. Thanks to my friends and other family members for listening and for forgiving me when I missed things because I was busy doing science. Thanks to my fellow Botany grad students who were plain old fun to bump into everyday. If I’ve forgotten someone here, I apologize. This is a very stressful time and I’m extremely grateful for all your help and support.  !  #"""!  Dedication  To Mom and Dad  ! ! ! ! !  !  "'!  Chapter 1. Introduction 1.1  On the Evolution of Hyphae It is possible that life as we know it could not exist if it were not for the hypha. Most  fungi, whether mould from a piece of fruit and or a grocery store mushroom, are made up of microscopic threads. These threads are the hyphae, the multinucleate, filamentous cells contained in a chitinous wall that make up fungal bodies. A network of fungal hyphae is called a mycelium. Hyphae acquire nutrients, colonize substrates and form spores and gametes for reproduction (Alexopoulos et al., 1996). Hyphae can take various forms by branching, fusing with one another, and can grow parallel and in close association with one another or individually spread out across a substrate (Harris, 2011). These humble but endlessly versatile cells may have given plants a huge “leg-up” in the colonization of land. Hyphae encased in the cells of fossilized early land plants tell the story of how the earliest land plants might have invaded terrestrial environments. Creeping, exploratory hyphae extending from inside plant cells presumably gave their hosts access to a wealth of nutrients before plants had true, nutrient absorbing roots (Pirozynski and Malloch, 1975; Remy et al., 1994). Billions of years later, the importance of hyphae in our lives remains inestimable. Fungal hyphae cycle nutrients in the environment and help trees grow. Hyphae can make us sick, but they can also produce compounds that heal. But fungi did not always make hyphae. As fungi share a common ancestor with the animals and the collar-flagellates as well as a slime mold and other crawling amoebae, their most ancient predecessors were most likely unicellular, uniflagellate amoebae (Baldauf and Palmer, 1993; Brown et al., 2009; Liu et al., 2009; Medina et al., 2003; Ruiz-Trillo et al., 2004; Steenkamp et al., 2006). Moreover, not all extant fungi exhibit the hyphal habit. This observation raises numerous questions about the evolution of hyphae. In the course of my thesis work, I intended to trace the evolution of hyphae in the fungi and in doing so answer the question: Where in the fungal tree did hyphae originate? Specifying where hyphae arose is a formidable task that will benefit greatly from two kinds of analysis: 1) a phylogenetic analysis of !  (!  early-diverging hyphal and unicellular fungi and 2) critical study of hyphal cytology to evaluate underlying patterns of homology across representative taxa.  1.2  Literature Review  1.2.1 The Distribution of Hyphae Through the Evolutionary Tree of Fungi To study the evolution of hyphae, I needed to know how hyphal fungi are related to one another as well as to other non-hyphal fungi. The phylogeny of the fungi is not fully resolved and therefore interpreting the origins of the filamentous form in the fungi is problematic. In discussing fungal phylogeny and classification, I will follow the scheme proposed by Hibbett and co-workers (2007) summarizing the results of recent studies. For a generalized fungal tree showing phyla with names of representative hyphal genera, see Ch. 2, Fig. 2.3, p. 41. The hyphal form dominates in terrestrial fungal groups: the Ascomycota, Basidiomycota, Glomeromycota and zygomycetous fungi. James et al. (2006a) sampled a set of six genes from 199 fungal species to produce the most taxon and gene-rich phylogeny of the fungi to date. As in other phylogenies using fewer genes (Liu et al., 2006), different genes (Steenkamp et al., 2006), genomic data (Fitzpatrick et al., 2006), mitochondrial genomes (Seif et al., 2005) or that sampled more heavily from certain fungal lineages (White et al., 2006), this work supported the sister group relationship between the Ascomycota (sac fungi) and Basidiomycota (club fungi). Together the two phyla form a clade often referred to as the Dikarya (Hibbett et al., 2007). The Dikarya consist mainly of hyphal fungi, some of which gave rise to unicellular yeasts (Harris, 2011). The Dikarya in turn was the sister group to the Glomeromycota, also hyphal, containing the arbuscular mycorrhizae (James et al., 2006a; Liu et al., 2006; White et al., 2006). Filamentous forms are also common in the zygomycetous fungi whose phylogeny is uncertain (Tanabe et al., 2004). Numerous molecular phylogenies have indicated that the Phylum Zygomycota is not a natural group (Bruns et al., 1992; James et al., 2006a; James et al., 2000; Keeling et al., 2000; Schüβler et al., 2001; Tanabe et al., 2004; White et al., 2006). Zygomycetous fungi have also appeared to be monophyletic in other phylogenies (Liu et al., 2006).  !  %!  In contrast, hyphal forms are much less common in the remaining fungal groups the chytrids, microsporidia and the cryptomycota, which are thought to have arisen from the most basal nodes. Resolving the phylogeny of these fungi is a hurdle on the path to elucidating the origins of hyphae. The majority of the aquatic zoosporic fungi, also known as chytrids, do not produce hyphae. Zoospores are uniflagellate, wall-less cells used in reproduction and dispersal. All zoosporic fungi were traditionally placed in the Phylum Chytridiomycota and then further classified based on their zoospore ultrastructure (James et al., 2000). However, molecular phylogenies offer compelling evidence that chytrids are paraphyletic (James et al., 2006b; James et al., 2000; White et al., 2006); Sekimoto et al. manuscript under review). While the majority of known chytrids belong to Chytridiomycota, others belong to the cryptomycota, the hyphal Blastocladiomycota and one genus of unicellular chytrids, Olpidium, belongs among the zygomycetous fungi (James et al. 2006b, Sekimoto et al. manuscript under review). The “hidden fungi”, the uniflagellate cryptomycota (Jones et al., 2011), also known as LKM11 (Lara et al., 2009; van Hannen et al., 1999) or Rozellida from environmental sequence data (Lara et al., 2009), are important to the evolutionary story because they appear to have arisen from the deepest split in the fungal tree (James et al., 2006a; Jones et al., 2011). Jones et al. (2011) observed unicellular, wall-less, flagellated and nonflagellated cryptomycotan cells either free or attached to algae and proposed a general life cycle for these fungi, but little is known about their morphology and life history. The only well-characterized member of this clade at the moment is Rozella, which has a unicellular body and is an intracellular parasite of fungi and fungus-like algae (Alexopoulos et al., 1996). Rozella allomycis infects the chytrid, Allomyces and produces zoosporangia, zoospores and resistant sporangia inside of its host (Held, 1980). Whether the biology of Rozella is truly representative of the cryptomycota is currently unknown. The rise of the obligate intracellular pathogens, the unicellular microsporidia, was formerly thought to pre-date the acquisition of mitochondria by early eukaryotes (Vossbrinck et al., 1987). Later phylogenies uncovered their close affinity to the fungi (Edlind et al., 1996; Hirt et al., 1999; Keeling and Doolittle, 1996) and in particular to the zygomycetes (Keeling, 2003; Lee et al., 2008). James et al. (2006a) found that the Microsporidia were the sister group to Rozella at the base of the fungi, although this could be the result of long-branch attraction artifact. !  )!  Phylum Chytridiomycota and origin of hyphae Phylogenies show that the divergence of the chytrids from other fungi followed after the origin of the stem lineage of the unicellular cryptomycota and microsporidia (James et al., 2006a; Jones et al., 2011). Therefore, a version of the hyphal form must have originated in the Chytridiomycota (Stajich et al., 2009) and knowledge of hyphae in representative chytrids is essential for understanding hyphal biology and evolution. Many Chytridiomycota produce unicellular bodies supported by a network of rhizoids. The unicellular bodies develop into sporangia, eventually releasing zoospores. Rhizoids are different from hyphae because they lack nuclei and they grow progressively narrower as they branch (Alexopoulos et al., 1996). Other chytridiomycotan fungi have true hyphae, including Polychytrium aggregatum (Ajello, 1948) and several members of the Class Monoblepharidomycetes (Alexopoulos et al., 1996; Hibbett et al., 2007). Class Monoblepharidomycetes, phylum Chytridiomycota For this study, I have focused on the Monoblepharidomycetes because members of its six known genera demonstrate a wide range of morphologies (Alexopoulos et al., 1996; Bullerwell et al., 2003; Ustinova et al., 2000). While the placement of the Monoblepharidomycetes in the fungal tree is uncertain, most studies indicate that they diverged early within the Chytridiomycota (Bullerwell et al., 2003; James et al., 2006a; White et al., 2006). The unicellular genus Hyaloraphidium produces crescent-shaped cells and lacks a zoospore stage but reproduces by autosporulation (Ustinova et al., 2000). Other genera reproduce by zoospores (Alexopoulos et al., 1996). Two genera, Harpochytrium and Oedogoniomyces form rod-shaped cells (Gauriloff et al., 1980a), and three genera, Monoblepharis, Monoblepharella and Gonapodya, are hyphal. The variety of forms in the group makes it wellsuited for a case study in hyphal evolution. I wish to draw comparisons between the growth and cytology of hyphae in Monoblepharidomycetes and other fungi. Studies to date on Monoblepharidomycetes have been few in number. Travland and Whisler (1971) investigated the ultrastructure of vegetative cells and of zoospores of Harpochytrium hedinii. Ustinova et al. (2000) studied the fine structure of Hyaloraphidium curvatum. Earlier light microscopic studies showed that, uniquely among Fungi, Monoblepharidomycetes reproduce sexually by means of flagellate sperm and stationary eggs !  *!  (Alexopoulos et al., 1996; Johns and Benjamin, 1954; Perrott, 1958; Sparrow, 1953). Later, Gauriloff et al. (1980b) and Mollicone and Longcore (1994; 1999) examined zoospore ultrastructural characters that were important in chytrid classification (Barr, 1990; James et al., 2000).  1.2.2 Hyphae, the Defining Growth Form of Fungi Knowledge about the biology of hyphae has defined mainly by studies of cytology and of hyphal function in the model systems organisms in the Ascomycota and Basidiomycota (Harris, 2011). However, fungal phylogenetic diversity is concentrated among the early-diverging zoosporic fungi where almost nothing is known about hyphal growth. In my thesis, I interpret the structures underlying hyphal growth in Monoblepharidomycetes in the light of what is currently known about structures related to filamentous growth and development in wellstudied fungi. I will next summarize the results of many studies of the mechanisms underlying hyphal growth. How Hyphae Grow and Take Shape: Apical Growth and the Spitzenkörper Fungal hyphae grow at their tips. Admittedly, they also branch, they sometimes form septa, and some fungal hyphae undergo intercalary growth (Christensen et al., 2008; Read, 2011). However, the most commonly studied, readily observable and most significant form of hyphal growth is tip-mediated growth (Read, 2011). Since my study represents the first close examination of hyphae in a chytridiomycotan fungus, I focused my study on the hyphal tip and the structures underlying polarized growth. In the mid-20th century, Girbardt (1957) first observed the close association between the movement of the Spitzenkörper, a phase-dark “tip body” (Brunswik, 1924) and the directionality of apical growth in fungi. This influential work spawned numerous studies that have uncovered the dynamic nature and ultrastructural equivalents of this phase-dark region and pursued the mechanism by which it defines the shape of hyphae. Early transmission electron microscopic (TEM) images of hyphal tips suggested that the Spitzenkörper was composed of an accumulation of secretory vesicles (Girbardt, 1969; !  &!  McClure et al., 1968). Girbardt recognized two types of Spitzenkörper, type 1 and type 2. Type 1 was a cap-like organization of secretory vesicles, now called a “crescent-shaped” organization of vesicles. Type 2 was the spheroid aggregation (Girbardt, 1969). Grove and Bracker (1970) examined hyphal ultrastructure in diverse fungi, including the oomycete, Pythium ultimum and defined three forms of apical tip organization, the third belonging to the regularlyseptate, Ascomycota and Basidiomycota. Following Girbardt, they labelled the cluster of vesicles in the third type of hyphal apex as the Spitzenkörper. Grove and Bracker’s (1970) study further identified and compared Spitzenkörper revealing that while the basic components of Spitzenkörper were the same in many fungi, they were arranged in a variety of ways. All Spitzenkörper contained apical vesicles (70-90 nm diam.), microvesicles (30 nm diam.) and microfilaments but they were organized in unique ways in different fungal species. For instance, Neurospora crassa Spitzenkörper contained a core region of ribosomes and microvesicles in a cluster of larger, electron-transparent apical vesicles. Conversely, the core region of the Spitzenkörper in Fusarium lacked microvesicles but had a posterior cluster of ribosomes and some tubular structures. Following a pioneering study by Howard and Aist (1979), ultra-rapid freezing of cells has been leading to close-to-life preservation of cellular organization in mostly asco- and basidiomycetous fungi (Hoch and Howard, 1980; Hoch and Staples, 1983; Howard, 1981; Howard and Aist, 1979; Newhouse et al., 1983; Roberson and Fuller, 1988). Unlike the conventional chemically fixed material, the quality of images from freeze-substituted fungi was stunning, free from artifacts such as wavy membranes and revealed fungal tip ultrastructure in incredibly fine detail. Howard and Aist (1979) compared TEM images from conventional chemical and freeze-substituted material from a single species, Fusarium acuminatum. Where chemical fixation had revealed oblong apical vesicles and indistinct microvesicles, freezing showed different types of spherical apical vesicles and hexagonal microvesicles. Later, Howard (1981) demonstrated the presence of microvesicles and microfilaments in the phase-light core region of F. acuminatum where they had not previously been observed in chemically fixed material (Grove and Bracker, 1970). He also described an unrecognized class of vesicle coated in microfilaments, which he named the “filasome”. Later, Bourett and Howard (Bourett and Howard, 1991) and Roberson (1992) showed with gold-labelling that the microfilaments surrounding filasomes were made of actin. Moreover, ribosomes frozen on the spot have also !  +!  been noted around the Spitzenkörper (Roberson et al., 2011; Vargas et al., 1993) or in a cluster just behind it (Howard, 1981; Howard and Aist, 1979). To extend knowledge about Spitzenkörper outside of model organisms and beyond the Dikarya, Roberson and colleagues surveyed the hyphal tips of chytrids and zygomycetes. Prior to Roberson’s work, the only study of chytrid hyphal tips was that of Roos and Turian (Roos and Turian, 1977) who proposed a fourth type of apical region, in addition to Grove and Bracker’s (1970) three forms, at the hyphal tip of Allomyces arbuscula. They based their conclusions on light micrographs of living material and TEM images of chemically fixed material. In A. arbuscula, the apex contained mainly vesicles and non-specific membranous cisternae in a roughly crescent-shaped organization and microtubules in a radial array. Taking another look at the genus, Vargas et al. (1993) found that the Spitzenkörper in Allomyces macrogynus appeared as a phase-light spherical region in the light microscope. TEM imaging showed that this region corresponded to a region of granular cytoplasm and some sparse actin microfilaments but no microvesicles (diam < 40 nm). Vargas et al. (1993) and later Srinivasan et al. (1996) found that the phase-light region of A. macrogynus was not surrounded by a conspicuous vesicle cloud. Recently, Roberson and others (2011) described the first Spitzenkörper in a zygomycete fungus, Basidiobolus ranarum, as an undifferentiated phase-dark spheroid body without apical microvesicles and without a differentiated core region. The B. ranarum Spitzenkörper appeared simply as an aggregation of large vesicles. Confirming the complex and highly variable nature of the Spitzenkörper, López-Franco and Bracker (1996) made video-recordings of living hyphae which revealed that Spitzenkörper do not have a defined boundary and are constantly moving and being remodeled. They defined eight unique forms of Spitzenkörper based on phase-contrast imaging of 32 different fungi that differed in the shape, position and presence of phase-light and dark regions. They also referred to a ninth unique configuration from Allomyces macrogynus (Roberson et al., 2011; Srinivasan et al., 1996; Vargas et al., 1993). In line with their curiosity regarding the structural mysteries of the Spitzenkörper, investigators have sought to uncover its role in hyphal tip elongation. Based on early TEM images of vesicles fused with the plasma membrane at hyphal apices, McClure et al. (1968) proposed that tip vesicles, assumed to be derived from the Golgi, added membrane and cell !  ,!  wall building materials to the expanding hyphal apex. Girbardt (1969) also suggested that apical vesicles were involved in routine secretion of extracellular enzymes. Taking these ideas into account, Bartnicki-Garcia et al. (1989) proposed the mathematical Vesicle Supply Center (VSC) model for hyphal extension. Under this model, the Spitzenkörper is propelled forward as a unit and as it moves, it releases vesicles received from the Golgi thus giving the hypha its characteristic shape. Since then, Gierz and Bartnicki-Garcia (2001) have articulated the VSC model to account for the three-dimensional structure of the hypha. Supporting the role of the Spitzenkörper as a VSC, workers have detected chitin synthases, which contribute to chitin cell wall synthesis, in discrete vesicles in the region of the Spitzenkörper. Verdín et al. (2009) discovered that vesicles containing certain enzymes formed functional layers in Neurospora crassa leading to the identification of an outer ring, termed the “Spitzenring”, and a central core within the Spitzenkörper. Sánchez-Léon et al. (2011) found that different classes of chitin building enzymes, the chitin synthases are held in different subpopulations of vesicles. Suggesting that its roles may vary across fungi, the Spitzenkörper in Allomyces may act as a microtubule-organizing center (MTOC). McDaniel and Roberson (1998) detected #-tubulin, the protein that nucleates microtubule formation, in the region corresponding to the Spitzenkörper in Allomyces. Thus far, #-tubulin has not been observed in hyphal apices in any other fungus though microtubules are frequently found in the apical region of hyphae (Harris et al., 2005) RW Roberson and RR Mouriño-Pérez unpublished). The Role of the Cytoskeleton in Hyphal Extension So far, I have emphasized the relationship of the Spitzenkörper to tip growth. The next questions concern how vesicles move to the tip. At least part of the answer lies in the cytoskeleton, which forms tracks for the motor proteins that transport vesicles to the hyphal tip (Schuchardt et al., 2005; Seiler et al., 1997). Besides being directly involved in polar transport, the cytoskeleton can generate a physical force that can push cytoplasm out into growing tips (Heath and Steinberg, 1999). The polarized orientation of actin and microtubules in the hypha is observable under the TEM and by using the actin-binding toxin, phalloidin or immunofluorescence microscopy (Bourett and Howard, 1991; Howard, 1981; Roberson, 1992; Roberson et al., 2011). The roles of microtubules and actin have also been studied in some detail using chemicals that affect the stability of these proteins (Taheri-Talesh et al., 2008; Torralba et al., 1998). Analyses of loss of function mutants (Virag and Griffiths, 2004), of !  -!  fluorescently tagged actin (Berepiki et al., 2010) and tubulin (Finley and Berman, 2005; Horio and Oakley, 2005; Uchida et al., 2008), have revealed the cytological locations and the functions of these structures in certain model fungi. Actin Polarized growth is impossible without actin. When an actin polymerization inhibitor such as a cytochalasin is imposed on a hypha, the hyphal tip loses its polarity and balloons instead of forming a tube (Heath et al., 2000; Srinivasan et al., 1996; Torralba et al., 1998). Furthermore, Virag and Griffths (2004) found that N. crassa mutants deficient in actin activity branched profusely instead of growing continuously at their apex. Differentiated roles in polarized growth have been linked to specific forms of actin. Different forms are generated through the binding of actin microfilaments with actin binding proteins (Walker and Garrill, 2006). The most conspicuous of these forms are actin patches, also called plaques. Actin patches are concentrated in the hyphal apices of several fungi. Specifically, they form a cortical collar in the subapical region of the hyphae of Neurospora crassa (Berepiki et al., 2010; Delgado-Álvarez et al., 2010), Aspergillus nidulans (Taheri-Talesh et al., 2008; Upadhyay and Shaw, 2008) and Sclerotium rolfsii (Roberson, 1992) where they are thought to mediate endocytosis. Contrary to intuition and early experiments with membrane dyes (Girbardt, 1969), endocytosis occurs at hyphal tips and actin patches are believed to represent sites of endocytosis. Removing parts of the plasma membrane during hyphal growth seems counterproductive (Shaw et al., 2011). Consider, however, that if vesicles are continually added to the hyphal apex, the part of the membrane that was once at the very apex necessarily moves downward. Therefore, anything embedded in the plasma membrane that indicates to the cell machinery where materials for growth should be targeted would be continually edging down the hypha rather than remaining at tip (Shaw et al., 2011). As a consequence, polarity would be lost. Since hyphae exhibit sustained polarized growth, they must have a means of keeping polarity-signaling equipment at the apex. In response to this problem, numerous researchers formed their own very similar ‘Apical Recycling Models’ (Fischer-Parton et al., 2000; Higuchi et al., 2009; Steinberg, 2007; Taheri-Talesh et al., 2008; Upadhyay and Shaw, 2008). Briefly, actin patches are meshworks of actin filaments that surround clathrin in the membrane regions flanking the hyphal apex. Motor proteins acting on the actin filaments pull the plasma membrane inwards to form an endocytic vesicle. These vesicles fuse with the endosome. Reports conflict !  .!  as to what happens next. Vesicles may then bud off the endosome and fuse with the Golgi (Taheri-Talesh et al., 2008), they might travel directly to the Spitzenkörper (Fischer-Parton et al., 2000) or they may do both. At any rate, vesicles end up at the Spitzenkörper and are shuttled back to the hyphal tip (Fischer-Parton et al., 2000; Higuchi et al., 2009; Taheri-Talesh et al., 2008; Upadhyay and Shaw, 2008). Cortical actin cables also form a network throughout the hypha (Roberson, 1992). The role of actin cables in intracellular transport in hyphal fungi has not been directly determined. However, in the budding yeast, Saccharomyces cerevisiae actin cables account for the overall polarity of the cell and are the primary transportational highway for vesicle transport (Pruyne and Bretscher, 2000). Actin cables can also move actin plaques (Taheri-Talesh et al., 2008). Note that elaborate networks of actin cables are easily observed in some fungi (Roberson, 1992; Taheri-Talesh et al., 2008) but are sparser or absent in others (Berepiki et al., 2010; Horio, 2007; Srinivasan et al., 1996). The implication of this is that fungi with fewer actin cables must rely more heavily on microtubule-based transport (Horio, 2007). Actin can also be found as individual microfilaments in the Spitzenkörper. Interestingly, they have not been seen in the Spitzenkörper using dyes or antibodies (Roberson, 1992), whereas TEM images (Bourett and Howard, 1991; Howard, 1981; Roberson and Fuller, 1988) and GFP fusions of actin and ABPs, have clearly demonstrated their presence there (TaheriTalesh et al., 2008). Myosin motors travelling along these microfilaments are thought to carry the vesicles that comprise the Spitzenkörper and shuttle them to the apical plasma membrane (Steinberg, 2007). Microtubules While actin is absolutely critical in polarized growth, microtubules seem to play a supplementary role. Extensive linear arrays of microtubule ‘highways’ in fungal hyphae often extend into the Spitzenkörper region (Freitag et al., 2004; Howard, 1981; McDaniel and Roberson, 1998; Roberson and Fuller, 1988; Roberson et al., 2011; Uchida et al., 2008). These are thought to be the highway along which vesicles are trafficked over long distances in the growing hypha (Fuchs et al., 2005; Steinberg, 2007). Hyphae can grow from spores without forming microtubular arrays, however, disruption of the microtubular cytoskeleton leads to crippled growth rates in mature hyphae (Horio and Oakley, 2005). And, if microtubules are the !  (/!  highway, plus-end directed kinesin motors are the trucks hauling materials for secretion and tip growth. When kinesin motor function in N. crassa is lost, the Spitzenkörper does not form (Seiler et al., 1997). In addition, kinesins carry mRNAs from subapical nuclei towards ribosomal clusters near the Spitzenkörper along microtubule tracks (Becht et al., 2006). In addition to their role in polarized hyphal growth, microtubules are also involved in the process of mitosis. Gambino and colleagues (1984) compared microtubule organization in interphase and during mitosis in Aspergillus nidulans protoplasts. During interphase, microtubules were organized in elaborate cytoplasmic arrays but during mitosis, microtubules were largely found in the spindle. Confirming this relationship in living cells, Szewczyk and Oakley (2011) examined microtubule dynamics in real-time in A. nidulans hyphae, observing the extremely rapid rearrangement of microtubules right before and after spindle formation. Surprisingly, this drastic microtubular renovation impeded neither growth nor the formation of Spitzenkörper (Pruyne et al., 2002; Riquelme et al., 2003). Later, examination of Green Fluorescent Protein-(GFP)-tubulin in A. nidulans demonstrated that while much of the microtubule cytoskeleton became involved in spindle function during mitosis, apically located microtubules remained in place, presumably facilitating continued growth (Horio and Oakley, 2005). Thus, while microtubules function in both growth and mitosis, different microtubule populations have unique roles in A. nidulans. Other Structures and Forces Implicated in Hyphal Growth As discussed, investigations into the mysteries of apical growth have been largely concerned with the Spitzenkörper, but the Spitzenkörper does not act alone. Studies of the genetic basis of polarization in, S. cerevisiae have shed light on the other components that control apical growth in fungi. I will briefly mention some of the highlights to provide a more comprehensive view of the mechanism of tip growth. Two protein complexes involved in polarity, the polarisome and the exocyst, which are distinct from the Spitzenkörper, have been well characterized in S. cerevisiae. The polarisome is comprised of proteins that facilitate actin filament assembly at the tip (Pruyne et al., 2002). Myosin motors ferry vesicles to the growing end of the yeast (He et al., 2007; Pruyne et al., 2002). Once they reach the budding site, vesicles dock at the exocyst then fuse with the plasma membrane (He et al., 2007). Jones and Sudbery (2010) localized Spitzenkörper proteins as well !  ((!  as polarisome and exocyst protein orthologs in the facultatively hyphal pathogen, Candida albicans. They consistently found exocyst and polarisome components in a crescent-shape at the hyphal tip, the site of growth. The Spitzenkörper was a separate entity and materials flowed steadily to and from it. From these observations, Jones and Sudbery (Jones and Sudbery, 2010) suggested a model in which vesicles from the Spitzenkörper arrived at the exocyst via the actin microfilaments that were nucleated by the polarisome. Other workers have made similar observations in Aspergillus (Sharpless and Harris, 2002) and Ashbya gossypii (Köhli et al., 2008). Furthermore, turgor pressure as long been believed to be responsible for tip growth (Lew, 2011) though there is some evidence to the contrary from the oomycete water moulds (Money and Harold, 1993). Water moulds were grown in hyperosmotic conditions then their internal osmotic pressure was measured. Because individuals exhibiting low osmotic pressure (low turgor) continued to grow, some other force must be mediating growth and it is likely the cytoskeleton, as mentioned above (Heath and Steinberg, 1999). Finally, a calcium gradient created internally at the hyphal tip is also thought to regulate hyphal growth, at least in N. crassa. Calcium was highly concentrated at the tips of actively growing hyphae whereas non-growing hyphal tips did not demonstrate this calcium gradient (Levina et al., 1995). Though the mechanism by which this gradient affects growth has yet to be determined, Silverman-Gavrila and Lew (2003) posited that the many proteins that are regulated by calcium were likely required for tip growth. Most hyphae produce crosswalls called septa Many fungi bear regularly spaced septa, cross-walls composed of cell wall components that appear to compartmentalize the hypha but permit the flow of organelles such as nuclei through septal pores (Giesy and Day, 1965; Hunsley and Gooday, 1974). Because materials can flow through pores, septa do not seem to function strictly in cellularization, though they can become plugged to seal off parts of the hypha with different functions (Beckett, 1981). Septa are also sites of wound healing following injury in ascomycetes and basidiomycetes (Aylmore et al., 1984; Buller, 1933; Collinge and Markham, 1985). Interestingly, exposing hyphae to low nutrient media, high temperature, or hypertonic solutions caused plugging or unplugging of septal pores near the hyphal apex suggesting that septa help some fungi survive stressful  !  (%!  conditions (van Peer et al., 2009). Septa might also add mechanical strength to the hypha (Gull, 1978).  1.3  Research Questions and Hypotheses Against this background, I identified lines of inquiry that address the central question of  where hyphae arose in the fungal tree. I had two alternative hypotheses. Hyphae either evolved convergently in the Chytridiomycota and in the Dikarya/zygomycete clade, or they evolved once, early in fungal history and their absence even among unicellular Chytridiomycota represents convergent loss. Addressing these hypotheses required ancestral state reconstruction and a phylogenetic tree that included Chytridiomycota with hyphal and unicellular bodies. To contribute to the sampling of Chytridiomycota with diverse body forms, I sequenced ribosomal genes from isolates of Monoblepharidomycetes. I also asked the question: How do hyphae in the Monoblepharidomycetes (Chytridiomycota) compare to those of other fungi? Because Monoblepharidomycetes' cellular organization and ultrastructural features were largely unknown, I planned to investigate their hyphal apex ultrastructure, cytoskeletal architecture and organellar composition. This involved analyzing the Spitzenkörper and cytoskeletal structure in the hyphae of Monoblepharis, Gonapodya and Monoblepharella using light and electron microscopy. These investigations would provide evidence for analysis of convergence and homology in hyphal construction across the most divergent members of kingdom Fungi.  !  ()!  Chapter 2. A Different Shade of Hypha: Cytological and Molecular Phylogenetic Evidence for the Independent Rise of the Hyphal Habit in the Monoblepharidomycetes (Chytridiomycota) 2.1 Introduction While the common ancestors of fungi and animals were swimming and crawling unicells, modern fungi exhibit a striking range of sizes both gigantic and microscopic (Liu et al., 2009; Smith et al., 1992; Steenkamp et al., 2006). Moreover, fungi are found in an array of habitats from rotting fruit in the fridge to more macabre places such as ant brains (Evans et al., 2011). The vast majority of the known morphological and ecological diversity of fungi is largely attributable to the rise of hyphae. Hyphae, the long, filamentous cells, enveloped in chitinous cell walls are characteristic of most known fungi. Members of the Ascomycota, the largely hyphal sac fungi, make up approximately 64% of Earth’s known fungal diversity (Stajich et al., 2009). Not all fungi are hyphal though. Non-hyphal fungi populate branch tips all over the fungal tree of life. Single-celled fungi, also known as yeasts, are found in the terrestrial Ascomycota, Basidiomycota, and zygomycetous fungi. Another non-filamentous group, the chytrids, are aquatic fungi that reproduce and disperse via a uniflagellate cell known as the zoospore. Species of chytrids from the Chytridiomycota, as well as Rozella and its close relatives, the cryptomycota, which are typically unicellular, arose from the deepest divergences in the Kingdom Fungi (James et al., 2006a; Jones et al., 2011). Other chytrid species, including some hyphal ones belong to the Blastocladiomycota and even the terrestrial, aseptate, zygomycetous lineages (James et al., 2006b). Given this distribution of unicellular and hyphal forms in the fungal tree, I became curious about the origins of hyphae in the fungi. Exploring the hyphal form in a group of early-diverging chytridiomycotan fungi thus became my goal. Therefore, I focused my research on the phylogeny and cellular organization of hyphae in the Class Monoblepharidomycetes. Monoblepharidomycetes is a monophyletic chytrid group that belongs to the Chytridiomycota (Bullerwell et al., 2003; James et al., 2006a; James et al., 2006b; White et al., 2006) under most phylogenetic reconstructions but could also have sprung from the basal divergence in the fungi (Sekimoto et al. manuscript under review). While hyphal forms are rare in the Chytridiomycota, several members of !  (*!  Monoblepharidomycetes are filamentous (Fig. 2.1). Others form crescent or rod-shaped unicells (Fig. 2.1) (Alexopoulos et al., 1996; Forget et al., 2002; Ustinova et al., 2000). However, no known species exhibit the simple thallus form comprised of a single zoosporangium tethered by rhizoids possessed by closely related chytrids such as the infamous amphibian pathogen, Batrachochytrium dendrobatidis (James et al., 2006a; Longcore et al., 1999). The hyphae of Monoblepharidomycetes are distinguishable from the rhizoids that simply anchor other chytrids to their substrate. Rhizoids do not contain nuclei and become finer as they branch (Alexopoulos et al., 1996). I wondered whether monoblepharidalean hyphae had a shared beginning with other fungal hyphae in the most recent common ancestor of the fungi or if they were uniquely derived. I aimed to answer these questions using a combination of molecular phylogenetics and microscopy. Our current understanding of the evolution of hyphae was recently summarized by Harris (2011) who focused on lineage specific variations on the hyphal form in Ascomycota, Basidiomycota and the zygomycetous fungi. I inferred a phylogeny of the fungi based on the 28S rDNA region using maximum likelihood (ML) analyses and Bayesian Inference. I sampled widely from the Kingdom Fungi but emphasized taxon-sampling within the Monoblepharidomycetes in an effort to better infer their position in the fungi as well as resolve intergeneric relationships. An ancestral state reconstruction based on the resulting phylogeny was used to explain the origins of these hyphae and the presence of pseudosepta, irregularly spaced cell wall thickenings in the genus Gonapodya. I then examined monoblepharidalean hyphae for similarities and differences with other fungal hyphae. Learning more about the biology and evolutionary beginnings of these hyphae could give us a glimpse of early hyphal fungi. I then sought to unveil the structures driving hyphal growth in the Monoblepharidomycetes. Hyphae grow at their tips, so, as a starting point, I scoured hyphal tips for a Spitzenkörper, or “tip body” (Girbardt, 1969). The Spitzenkörper is a discrete, often spherical body, visible at the light microscope level that acts as a ‘vesicle supply center’ that accumulates and guides vesicles containing materials for cell wall synthesis and extracellular enzymes from the Golgi to the growing tip (Bartnicki-García et al., 1989). Spitzenkörper are highly dynamic (López-Franco and Bracker, 1996) and vary in form across species (Fig. 2.2). Ultrastructurally the Spitzenkörper appears to be a cluster of vesicles with differing !  (&!  characteristics interacting with the cytoskeleton and sometimes containing ribosomes (Grove and Bracker, 1970; Harris et al., 2005; Roberson and Fuller, 1988; Roberson et al., 2011; Vargas et al., 1993). While Spitzenkörper are widely distributed in the fungal tree, some fungi do not appear to possess them (Fig. 2.3). To properly capture the accumulation of vesicles at the hyphal apex and the fine structure of the microtubule and actin cytoskeletons, I used rapid freezing and freezesubstitution methods. Past studies made careful and even taxon-extensive observations of cellular organization in hyphae using traditional chemical fixation (Girbardt, 1969; Grove and Bracker, 1970; McClure et al., 1968). Unfortunately, the slow penetration of chemical fixatives killed cells slowly leading to artifacts and reports of false organelles (Chandler and Roberson, 2009). Ultrarapid freezing techniques have improved the quality of preservation by leaps and bounds, producing stunning images of organelles frozen on the spot (Howard, 1981; Howard and Aist, 1979; Roberson and Fuller, 1988; Roberson et al., 2011; Vargas et al., 1993). In this thesis chapter, I will use phylogenetic analysis including all available cultures of the key basal fungi in the Monoblepharidomycetes to evaluate where hyphae evolved in the fungal tree. I will apply cytological approaches to examine hyphae of the Monoblepharidomycetes for the features that make these fungal hyphae distinctive. I will discuss the general characteristics, organellar composition, hyphal tip ultrastructure and cytoskeletal organization of monoblepharidalean hyphae. Together, these approaches will enable me to reconstruct patterns of early evolution of hyphae in the fungi.  2.2 Materials and Methods Cultures I received 39 Monoblepharidomycetes from Joyce Longcore (U. Maine) that were originally isolated by Longcore, Marilyn Mollicone (U. Maine) and Howard Whisler (U. Washington) from various algae, fruit and twig baits. One isolate, CR90 came from the Berkeley Microgarden (UC Berkeley). The 39 isolates represent the most complete set of cultures of Monoblepharidomycetes available in the world. All mycelial isolates were maintained at approximately 20°C on mPmTG agar (0.4 g peptonized milk, 0.4 g tryptone, 2.0 g glucose, and 5.0 g agar in 1L distilled water) and transferred every three months. Rod-shaped isolates, !  (+!  Harpochytrium (JEL 94, JEL 105, JEL 196) and Oedogoniomyces (CR90) were maintained by adding liquid media to the solid agar culture once per week and transferring colonies whenever the culture covered about 50% of the plate surface. All cultures included in this study were eventually preserved in a solution of 10% glycerol and skim milk and stored in liquid nitrogen. To ensure the cultures will be available to future researchers, I sent them, with Longcore’s encouragement to the American Type Culture Collection (ATCC) for permanent storage. Some isolates have not yet been accepted by the ATCC because they were deemed “too similar” to others. Identification numbers for the Monoblepharidomycetes and their ATCC numbers are listed in Table 2.1. Table 2.2 lists the species from the Assembling the Fungal Tree of Life (AFTOL) database included in this study along with their current classification to phylum. Culturing for Microscopic Observation I grew Monoblepharis macrandra JEL 501 and Gonapodya prolifera JEL 478 at 23°C on TYG media (2.5 g tryptone, 1.25 g yeast extract, 3.0 g glucose and 5.0 g agar in 1L distilled water) because experimentally, it facilitated fast growth. To observe individual hyphae, I grew isolates in monolayers in slide culture and on sterile dialysis membrane (Fisher Scientific) on nutrient agar. I made slide cultures by dipping flame sterilized glass slides in TYG with either agar or 12% gelatin (for phase contrast optics) and inoculating them with 2 cm2 chunks of mycelium. The slides were placed in sterile petri dishes with a dampened piece of filter paper and sealed. Dishes were placed in a closed container lined with damp paper towel. Light Microscopy I observed living cells from slide cultures after 3-4 days, gently placing a glass coverslip on the visible mycelium for Nomarksi Differential Interference Contrast (DIC) and phase contrast microscopy. I looked for familiar organelles such as nuclei and signs of active cytoplasmic streaming in hyphae. For traditional chemical fixation, Roberson poured 4% formaldehyde in 0.05M PBS, 0.1% BSA and 0.02% sodium azide at pH = 6.8 (PBS/BSA) on cut dialysis membrane with fragments of mycelium or whole, small mycelia. For freeze-substitution, I followed the protocols outlined by Roberson et al. (2011), plunging the material into cold acetone at -30°C then freeze-substituting the cells in methanol in the -80°C freezer for 72 h. Samples were then allowed to come to room temperature and were slowly re-hydrated with PBS-BSA. I then observed "-tubulin and #-tubulin distribution in mycelia using immunofluorescence labeling. I rinsed the PBS-BSA from the fixed specimens with distilled !  (,!  water, spread them out on ethanol-cleaned glass slides and allowed them to dry overnight in a covered glass petri dish so that the mycelia adhered to the slide. The following day, I rehydrated the mycelial fragments in PBS-BSA then added 0.1% Novozyme (Calbiochem Corp, La Jolla, CA), incubating the material for 10-20 min to partially digest the chitinous cell wall. Next, I rinsed off the enzyme and added 500-fold diluted monoclonal "-tubulin antibody (Ab) (Accurate Chemical and Scientific Corporation, Westbury, NY) or monoclonal #-tubulin Ab (Sigma Chemical Co., St. Louis, MO), incubating the mycelium for 2-12 h in at 4°C. I rinsed slides 3 times, incubating each rinse for 5 minutes then added the 100-fold diluted secondary Alexa 488-tagged Ab (Invitrogen, Carlsbad, CA), and incubated at room temperature for 1 h. I rinsed the slides again and then added a drop of 0.1 µg/mL DAPI (Sigma). For actin visualization, I stained chemically fixed, un-dried cells in PBS/BSA with 0.33 mM rhodamine-phalloidin (Invitrogen, Carlsbad, CA) in microfuge tubes for 30 min at room temperature after membrane permeabilization with Novozyme. All solutions added to mycelia were diluted in PBS-BSA. Prior to imaging, I added 0.1% n-propyl gallate (Sigma) in 90% glycerol to prevent photobleaching. I also preserved mycelial fragments of Coemansia sp., a zygomycetous fungus, by freezesubsitution and performed immunolocalization assays on them at the same time as the Monoblepharidomycetes. I took all of the conventional light and wide-field epifluorescence microscopic images using the Axioscop (Carl Zeiss Inc., Thornwood, NY) fitted with a Roper Cool SNAP ES Digital Camera (Roper Scientific Inc., Tucson, AZ). Subsequently, Roberson and I processed these images first in Metamorph 6.0/6.1 (Universal Imaging Corporation, Downington, PA) then added scale bars in Adobe Photoshop CS5. Transmission Electron Microscopy Roberson cut 0.5 mm2 pieces of M. macrandra and G. prolifera mycelium growing on dialysis membrane and placed them on TYG agar at 23°C to recover for 2 to 3 hours. We used the technique and apparatus described by Roberson et al. (2011) for rapid freezing, freezesubstitution, flat embedding and serial sectioning of fungal hyphae. Fragments of dialysis membrane were plunged into liquid propane at -186°C and were quickly transferred to vials of 2% osmium tetroxide and 0.05% uranyl acetate in acetone and incubated at -85°C. After 48 h, samples were placed at -20°C, then 4°C for 2 h each before being placed at room temperature. Samples were rinsed in anhydrous acetone then infiltrated gradually in vials containing increasing concentrations of Spurr’s resin. Once the material was in 100% resin, it was flat embedded on a cleaned glass slide coated with Teflon™. Polymerization was carried out at !  (-!  60°C for 36 h. I microscopically examined slides for intact, well-preserved hyphae and mounted the selected material on blocks of resin then hand trimmed them. Lowry used an ultramicrotome (Leica, Wetzlar, Germany) to cut ultrathin (60-80 nm) serial sections and picked them up on slot grids coated in Formvar™. Grids were post-stained in 2% uranyl acetate in 50% ethanol for 10 min then in lead citrate for 5 min. I studied the sections with the FEI CM12S TEM (FEI Electronics Instruments Co., Mahwah, NJ) and took images with an attached Gatan 689 CCD digital camera (Gatan Inc., Pleasanton, CA). TEM images were processed using Photoshop as well. DNA Extraction, Amplification and Sequencing I extracted genomic DNA from the Monoblepharidomycetes taxa using the DNeasy Plant Mini Kit (Qiagen, Toronto, ON) following the manufacturer’s protocols. Subsequently, I amplified an approximately 3 kb region of the 28S rDNA in our isolates with a total of six primers using PureTaq Ready-to-Go PCR beads (GE Healthcare, Baie d’Urfe, QC) and an Applied Biosystems GeneAmp PCR System 9700 thermocycler (Applied Biosystems, Foster City, CA). For amplifications, I used an initial 94°C wax bead melting step followed by 40 cycles with a 10 sec denaturation at 94°C, a 20 sec annealing step at 55°C and 30 sec elongation step at 72°C that increased by 4 sec each cycle. The final cycle ended with a 7 min. elongation step also at 72°C. I cleaned PCR products using ethanol precipitation and performed sequencing reactions with the primers used for the initial amplifications using ABI PRISM BigDye Terminator Cycle Sequencing Kit V3.1 chemistry (Life Technologies-Applied Biosystems, Streetsville, ON). I purified the sequencing products using the DyeEx Terminator Removal Kit (Qiagen, Toronto, ON) according to manufacturer’s instructions. Sequencing reaction products were analyzed with an ABI 3730 DNA Analyzer (Life Technologies-Applied Biosystems, Streetsville, ON). I obtained clean amplicons and sequence for 37 Monoblepharidomycetes using the primer pairs LROR/LR7 and LR3R/LR12 (Vilgalys and Hester, 1990). Sequencing with primers LR3R and LR12 left a gap in coverage roughly 500 bp long. I sequenced across this gap in 11 isolates, including one isolate of each species, using primer combinations: LR3R/LR9 and LR8R/LR12 (http://www.biology.duke.edu/fungi/mycolab/primers.htm) (Table 2.1). About 94% of the 28S fragment was sequenced in both directions for those 11 isolates.  !  (.!  Sequence Assembly and Alignment I used Sequencher ver. 4.10.1 (Gene Codes, Ann Arbor, MI) to assemble the sequence fragments. I then obtained 56 fungal sequences and sequences from Monosiga brevicollis and Homo sapiens from the AFTOL database (http://aftol1.biology.duke.edu/pub/alignments). The non-fungal sequences served as outgroups for rooting the fungal tree. I created a starting alignment for manual editing with MacClade 4.05 (Maddison and Maddison, 2000) using the MAFFT (Katoh et al., 2005) Q-INS-i algorithm found on the MAFFT server (http://mafft.cbrc.jp/alignment/server/index.html). The Q-INS-i algorithm accounts for the secondary structure of RNA and is useful for aligning highly divergent sequences (Katoh and Toh, submitted). Due to the complexity of the data set, I could not accurately assess positional homology in some regions. Since I was most interested in phylogenetic relationships within the Monoblepharidomycetes, I aligned the closely related Monoblepharidomycetes sequences together against large gaps and made character sets from the displaced regions of hard-to-align sequences. I also removed large regions of H. sapiens sequence from the alignment that caused small gaps to break up fungal sequence regions that were highly similar. I made exsets from sequences that did not overlap with any other species in the alignment. I made all modifications using MacClade then excluded the exsets and character sets using PAUP*4.0b10 (Swofford, 2003). The final alignment contained 3459 characters. Phylogenetic inference Based on results from MODELTEST 3.7 (Posada and Crandall, 1998; Tavaré, 1986), I used the General Time Reversible (GTR) model (Stamatakis, 2006; Tavaré, 1986) estimating site-to-site variation using a gamma distribution. I used RAxML 7.2.8 (Stamatakis, 2006) to infer maximum likelihood phylogenies and complete bootstrap replicate searches. RAxML performed 100 independent likelihood searches followed by 100 bootstrap replicates under the GTRGAMMA model. I interpreted bootstrap support values $70% as moderate support for a clade and values $90% as strong support. I considered branches with less than 50% support to be unreliable. I also inferred a phylogeny using MrBayes 3.1.2, a GTR model, a gamma distribution with an estimated shape parameter, and an estimated proportion of invariant sites (Huelsenbeck and Ronquist, 2001; Ronquist and Huelsenbeck, 2003). I ran two Markov fourchain analyses for 20, 000, 000 generations, sampling every 10000 generations and discarding the first 1092 of the 2001 sampled trees as burnin. Using Tracer (Rambaut and Drummond, 2007), I verified that the effective sample size in each of the two separate runs after burnin had !  %/!  exceeded 200, suggesting that equilibrium had been reached. I considered a clade to be considered strongly supported using if its posterior probability was $98% in Bayesian analyses. Trees were rooted and modified for uniformity in FigTree v.1.3.1 (available from http://tree.bio.ed.ac.uk/software/figtree). Ancestral State Reconstruction To implement ancestral state reconstructions, I imported my likelihood tree from RAxML into Mesquite (Maddison and Maddison, 2010) and created a matrix with the following characters: pseudoseptation and hyphal growth. I then traced the origins of pseudosepta using a parsimony reconstruction of ancestral states. I chose a parsimony-based inference method because only two species, from the same genus, had pseudosepta, which led me to assume that transitions to (or from) these states were rare. I used likelihood for ancestral state reconstruction of hyphae. Some species of fungi such as Candida albicans are dimorphic, producing both yeast cells and hyphae and even pseudohyphae (Harris, 2011). For character state coding, I considered these to be hyphal since the analysis required single, defined states. A likelihood ratio test indicated that a symmetrical model, with the same rate of forward as backward change, was more appropriate for my data than the more complex asymmetrical two-parameter model, with different rates for forward and backward character change. Consequently, I applied the Markov k-state 1 parameter likelihood model (Mk1) of Lewis (2001) in Mesquite (Maddison and Maddison, 2010) to trace the evolution of hyphae in the fungi. I inferred the state at internal nodes according to a likelihood decision threshold criterion. If likelihoods of the two states at a node differed by 2.0 or more the negative log units, I concluded that the more likely state was unambiguously present at that node. If the difference was less than 2.0 units, the character state was ambiguous. To assess how robust my reconstruction was to my choice of a model, I also tested more complex asymmetrical models, constraining and incrementally increasing the rate of hyphal loss.  ! ! !  !  %(!  2.3 Results Light Microscopic Observations of Vegetative Hyphae M. macrandra did not grow well in slide culture therefore, I include only one image of the hyphal tip at the light microscope level here (Fig.2.2a). Conversely, G. prolifera grew vigorously (Fig. 2.2c-d, Fig. 2.4). Hyphae of both species were ~3-5 µm in width (Figs. 2.1, 2.2, 2.4). Typical G. prolifera vegetative hyphae possessed irregularly spaced pseudosepta (Fig. 2.4a, d) and nuclei with distinct nucleoli (Fig. 2.4b). Hyphae branched at different angles (Fig. 2.4c) and sometimes had a catenulate appearance (Fig. 2.4e). Transmission Electron Microscopic Observations of Vegetative Hyphae Both species contained eukaryote-typical organelles such as mitochondria (Figs. 2.5a-b, 2.6, 2.7b, 2.9a), nuclei (Figs. 2.6 and 2.7), extensive rough endoplasmic reticulum (RER) covered in ribosomes (Figs. 2.5, 2.6, 2.7, 2.9, 2.10), stacked Golgi cisternae (Fig. 2.5a,c, 2.6b,d, 2.7a-d, 2.9a,c), lipid bodies and multivesicular bodies (Figs. 2.5a, 2.6a-b, 2.7c-e, 2.9a,c). Golgi bodies were either flat (Figs. 2.5c and 2.9a) or cup-shaped (Figs. 2.5a and Fig. 2.9c). Nuclei were usually roughly ovoid but could be angular or elongate (Figs. 2.6 and 2.7). Serial sections revealed an unusual grid-like or mesh-like (mesh size = 28 nm) structure that resembled a rectangular wire fence rolled around a vacuole near the nucleus in M. macrandra (Fig. 2.8). Golgi bodies were often closely associated with the microtubule-centriole complex (Figs. 2.6d and 2.7d). Elongate (Figs. 2.5a-b and 2.9a) and sometimes branching mitochondria (Fig. 2.7b) were oriented parallel to the length of the hypha. Hyphae also contained large, dense granular bodies that appeared to lack a delimiting membrane (Figs. 2.9a and 2.10). These bodies were prominent in G. prolifera, sometimes taking up the width of a hypha (Figs. 2.9 and 2.10) but they were rarer in M. macrandra (Fig. 2.10d). They were often found near RER (Fig. 2.10a,c). In more mature segments of M. macrandra hyphae, elongate to irregularly shaped membrane bound vacuoles were common (Figs. 2.5d and 2.6c). Towards the outside of these vacuoles, the contents were relatively electron-transparent, however the middle of these vacuoles fell out of section leaving behind an empty, white space (Figs. 2.5d and 2.6c). Furthermore, the G. prolifera cytoplasm appeared to be partitioned so that areas occupied by a low density of large particles were clearly separated from areas containing a high density of small particles (Fig. 2.10a). TEM also revealed the layering of cell wall material (Fig. 2.11a) in the irregularly spaced pseudosepta of G. prolifera, which appeared as opposing hump!  %%!  shaped cell wall thickenings (Fig. 2.11). Pseudosepta often occurred at branching points (Fig. 2.11c). The cytoplasm of G. prolifera was continuous through the pseudosepta (Fig. 2.11), allowing nuclei to squeeze through as demonstrated by the “barbell-shaped” nuclei viewed using wide-field epifluorescence microscopy (Fig. 2.12). The cell wall of M. macrandra appeared to consist of three layers, an outer (OL), middle (ML) and an inner layer (IL) (Fig. 2.13a). All layers were uneven in thickness. The middle layer was more opaque than the other two layers, which have roughly equal electron opacities. The outer surface of the outer layer was also slightly bumpy (Fig. 2.13a). The cell wall of G. prolifera was also layered. The outermost layer (OL) was roughened or flaky (Fig. 2.8b). The inner layer had an even texture while the middle layer was very thin. Hyphal apices I concluded that there was no spherical body in the hyphal apices of either M. macrandra or G. prolifera that was suggestive of a Spitzenkörper under the light microscope (Fig. 2.2a, c-d). Nor was there a crescent shaped body as in Rhizopus (zygomycetous fungus, Fig. 2.2g). Live culture filming confirmed that growth was truly occurring at the tips of hyphae (results not shown). I will follow the conventions defined by López-Franco and Bracker (1996) for discussing the different parts of a hypha. At the TEM level, vesicles were concentrated in hyphal apices of M. macrandra (Fig. 2.13a-c) and G. prolifera (Fig. 2.14a-c). The apical dome of M. macrandra, the part of the apex contained in the curved part of the hyphal tip, contained only vesicles, while the subapical region, just below the dome, contained mitochondria (Fig. 2.13b-c). In G. prolifera, only vesicles and filamentous actin were found (Fig. 2.14a-c). Neither species has a spherical, ring-shaped or crescent-shaped organization of vesicles in its hyphal apices. However, I observed different kinds of vesicles in hyphal tips (Figs. 2.13 and 2.14). Some vesicles between 30-50 nm had contents with high electron density while others had contents with low density. Microvesicles were ~ 30 nm in diameter. Unlike G. prolifera, M. macrandra tip sections also contained larger vesicles with electron-lucent contents, diameters between 50-80 nm, and an electron-opaque coating (Fig. 2.13a-c). Clumps of actin-like filaments were in close proximity to the plasma membrane in G. prolifera (Fig. 2.14c). The long fine filaments in G. prolifera labeled ‘A’ are likely actin microfilaments (Fig. 2.14a-b). Additionally, the inner face of the plasma membrane at hyphal apices was darkened and bumpy (Figs. 2.14 and 2.15).  !  %)!  The Microtubule Cytoskeleton Wide-field epifluorescence imaging revealed astral-like arrays of microtubules surrounding DAPI-stained nuclei (Fig. 2.15). Thin hyphae contained nuclei in single file whereas thicker hyphae sometimes housed clustered nuclei (Fig. 2.15d). The zygomycete Coemansia sp., which was fixed alongside the Monoblepharidomycetes had an extensive array of cortical microtubules as well as spindle microtubules (Fig. 2.16). In M. macrandra, microtubule nucleating microtubule organizing centres (MTOCs) were solitary or in pairs on opposite poles of the nuclei (Fig. 2.17b,d). The close association between MTOCs (centrioles), nuclei and microtubules was confirmed in TEM images (Figs. 2.5, 2.6, 2.18). TEM serial sections revealed nuclei with prominent nucleoli surrounded by microtubules emanating from a single centriole in M. macrandra (Fig. 2.18, G. prolifera not shown). The Actin Cytoskeleton Epifluorescence images demonstrated the presence of actin plaques, cables and monomers. Actin plaques (or patches) were highly concentrated at hyphal apices but seemed to be absent from the very tips of hyphae (Figs. 2.19 and 2.20). These plaques may correspond to what appear to be clusters of actin-like filaments docked on the plasma membrane in TEM images (Fig. 2.14c). Cortical plaques also dotted the hyphal length (Fig. 2.20). Long cables of actin formed a network in the hyphal cortex that ran down the hyphae (Fig. 2.21). Faint, diffuse actin staining was found throughout the hyphae (Fig. 2.21). Actin did not appear to form rings at pseudosepta in G. prolifera (Fig. 2.22). Actin did not stain at all in air-dried material. Phylogeny of the Fungi In the phylogeny of the fungi, the same branches received strong support from both likelihood and Bayesian inference. I recovered strong support for the Fungi as a monophyletic lineage. In addition, each of the hyphal fungal phyla recognized by Hibbett et al. (2007), namely the Ascomycota, Basidiomycota, Glomeromycota and Blastocladiomycota, were monophyletic (Fig. 2.23). The monophyly of the ‘core chytrid clade’ excluding the Monoblepharidomycetes received moderate support. The zygomycete insect parasites, Conidiobolus coronatus and Entomophthora muscae formed a sister clade to Blastocladiomycota, which contains the hyphal chytrid, Allomyces. The blastocladiomycotan fungi, particularly Coelomomyces stegomyiae, and some of the zygomycete such as Coemansia reversa were extremely divergent, producing long  !  %*!  branches in the phylogeny. The placement of Rozella at the basal split in the Fungi received moderate to strong support (Fig. 2.23). The Monoblepharidomycetes appeared monophyletic with Hyaloraphidium as the sister group of the rest of the Monoblepharidomycetes (Fig. 2.23, 2.24). I recovered strong support for the monophyly of mycelial Monoblepharidomycetes consisting of the three genera, M. macrandra, Monoblepharella and Gonapodya (Fig. 2.24). The moderately supported Gonapodya clade formed a sister group to the Monoblepharis-Monoblepharella clade. Furthermore, with strong support, the rod-shaped Oedogoniomyces were nested among species of Harpochytrium. This Oedogoniomyces/Harpochytrium clade was the sister group to the hyphal Monoblepharidomycetes. Isolate L9 fell into the Monoblepharella clade while the three other unidentified monoblepharidalean isolates (w20, w187, and unk7) belonged to the Monoblepharis clade. Interspecific relationships were not well resolved and Gonapodya polymorpha isolate JEL452 was strikingly divergent from other isolates identified as being conspecific. Although isolate JEL 490 was identified as a species of Monoblepharis, this phylogeny placed it in Monoblepharella (Fig. 2.24). Conversely, JEL 494 was labeled as Monoblepharella but it appeared in the Monoblepharis clade (Fig. 2.24). Ancestral State Reconstruction The evolutionary history of hyphae reconstructed in this study is presented in Fig. 2.25. Character state reconstruction at all of the internal nodes unambiguously implied that the common ancestor of the Fungi was unicellular, not mycelial. Hyphae originated independently in the ancestor of the mycelial Monoblepharidomycetes (Monoblepharis, Gonapodya, Monoblepharella), in the ancestor of the terrestrial Fungi, and in Polychytrium aggregatum. By testing different rates of transition to and from the hyphal state I found the results of the ancestral state reconstruction to be robust to alternative models of evolution. While in the most likely model, rates of gain and loss of hyphae were the same, I had to constrain the rate of hyphal loss to be at least thirteen times greater than the rate of hyphal gain before the fungal ancestor was no longer unambiguously reconstructed as non-hyphal. Even with this constraint, the likelihood of a unicellular ancestral state remained higher than the likelihood of a hyphal ancestry (Fig. 2.26). I had to constrain the rate of hyphal loss to be more than twenty-four times greater than the rate of gain before the likelihood that the fungal ancestor was hyphal dropped to 50% (result not shown). As expected, the parsimony based reconstruction !  %&!  indicated that pseudosepta arose only once within the Monoblepharidomycetes in the ancestor of Gonapodya (Fig. 2.27).  2.4 Discussion On the Origins of Hyphae My analyses provide evidence for three independent origins of the hyphal habit from a unicellular ancestor, under a likelihood reconstruction of ancestral states (Fig. 2.25). Hyphae independently originated in the common ancestor of terrestrial fungi, in the Monoblepharidomycetes and in the ancestor of P. aggregatum. I did not include the ascomycotan yeasts most closely related to the hyphal cotton pathogen, Ashbya gossypii, in my analyses. However, hyphae are also a derived character in Ashbya based on the nesting of this species among yeasts (Dujon, 2010; Schmitz and Philippsen, 2011). Thus, hyphae have evolved at least four times in the fungi. Based on a likelihood ratio test, I applied a single-rate parameter model to a likelihood ancestral state reconstruction. This single-rate model assumed that evolutionary hyphal gain and hyphal loss occurred at the same rate. I then demonstrated that the ancestral character state for fungi was inferred to be unicellular, even when using an asymmetrical model where I specified that hyphal loss was up to 24 times as likely as the gain of hyphae. Although the evolutionary history of the fungi provides several examples of hyphal loss, it does not indicate an extreme bias towards hyphal loss (Harris, 2011). A four protein-coding gene phylogeny of the fungi suggests that the chytrid-typical thallus of Olpidium (Sekimoto et al. manuscript under review) also resulted from the loss of the hyphal habit. Any future ancestral state reconstructions exploring hyphal evolution would benefit more from the use of a robust phylogeny with more extensive taxon sampling and from testing different tree topologies rather than from using a different likelihood model. Cellular Organization in Monoblepharidomycetes and its Implications Consistent with their convergent origins, the hyphae of the Monoblepharidomycetes show structural differences compared with the hyphae of all other fungi examined so far. I achieved excellent preservation in these specimens by rapid freezing and freeze-substitution as demonstrated by the sharpness of the plasma membrane and the well-defined mitochondrial !  %+!  cristae in TEM images. Also, the orientation of the mitochondria parallel to the length of the hypha indicated that the hyphae were growing normally prior to fixation. In TEM and light micrographs, monoblepharidalean hyphae lack the Spitzenkörper or a crescent-shaped array of apical vesicles found in other hyphal fungi (Grove and Bracker, 1970; Howard, 1981; López-Franco and Bracker, 1996; Roberson and Fuller, 1988; Roberson et al., 2011; Vargas et al., 1993). Even though vesicles are not arranged in a recognizable, compact cluster in Monoblepharidomycetes, the presence of diverse vesicle populations suggests some level of organization. In other fungi, apices contain macrovesicles and microvesicles with different electron-opacities (Grove and Bracker, 1970; Howard, 1981; López-Franco and Bracker, 1996; Roberson and Fuller, 1988; Roberson et al., 2011; Vargas et al., 1993). These vesicles have distinct functions. Macrovesicles are thought to contain the precursors for cell wall formation as well as enzymes destined for secretion (Bartnicki-García, 1990) and the same may be true for macrovesicles in the hyphae of M. macrandra and G. prolifera. One class of microvesicles, known as chitosomes, possess chitin synthase activities (Ruiz-Herrera and Bartnicki-García, 1974). Riquelme et al. (Riquelme et al., 2007; Riquelme et al., 2011) and Sanchez-Leon et al. (2011) found that different chitin synthase paralogs, while they localized in similar patterns, were not found in the exact same compartments. The microvesicles that I observed in monoblepharidalean hyphal apices could be chitosomes, although, I would have to isolate the vesicles and assay them for chitin synthase activity or use gold labeled anti-chitin synthase antibodies to confirm this. My studies suggest that Spitzenkörper arose after the Monoblepharidomycetes split from all other fungi. That is, filamentous chytrids found their own way to grow at their apices without a Spitzenkörper. The unclustered distribution of vesicles may be related to the irregular growth patterns of the Monoblepharidomycetes (B. Couch, UBC Botany, personal communication). Obviously, growth conditions and other metabolic characteristics of these fungi that I was unable to investigate, also have roles in differences in growth rates. However, the uneven layering and surfaces of cell walls in Monoblepharidomycetes could be the result of uneven deposition of cell wall materials due to variation in the direction and rate of transport of vesicles to the plasma membrane. Perhaps vesicle targeting and transport is less efficient in Monoblepharidomycetes, leading to slower tip growth. In culture, these fungi grew only a few micrometers per hour (data not shown), much slower than Neurospora crassa that can grow at !  %,!  a rate of 4 mm per hour (Riquelme et al., 2011). The loss of Spitzenkörper in growing hyphae has been shown to reduce rates of hyphal elongation (Reynaga-Peña et al., 1997). On the other hand, zygomycetous fungi also lack Spitzenkörper but can grow rapidly at a rate of 20 µm per minute, covering substrates overnight (RW Roberson unpublished), although they at least have some discernible form of vesicle clustering. Similarly, the long, parallel microtubular arrays found in most filamentous fungi, that appear to be absent from M. macrandra and Gonapodya may have evolved after the divergence of Monoblepharidomycetes. In other hyphal fungi, molecular motors use microtubule tracks to carry vesicles over long distances (Steinberg, 2007). Since monoblepharidalean hyphae seem to have only nucleus-associated microtubules, it is improbable that long-distance vesicle trafficking is accomplished using microtubules. I hypothesize that the elaborate actin cables in the hyphal cortex are more active in long-distance transport in Monoblepharidomycetes. Even in other fungi, actin is more critical than tubulin in polarized growth. In Ustilago maydis, polarized hyphal growth occurred even when microtubule-based motors were non-functional (Schuchardt et al., 2005). Additionally, workers have found that when actin fails to polymerize in growing hyphae, hyphal tips expand symmetrically suggesting a loss of polarized vesicle transport (Srinivasan et al., 1996; Upadhyay and Shaw, 2008). Admittedly, our methods of sample preparation and tubulin detection might explain the apparent lack of cortical microtubules. Observing the microtubular cytoskeleton in living material using GFP-tubulin constructs would be helpful but technically difficult as there is not yet a transformation system for these fungi. Suggesting that the absence of cortical microtubules was not an artifact of fixation, it was consistent through both chemical fixation and freeze substitution. The staining of microtubules associated with nuclei argued against other explanations such as lack of penetration of the antibodies or unknown proteins blocking antibody binding sites. Moreover, I fixed and stained Coemansia, a zygomycete that demonstrates a “classical” microtubule cytoskeleton, alongside the Monoblepharidomycetes also arguing against preservation or immunofluorescence artifact. Arguing against the possibility that microtubules that would normally have formed cortical arrays had all reorganized to form a spindles (Gambino et al., 1984; Szewczyk and Oakley, 2011), it was unlikely that none of the hyphae were in interphase. I examined several 2 cm2 mycelial fragments and none of them had any cortical microtubules. Even at hyphal tips, where presumably, cortical microtubules !  %-!  involved in apical growth reside during mitosis (Horio and Oakley, 2005), there were none to be found. In addition, I found only nucleus-associated microtubules in electron micrographs. Spindle formation is associated with duplication of the MTOCs. Serial sections through nuclei demonstrated that even single MTOCs had microtubular arrays suggesting that astral-like microtubules were not only associated with spindle formation. Although additional evidence would be welcome, my results argue against a cryptic population of cortical microtubules in Monoblepharidomycetes. The patches of actin in hyphal apices M. macrandra and G. prolifera could perhaps function in endocytosis in the Monoblepharidomycetes, as in other fungi such as A. nidulans (Upadhyay and Shaw, 2008). Clumps of filamentous actin-like material in TEM images could correspond to the highly concentrated actin plaques observed in light micrographs of G. prolifera and M. macrandra. In Sclerotium rolfsii (Roberson, 1992) and Allomyces macrogynus (Srinivasan et al., 1996), actin plaques corresponded to filasomes, microvesicles with electronopaque cores, coated in fibrillar actin (Bourett and Howard, 1991; Howard, 1981). Beripiki and colleagues (2010) observed actin patches in living N. crassa hyphae that they thought may have been filasomes. It is an attractive but unproven possibility that these filasomes are products of endocytosis (Fischer-Parton et al., 2000). In the Monoblepharidomycetes, clusters of actin-like filaments seemed to lack a microvesicle core, a problem in interpreting them as filasomes. Even in serial sections through the same hypha, no membrane-bound vesicle was visible within the fibrillar mass, although more observations are needed. As in animals, other Chytridiomycota, and most other eukaryotes, G. prolifera and M. macrandra have stacked Golgi cisternae and centrioles that nucleate microtubules around their nuclei (Alberts et al., 2002; Taylor and Fuller, 1981). This contrasts with other hyphal fungi, which have tubular organelles or 'Golgi equivalents' and spindle pole bodies rather than centrioles (Alexopoulos et al., 1996). Having a centriole may not have detectable consequences for polarized growth but it underscores the fact that Monoblepharidomycetes adopted the hyphal habit while retaining ancestral characteristics. Perhaps early hyphal fungi also had centrioles, stacked Golgi compartments and relied more heavily on their actin cytoskeleton than on cortical microtubules for tip growth and secretion of digestive enzymes.  !  %.!  Not all of the structures in the Monoblepharidomycetes have obvious homologs in other fungi. The function and identity of the large, dense, granular bodies in G. prolifera's cytoplasm remains unknown. Similar dense bodies have been reported in Harpochytrium hedinii (Travland and Whisler, 1971) and the authors likened them to microbodies. The term microbody has been loosely applied to an assortment of fungal structures, some of which are characterized, such as peroxisomes (de Duve and Baudhuin, 1966). More formally, microbodies are defined as electron-dense structures ranging from 0.7-1.7 µm bound by a single-unit membrane (Carson and Cooney, 1990). However, the bodies in G. prolifera do not appear to be membrane-bound, though the membrane could conceivably have been extracted during the substitution process. We were unable to find any of these dense bodies enveloped in a membrane in chemically fixed cells (results not shown). Unless we perform chemical assays on these structures, their function and composition will remain elusive. The grid-like structures in M. macrandra are also mysterious. Grid-like structures were also reported in Hyaloraphidium curvatum by Ustinova et al. (2000), who raised the possibility that the chequered-pattern was indicative of a phospholipid dense body. Superficially, the gridlike structures resemble rumposomes, which are however membranous cisternae appearing honeycomb-like in cross-section. Rumposomes were first characterized in zoospores of Monoblepharella (Fuller and Reichle, 1968) and later found in zoospores of Harpochytrium (Travland and Whisler, 1971). Because the mesh in M. macrandra did not persist through many sections, unlike a rumposome, it does not represent a tall tubular or stacked, fenestrated cisterna. Furthermore, the lines delimiting the mesh are too thin to be lipid bilayer membranes. Unlike a rumposome, the structure in M. macrandra is positioned at an angle towards the nucleus in a hypha rather than in the posterior of a zoospore. The holes in the “mesh” are distinctively square rather than hexagonal as in rumposomes in zoospores of Monoblepharis polymorpha (Mollicone and Longcore, 1994) and although they are similar in size, they are not exactly the same. Therefore, despite its general similarity to a rumposome, I suspect that it is an unrelated structure with an unknown function. I propose that it be called a "reteasome” from the Latin “rete”, meaning “net” and the Greek “soma” meaning “body”. Phylogeny of the Fungi James and colleagues presented the most comprehensive phylogenetic reconstruction of the Fungi to date based on six concatenated gene regions sampled from over 200 taxa (James !  )/!  et al., 2006a). Though I did not expect to achieve this depth of resolution using only 28S sequence data, I recovered many of the clades found previously (James et al., 2006a). The most taxon-rich, well-supported phylogeny of the zygomycetes (White et al., 2006) suggests that the group is paraphyletic, a result also found in my analyses. On the other hand, a recent fourprotein coding gene phylogeny suggested that zygomycetous fungi formed a monophyletic group (Sekimoto et al., manuscript under review). Monoblepharidomycetes was a monophyletic clade that was the weakly supported sister group to the core chytrid clade. While most investigations support this relationship (Bullerwell et al., 2003; James et al., 2006a; White et al., 2006), other evidence suggests that it arose from the deepest split in the fungal tree (Sekimoto et al., manuscript under review). Either way, the Monoblepharidomycetes diverged early from all other fungi. Phylogeny of the Monoblepharidomycetes Chamber's PhD dissertation had previously provided the only phylogenetic reconstruction of the Monoblepharidomycetes that included several representative taxa. His analysis, based on maximum parsimony analysis of partial 28s rDNA sequence, supported the monophyly of the Oedogoniomyces, Harpochytrium and Monoblepharis and separated Monoblepharidomycetes into two distinct clades. Relationships between genera were not well resolved and Gonapodya appeared to be non-monophyletic (Chambers, 2003). In contrast, my analyses demonstrated moderately to well-supported intergeneric relationships within the Monoblepharidomycetes. For instance, Gonapodya, Monoblepharis and Monoblepharella, which were all hyphal, formed a monophyletic group implying a single gain of the hyphal form. Their sister group comprised the unicellular, rod-shaped genera: Oedogoniomyces and Harpochytrium. Since Oedogoniomyces was nested among Harpochytrium species in these analyses, the delimitation of these two genera may need to be re-evaluated. Hyaloraphidium, which is pelagic and crescent-shaped, was a sister group to all other Monoblepharidomycetes. From this pattern, the ancestral state reconstructions suggest that the monoblepharidalean ancestor was a unicell and that hyphae as well as pseudosepta arose only once in the class. Although I did not try to delineate species here, Gonapodya polymorpha JEL 452 was more divergent than expected if it were conspecific with the G. polymorpha isolates. Finding isolate JEL 490 in the Monoblepharella clade and JEL 494 in the Monoblepharis clade could be the result of misidentification and should be rechecked; when specimens do not display sexual structures, they are difficult to identify accurately by morphology alone. !  )(!  The Origins and Possible Roles of Pseudosepta According to my reconstruction, pseudosepta arose only once in the ancestors of Gonapodya. Assuming that depositing large amounts of cell wall material in a pseudoseptum comes at a high cost, they likely perform some important function in Gonapodya. However, inferring their role is problematic. Although Monoblepharis and Gonapodya share the same lifestyle and habitat, Monoblepharis lacks pseudosepta. Both genera have been collected from submerged twigs in freshwater (Johns and Benjamin, 1954; Mollicone and Longcore, 1994). Though pseudosepta give hyphae the appearance of multicellularity, it is unlikely that their most significant function is in dividing hyphae into functional compartments since pores do not actually prevent migration of organelles such as nuclei (Fig. 2.12). However, as nuclei can also squeeze through pores in Ascomycota (Markham, 1994), and even Basidiomycota (Giesy and Day, 1965), most fungi straddle the line between coenocytic and cellular organisms. Although the shallow form of pseudosepta could be responsible for increasing absorptive surface area, this is inconsistent with the fact that pseudoseptal pores close in older parts of the mycelium. Other more likely hypothetical roles for septa and pseudosepta include imparting mechanical strength to hyphae (Gull, 1978). Healing of wounds could also be one of the roles of pseudosepta. In Ascomycota, septa and their associated Woronin bodies prevent cytoplasmic “bleeding” and death after injury to the mycelium (Markham, 1994). The pores in Gonapodya could also be closing in order to seal off the older mycelium after cytoplasm migrates into the younger, growing hyphal tips so that hyphae can colonize a greater area without needing to take up more materials to occupy an increasing cell volume. In fungi including M. macrandra, older parts of a mycelium are highly vacuolated. G. prolifera is not as obviously vacuolated so it may be that one of the roles of pseudosepta is to the cytoplasm to concentrate in younger hyphae in the absence of large vacuoles. This is all speculative but perhaps experiments including severing and microscopically examining Gonapodya hyphae could help define the function of pseudosepta. Conclusion The word “hypha” is almost inseparable from the word “fungus” yet this study is one of the first to begin to address how and where they evolved in the fungal tree. To elucidate the evolutionary origins of the versatile hypha, I performed a detailed investigation of the cellular !  )%!  organization of two hyphal chytrids that descend from one of the basal-most nodes in the fungi and placed this information in the context of fungal phylogeny. Plunge freezing and freezesubstitution techniques enabled me to beautifully preserve the internal organization of hyphae. In this way, I uncovered a novel hyphal form that lacks a Spitzenkörper and has only nucleus associated microtubules instead of the prominent, parallel arrays of microtubules that are the tracks for long-distance transport in every other hyphal fungus studied so far. In the absence of these parallel microtubular arrangements, it is possible that actin cables play a more central role in vesicle and organelle trafficking. Ancestral state reconstruction suggests that this unconventional hyphal form reflects the fact that it was devised within Monoblepharidomycetes rather than in the most recent common ancestor of the fungi. Three other lineages also converged on the hyphal body plan. Studying monoblepharidalean hyphae invites us to ponder the ancestors of the fungi in and around us. It is possible that these hyphae bear the vestiges of the long-extinct pioneers of the hyphal thallus.  !  ))!  2.5 Tables Table 2.1 List of Monoblepharidomycetes specimens sampled in this study and their isolate identification numbers from the University of Maine Collection and Berkeley Microgarden (Oedogoniomyces only). Species Harpochytrium sp. Harpochytrium sp. Gonapodya sp. Harpochytrium sp. Gonapodya polymorpha Gonapodya polymorpha Gonapodya polymorpha Gonapodya polymorpha Gonapodya polymorpha Gonapodya prolifera Monoblepharella sp. Monoblepharis polymorpha Monoblepharis polymorpha Monoblepharis sp. Monoblepharella sp. Monoblepharis polymorpha Monoblepharis macrandra Monoblepharis macrandra Monoblepharis sp. Monoblepharis macrandra Monoblepharis macrandra Unknown Unknown Unknown Unknown Unknown Monoblepharis hypogyna Monoblepharis hypogyna Monoblepharis hypogyna Monoblepharis hypogyna Monoblepharis macrandra Monoblepharis micrandra Monoblepharis sp. Monoblepharis polymorpha Monoblepharis insignis Monoblepharis insignis Oedogoniomyces sp.  Isolate Number JEL 94* JEL 105 JEL 183* JEL 196* JEL 451* JEL 452* JEL 453 JEL 455 JEL 456 JEL 478* JEL 485* JEL 486*  ATCC Accession Number MYA-2892 MYA-2893  MYA-4798 MYA-4799 MYA-4800 MYA-2894 200005  Longcore Longcore Longcore Longcore Mollicone Mollicone Mollicone Mollicone Mollicone Mollicone Mollicone Mollicone  JEL 488  MYA-4801  Mollicone  JEL 490 JEL 494* JEL 496  MYA-4802 MYA-4803  Mollicone Mollicone Mollicone  JEL 500 JEL 501* JEL 502 JEL 503 JEL 504 JEL 612 L9 W20 W187 Unk7 Hypho3 Hypho7 Hypho17* Hypho19 Mac104b Mic5 Mor5 Poly30 Sig103 Sig104 CR90  MYA-160  MYA-4804 MYA-4805 MYA-4806 MYA-4807  MYA-4808 MYA-4809 MYA-4810 MYA-4811 MYA-4812 MYA-4813  MYA-4814  Collector  Mollicone Mollicone Mollicone Mollicone Mollicone Longcore Whisler Whisler Whisler Whisler Whisler Whisler Whisler Whisler Whisler Whisler Whisler Whisler Whisler Whisler Emerson and Whisler  * I obtained sequence used to fill in sequence gap between LR3R and LR12 in this isolate. !  )*!  Table 2.2 List of species from AFTOL database used in this study, their identification numbers (AFTOL ID). Their present classification is noted at the phylum-level where possible according to the classification scheme of Hibbett et al. (2007). Blank boxes indicate uncertainty in classification. AFTOL ID N/A N/A 18 21 24 25 26 27 28 29 43 52 56 60 65 71 134 137 139 140 141 144 184 185 274 279 297 300 301 449 492 505 539 633 635 638 689 724 844 865 891 !  Species Monosiga brevicollis Homo sapiens Coelomomyces stegomyiae Batrachochytrium dendrobatidis Polychytrium aggregatum Monoblepharella sp. Hyaloraphidium curvatum Cladochytrium replicatum Entomophthora muscae Smittium culisetae Rhizophlyctis rosea Hypocrea citrina Geoglossum nigritum Morchella esculenta Aleuria aurantia Peziza proteana Peltigera degenii Conidiobolus coronatus Glomus mosseae Coemansia reversa Mortierella verticillata Umbelopsis ramanniana Phycomyces blakesleeanus Spiromyces aspiralis Dothidea sambuci Monilinia fructicola Rozella allomycis Allomyces arbusculus Basidiobolus ranarum Armillaria mellea Stereum hirsutum Ustilago maydis Endogone pisiformis Olpidium brassicae Synchytrium macrosporum Neocallimastix sp Rhizophydium macroporosum Ramaria rubella Paraglomus occultum Tilletiaria anomala Peltula umbilicata  Phylum (or Order if Phylum Not Applicable) ----Blastocladiomycota Chytridiomycota Chytridiomycota Chytridiomycota Chytridiomycota Chytridiomycota Entomophthorales Chytridiomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Entomophthorales Glomeromycota Mucorales Mucorales Mucorales Ascomycota Ascomycota Rozellida/Cryptomycota* Blastocladiomycota Basidiomycota Basidiomycota Basidiomycota Mucorales Chytridiomycota Neocallimastigomycota Chytridiomycota Basidiomycota Glomeromycota Basidiomycota Ascomycota )&!  AFTOL ID 906 1069 1072 1074 1076 1078 1079 1081 1082 1084 1087 1088 1192 1199 1459  Species Orbilia auricolor Saccharomyces cerevisiae Ashbya gossypii Candida albicans Yarrowia lipolytica Neurospora crassa Aspergillus fumigatus Magnaporthe grisea Fusarium graminearum Coccidioides immitis Coprinopsis cinerea Cryptococcus neoformans Pneumocystis carinii Schizosaccharomyces pombe Puccinia graminis  Phylum (or Order if Phylum Not Applicable) Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Ascomycota Basidiomycota Basidiomycota Ascomycota Ascomycota Basidiomycota  --- Not part of Hibbett et al. classification scheme *Based on findings of Lara et al. (2010) and Jones et al. (2011) ! !  !  )+!  2.6 Figures  a  b  c Figure 2.1 Three species in the Class Monoblepharidomycetes possess a range of body forms a) Hyaloraphidium curvatum. Scale = 10 µm. (Photo: Marilyn Mollicone) b) Harpochytrium sp. with a holdfast towards the top of the image. Scale = 5 µm. c) Gonapodya prolifera. Scale = 5 µm.  !  ),!  a *  b  4 µm  c  d  *  e  3.5 µm  f  1 µm  g  Figure 2.2 Hyphal tips of phylogenetically diverse fungi with and without Spitzenkörper viewed under phase contrast or Nomarski differential interference optics (DIC). a) Monoblepharis macrandra lacks Spitzenkörper under phase contrast optics. b) Sclerotium rolfsii under DIC optics. c) Gonapodya prolifera also lacks a Spitzenkörper under DIC optics. d) Gonapodya prolifera hyphae under phase contrast optics. e) Phase contrast image of Neurospora crassa hyphal tip. f) Allomyces macrogynus viewed under DIC optics. g) Hyphal tip of Rhizopus oryzae has a crescent-shaped dark body when viewed using phase contrast optics. Asterices indicate presence of Spitzenkörper. Images b, f and g courtesy of Robert W. Roberson. Scale = 5 µm unless otherwise noted in the figure.  !  )-!  6  !"#$%$&'()&*"+ !  !"#$%&'()* ++++%&#,-((+  6  ,#)&'()&*"+ !  .$)%&-/&%0+ 1&'%2'(-+ .$"'%(0++  -.&'/0&'()&*"+  3-/$%4(##)-+  3-5620+  7(40-/&%0+ 85(9&/)-+ :&%'($%$##0+ ;&$*0<-(0+  !  1(2&'()&*" !  6  !"#$%$&3&."./#+ !  10-(=(&6&#)-+  6  !."#*&)."%$&'()&*"+ !  3##&*2"$-+  !"  45(*0$%$&'()&*"+ !  7&<0/&=20+  !" :&<&>+ 6#$/50%(-+  Figure 2.3 Phylogenetic distribution of Spitzenkörper in the Kingdom Fungi. From Robert W. Roberson, unpublished. Taxa with asterices possess Spitzenkörper. The ultrastructural organization of hyphal apices in the Chytridiomycota was previously unknown. The arrowhead indicates that Spitzenkörper were secondarily derived in Ashbya (Dujon 2010, Schmitz and Philippsen 2011).  !  ).!  Figure 2.4 Typical appearance of vegetative hyphae of Gonapodya prolifera under the light microscope. a) Pseudosepta appeared as raised, paired humps under Nomarski differential interference contrast optics. Small inclusions (I) were visible in the cytoplasm. b) A nucleus with a prominent nucleolus near a hyphal tip. c) Hyphae with branch points indicated by asterices. Hyphae branched at different angles. d) Pseudosepta (black arrowheads) viewed under phase contrast optics gave hyphae a multicellular appearance. e) Hyphae occasionally had a catenulate appearance. I = inclusion, N = nucleus. Scale = 5 µm.  !  */!  N I  b  a  c  d  e !  *(!  Figure 2.5 Ultrastructural features of a typical, vegetative Monoblepharis macrandra hypha. a) The apical dome is devoid of organelles besides vesicles but subapical regions contain numerous mitochondria oriented parallel to the hyphal length. Hyphae also contained lipid bodies and cup-shaped Golgi bodies. b) Extensive rough endoplasmic reticulum and elongate mitochondria were common in hyphae. c) In addition to cup-shaped Golgi bodies, there were flat, stacked Golgi cisternae. d) More distal regions of the hypha were highly vacuolated. The centre contents of irregularly shaped, membrane-bound vacuoles fell out of section leaving behind white spaces. Golgi Bodies = GB, Lipid bodies = Li, Mitochondria = M, Rough Endoplasmic Reticulum = RER, Vacuole = V. Scale = 0.5 µm.  !  *%!  GB  M M  RER  b M Li  c  GB  GB MVB  a  !  V  RER  d  *)!  N M M  V  Nu  Li Li  a  Li V  GB  b  MVB M  M N  C  V RER N  c  GB  d  Mts  Figure 2.6 Nuclei and surrounding organelles in subapical regions of hyphae in Monoblepharis lipid bodies. b) Stacked Golgi bodies were found in close proximity to nuclei. Lipid bodies and multivesicular bodies were also common in subapical regions of the hypha. c) Nuclei in more distal regions could be surrounded by vacuoles and rough endoplasmic reticulum. d) A high magnification view of a centriole nucleating a microtubule aster with a nearby Golgi body. Golgi cisternae were commonly found near nucleus-associated microtubules. Golgi Body = GB, Lipid Bodies = Li, Mitochondria = M, Multivesicular body = MVB, Nucleolus = Nu, Rough Endoplasmic Reticulum = RER, Vacuole = V. Scale = 0.5 µm.  !  **!  Figure 2.7 Nuclei and surrounding organelles in subapical regions of hyphae in Gonapodya prolifera. a) An oblong nucleus with a prominent nucleolus. b) Nucleus surrounded by Golgi bodies, extensive rough endoplasmic reticulum, mitochondria, lipid bodies and microtubules. c) Nucleus surrounded by Golgi and lipid bodies. d) Golgi body and microtubules from image ‘c’ at higher magnification. e) Mitochondria with plate-like cristae were often found near nuclei. f) Single astral-like microtubular array. Golgi Body = GB, Lipid bodies = Li, Microtubules = Mts, Mitochondria = M, Nucleolus = Nu, Rough endoplasmic reticulum = RER. Scale = 0.5 µm.  !  *&!  GB  Nu  RER  a  b  N  M Mt  N Li GB  Nu  GB  c  d  Mts  Mts  Li  N N  e !  Li  M  Mts  f *+!  Figure 2.8 Mesh-like ‘reteasomes’ surrounded vacuoles near nuclei in Monoblepharis macrandra. Images a-e are serial sections through the reteasome (black arrowhead). f) High magnification of the reteasome showing the square shape of the mesh. Nucleus = N. Scale = 0.5 µm.  !  *,!  N N  a  b  N  N  c  d  N  e !  f *-!  Figure 2.9 Gonapodya prolifera hyphae contained eukaryote-typical organelles as well as uncharacterized dense granular bodies. a) Dense granular bodies were prominent in the cytoplasm. Mitochondria, Golgi Bodies, rough endoplasmic reticulum and lipid bodies were also found in the cytoplasm. The cell wall exterior is roughened and has an uneven texture. b) High magnification view of the cell wall showing the inner, middle and outer layers. Small black arrows indicate the flaky exterior of the cell wall. c) Cup-shaped Golgi bodies and multivesicular bodies populated the hyphae. Cell wall = CW, Dense granular bodies = D, Golgi Body = GB, Inner layer = IL, Lipid bodies = Li, Mitochondria = M, Middle Layer = ML, Multivesicular body = MVB, Outer layer = OL, Rough endoplasmic reticulum = RER. Scale = 0.5 µm.  !  *.!  M D  GB  RER  Li  D M CW  a  OL  b !  GB  IL  ML  GB  MVBs  c &/!  D D  b  RER  a  RER  RER D D  c  d  Figure 2.10 Dense granular bodies were the most prominent structure in the cytoplasm of Gonapodya prolifera. a) Dense granular bodies often took up most of the width of the hypha. Also, note the partitioning of the cytoplasm. The boundaries demarcating this cytoplasmic partitioning are indicated by the white arrowheads. b) Granular bodies lacked a bounding membrane. c) Rough endoplasmic reticulum was common near dense granular bodies. d) Dense granular bodies were also found in Monoblepharis macrandra but they were not as prominent. Dense granular bodies = D, Rough endoplasmic reticulum = RER. Scale = 0.5 µm. !  &(!  Figure 2.11 Ultrastructure of Gonapodya prolifera pseudosepta. a) Cytoplasm and organelles are continuous through the pseudosepta. Pseudosepta are formed by gradual deposition of the cell wall materials. Black arrowheads show the individual layers of cell wall material in the pseudoseptum. Pseudosepta were irregularly spaced. b) A single pseudoseptum. c) Pseudoseptum at a branching point. Scale = 0.5 µm.  !  &%!  a  b  !  c  &)!  Figure 2.12 Barbell-shaped nuclei demonstrated that nuclei often squeezed through pores in pseudosepta. a) Phase contrast image of a hypha with pseudoseptum indicated by white arrowheads. b) DAPI-stained barbell-shaped nucleus (blue) squeezing through pseudoseptal pore. Scale = 10 µm.  !  &*!  Figure 2.13 Ultrastructure at the hyphal apex of Monoblepharis macrandra. a) Near median section of hyphal apex illustrating a concentration of apical vesicles and uneven trilayered cell wall. The apical dome is devoid of organelles but mitochondria can be found just below this region. White arrows indicate coated large apical vesicles with electron-lucent contents. Cell wall layers are evident. b) Different vesicle populations of the hyphal apex. Apical vesicle contents were electron-opaque (large black arrowheads), electron-lucent (white arrows) or had an intermediate electron opacity (large white arrowheads). Microvesicles had electronopaque cores (small black arrowheads). c) Apical vesicles and microvesicles under higher magnification. Inner layer = IL, Middle layer = ML, Mitochondria = M, Outer layer = OL. Scale = 0.5 µm.  !  &&!  ML OL IL  M  a  b !  c &+!  Figure 2.14 Ultrastructure at the hyphal apex of Gonapodya prolifera. a) Median section of hyphal tip and subapical regions demonstrating the presence of vesicles but not the tight clustering suggestive of a Spitzenkörper. What appear to be peripheral actin cables are also found in the tip. b) Higher magnification image of the same hyphal tip demonstrating the different vesicle types. Electron-opaque apical vesicles (large black arrowhead) and microvesicles (small black arrowhead), as well as electron-lucent vesicles (large white arrowhead) populate the hyphal apex. c) Same hyphal tip at higher magnfication again illustrating different vesicle populations as well as actin-like plaques (left of asterisk). Actin = A. Scale = 0.5 µm.  !  &,!  A  a  b  !  A  c  &-!  Figure 2.15 Immunolocalization of "-tubulin in Monoblepharis macrandra hyphae revealed astral-like arrays of microtubules flanking nuclei. a) Phase contrast image of hypha. b) Paired astral-like arrays of microtubules surrounding nuclei. c) DAPI stained nuclei. d) Overlay of images a, b and c. Scale = 2 µm.  !  &.!  a  b  c Figure 2.16 Immunolocalization of "-tubulin in hyphae of Coemansia sp. demonstrated both canonical cortical arrays of microtubules as well as spindle microtubules. a) DAPI stained nuclei. b) Spindle microtubules surrounding nuclei were found in some hyphae (white arrow), whereas others contained the usual cortical microtubules (white arrowhead). c) Overlay of images a and b. Scale = 10 µm.  !  +/!  Figure 2.17 Immunolocalization of #-tubulin in Monoblepharis macrandra hyphae. a) Phase contrast image of hyphae. b) Distribution of #-tubulin representing microtubule organizing centres in hyphae. c) DAPI stained nuclei. d) Overlay of images a, b, and c. Scale = 2 µm.  !  +(!  !  +%!  M C MTs  Li N  Figure 2.18 An ultrathin section through a subapical region in a hypha of Monoblepharis macrandra showing a centriole nucleating an astral-like array of microtubules adjacent to a nucleus. Mitochondria and lipid bodies are also visible. Serial sections through nuclei only revealed single asters. Centriole = C, Lipid bodies = Li, Microtubules = MTs, Nucleus = N. Scale = 0.5 µm.  !  +)!  +*!  Figure 2.19 Actin plaques concentrated at hyphal apices in Gonapodya prolifera. a) Phase contrast view of mycelium. b) Hyphae stained with rhodamine-phalloidin to detect actin. c) Overlay of images a and b. Scale = 20 µm.  !  !  +&!  a  b  c  Figure 2.20 Actin plaques concentrated at hyphal apices in Gonapodya prolifera but also dotted hyphal length at 400x magnification. The very tips of hyphae did not stain strongly indicating that actin plaques were mostly located in the regions directly flanking the very apex. a) Phase contrast view of mycelium. b) Hyphae stained with rhodamine-phalloidin to detect actin. c) Overlay of images a and b. Scale = 10µm.  !  !  Figure 2.21 Series of images through a hypha of Monoblepharis macrandra taken by optically focusing through the hypha. a) Nomarski differential interference contrast microscopic view of hypha. Images b-d demonstrate the peripherally located actin cables (white arrowheads) and plaques as well as diffuse cytoplasmic staining suggestive of actin in monomeric form. Scale = 5 µm.  !  ++!  a  b  c  d !  +,!  Figure 2.22 Actin plaques in the periphery of Gonapodya prolifera hyphae in distal parts of the mycelium. White arrowheads indicate location of pseudosepta. There was no evidence of actin rings at pseudosepta. a) Phase contrast view of hyphae. b) Hyphae stained with rhodamine-phalloidin to detect actin. c) Overlay of images a and b. Scale = 10 µm.  !  +-!  Figure 2.23 Phylogeny of the fungi based on 28s rRNA sequence using maximum likelihood and Bayesian inference methods. Values above branches indicate bootstrap support (left) and posterior probabilities (right) for clades. Support values are only given for basal clades and only if the bootstrap support values and Bayesian posterior probabilities are over 50% *Blastocladiomycota  !  +.!  *)#!""$ ))#)*$ !""#!""$  )!#)*$  *'#!""$  )&#'($ *)#!""$  Puccinia graminis Cryptococcus neoformans Tilletiaria anomala Ustilago maydis Schizosaccharomyces pombe Pneumocystis carinii Candida albicans  Saccharomyces cerevisiae Ashbya gossypii Dothidea sambuci Fusarium graminearum Hypocrea citrina Neurospora crassa Magnaporthe grisea Monilinia fruticola Peziza proteana Aleuria aurantia Morchella esculenta Peltigera degenii Coccidiodes immitis Aspergillus fumigatus Orbilia auricolor Peltula umbilicata Geoglossum nigritum Paraglomus occultum Glomus mosseae Mortierella verticillata Endogone pisiformis Umbelopsis ramanniana Phycomyces blakesleeanus Olpidium brassicae Basidiobolus ranarum Smittium culisetae Spiromyces aspiralis !""#!""$  Gonapodya prolifera JEL 478 Gonapodya sp. JEL 183  %&#!""$  %"#!""$  !""#!""$  '(#!""$  unidentified isolate L9 Monoblepharis sp. JEL 490 Monoblepharella sp. Monoblepharella sp. JEL 485 Monoblepharis insignis sig104 Monoblepharis micrandra mic5 Monoblepharis insignis sig103 Monoblepharis polymorpha JEL 488 Monoblepharis polymorpha JEL 496 Monoblepharis polymorpha JEL 486 unidentified isolate unk7 Monoblepharis macrandra JEL 502 Monoblepharis hypogyna hyp7 Monoblepharis hypogyna hyp17 Monoblepharis hypogyna hyp19 unidentified isolate w20 Monoblepharis hypogyna hyp3 Monoblepharis sp. mor5 unidentified isolate w187 Monoblepharis polymorpha pol30 Monoblepharis macrandra mac104b Monoblepharis macrandra JEL 500 Monoblepharis macrandra JEL 504 Monoblepharis macrandra JEL 503 Monoblepharella sp. JEL 494 Monoblepharis macrandra JEL 501 Harpochytrium sp. JEL 196 Harpochytrium sp. JEL 105 Oedgoniomyces sp. CR90 Harpochytrium sp. JEL 94 Hyaloraphidium curvatum Rhizophydium macrosporum Rhizophlyctis rosea Polychytrium aggregatum Cladochytrium replicatum Neocallimastix sp. Rozella allomycis Monosiga brevicollis  Allomyces arbusculus  Basidiomycota Yarrowia lipolytica  Ascomycota  Glomeromycota zygomycetous fungi !!"  Conidiobolus coronatus  Coemansia reversa Entomophthora muscae Coelomomyces stegomyiae  *  Gonapodya polymorpha JEL452 Gonapodya polymorpha JEL 451 Gonapodya polymorpha JEL 453 Gonapodya polymorpha JEL 455 Gonapodya polymorpha JEL 456  Monoblepharidomycetes  Batrachochytrium dendrobatidis  core chytrid clade  Synchytrium macrosporum  Homo sapiens  "+,$-./0123$  Rozella outgroup !  !""#!""$  ,/!  Coprinopsis cinerea Armillaria mellea Stereum hirsutum Ramaria rubella  Modified from tree 1  Figure 2.24 Portion of phylogenetic tree from Fig. 3.23 showing the phylogeny of the Monoblepharidomycetes. Values above branches indicate bootstrap support (left) and posterior probabilities (right) for clades. Only support values and probabilities over 70% are shown. Isolates within the dotted box were mycelial.  !  ,(!  ,%!  Umbelopsis ramanniana Phycomyces blakesleeanus Olpidium brassicae Basidiobolus ranarum Smittium culisetae Spiromyces aspiralis  Co Allomyces arbusculus  Gonapodya prolifera JEL 478 Gonapodya sp. JEL 183  462011*  0112011*  Gonapodya polymorpha JEL452 Gonapodya polymorpha JEL 451 Gonapodya polymorpha JEL 453 4:2011* Gonapodya polymorpha JEL 455 7%1"$%/("3 Gonapodya polymorpha JEL 456  unidentified isolate L9 Monoblepharis sp. JEL 490 Monoblepharella sp. Monoblepharella sp. JEL 485 insignis sig104 0112011* Monoblepharis Monoblepharis micrandra mic5 Monoblepharis insignis sig103 Monoblepharis polymorpha JEL 488 302011* Monoblepharis polymorpha JEL 496 Monoblepharis polymorpha JEL 486 unidentified isolate unk7 Monoblepharis macrandra JEL 502 Monoblepharis hypogyna hyp7 472011* Monoblepharis hypogyna hyp17 Monoblepharis hypogyna hyp19 unidentified isolate w20 Monoblepharis hypogyna hyp3 Monoblepharis sp. mor5 unidentified isolate w187 Monoblepharis polymorpha pol30 Monoblepharis macrandra mac104b Monoblepharis macrandra JEL 500 Monoblepharis macrandra JEL 504 Monoblepharis macrandra JEL 503 Monoblepharella sp. JEL 494 Monoblepharis macrandra JEL 501 Harpochytrium sp. JEL 196 0112011* Harpochytrium sp. JEL 105 Oedgoniomyces sp. CR90 Harpochytrium sp. JEL 94 Hyaloraphidium curvatum 442011*  312011*  Rhizophydium macrosporum !"#$%&'()#*+%",'-)* Rhizophlyctis rosea * Polychytrium aggregatum .!)%.)/-$%&'()#*+%",'-)* Cladochytrium replicatum Neocallimastix sp. Rozella allomycis Monosiga brevicollis  4%1%56.$'"#.66"3  4%1%56.$'"#*23  !"#$%&'()#*+,2-./%0%1*%,(&.23 !("6%#"$'*/*+,3 Batrachochytrium dendrobatidis 186*.&'/9)%*  Synchytrium macrosporum  !  332011* 352011*  Figure 2.25 Evolutionary history of hyphae in the fungi inferred by a likelihood model with a single backward and forward rate of transition between the hyphal and non-hyphal forms. Filled circles next to species names indicate hyphal species. Empty circles indicate non-hyphal species. Pie charts indicate the proportional negative log likelihood of ancestral states at nodes. The most recent common ancestor of the fungi (Node 1) was most likely not hyphal. Arrowheads indicate where hyphae must have arisen in the fungal tree.  !  ,)!  hyphal not hyphal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Figure 2.26 Evolutionary history of hyphae in the fungi under a likelihood model where the rate of hyphal loss was constrained to 13 times the rate of hyphal gain. Filled circles next to species names indicate hyphal species. Empty circles indicate non-hyphal species. Pie charts indicate the proportional negative log likelihood of ancestral states at nodes. The most recent common ancestor of the fungi (Node 1) was no longer unambiguously reconstructed as nonhyphal (asterisk). Arrowheads indicate where hyphae must have arisen in the fungal tree.  !  ,&!  hyphal not hyphal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Figure 2.27 Pseudosepta evolved only once within the Monoblepharidomycetes in the ancestor of Gonapodya based on a parsimony reconstruction of ancestral states (black arrowhead). Filled circles represent species that possess pseudosepta. Empty circles represent species that do not have pseudosepta. Two species included in my analysis that lie outside of the Dikarya, which are regularly septate, Allomyces arbusculus (Blastocladiomycota) and Coemansia reversa (zygomycetous fungus) also possess regular septa as indicated by the asterisk.  !  ,,!  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Chapter 3. Concluding Chapter 3.1 Outcomes of this Project in Light of Current Research Studies on the Monoblepharidomycetes, which began in the 1940s, focused largely on their sexual phases (Johns and Benjamin, 1954; Perrott, 1958; Sparrow, 1953). Barr (1990) proposed classifying chytrids into orders on the basis of their zoospore ultrastructure. This led to TEM studies of zoospores in the Monoblepharidomycetes (Mollicone and Longcore, 1994; Mollicone and Longcore, 1999). More recently, in the search to reconstruct the eukaryotic tree of life, Monoblepharidomycetes are represented in mitochondrial genome sequencing projects (Bullerwell et al., 2003) and in large-scale fungal phylogenetic reconstructions (James et al., 2006a). These phylogenetic analyses have placed the Monoblepharidomycetes as the sister group to class Chytridiomycetes in the Phylum Chytridiomycota. While Chytridomycetes contains mostly non-hyphal, zoosporic fungi, Monoblepharidomycetes contains unicellular and hyphal forms, thus providing an exceptional study system for investigating the rise of hyphae. Through my M.Sc. research, I have participated in a large, collaborative project funded by the US National Science Foundation, titled 'Assembling the Fungal Tree of Life project 2'. Our project aims to reconstruct the phylogeny of the fungi by combining both molecular sequence data and morphology. In an effort to find and catalog phylogenetically informative structures in fungi, Roberson has investigated and compared hyphal tip ultrastructure and cytoskeletal organization among the early-diverging, paraphyletic fungi informally known as the zygomycetes and chytrids (McDaniel and Roberson, 1998; Roberson et al., 2011; Srinivasan et al., 1996; Vargas et al., 1993). In line with these goals, my observations of hyphae of Monoblepharis and Gonapodya provide the first insights into the cellular organization, hyphal tip ultrastructure and mechanism of hyphal growth in Chytridiomycota. From my results, I compared monoblepharidalean hyphae with hyphae of other fungi. To provide an evolutionary framework to interpret the cytological results, I used ancestral state reconstruction, based on a molecular phylogeny of the fungi. The phylogeny inferred using maximum likelihood methods and Bayesian inference based on ribosomal 28S sequence regions agreed with previous studies in placing the Monoblepharidomycetes within the Chytridiomycota, while also confirming the monophyly of the Ascomycota, Basidiomycota, Glomeromycota and Blastocladiomycota (James et al., 2006a; James et al., 2006b; James et al., 2000; White et al., 2006). By including many monoblepharidalean species in my phylogeny, I was the first to be able to resolve intergeneric !  ,.!  relationships and to explain the evolution of hyphae and pseudosepta within the class. Thus, in the course of this study I discovered evidence for a case of convergent origin of hypha in a unique fungal lineage. Comparison of Hyphae in the Monoblepharidomycetes Monoblepharidalean microtubules were organized in astral-like arrays emanating from centrioles flanking nuclei. Even in cells that were, based on their single asters in serial sections, in interphase, microtubules were in astral-like arrays rather than in parallel arrays. In other fungi, during nuclear division, disk-shaped SPBs nucleate and organize microtubules to form the astral arrays and the mitotic spindle, and cortical microtubules become temporarily scarce (Gambino et al., 1984; Szewczyk and Oakley, 2011). However, in interphase, hyphae of most other fungi have extensive cortical arrays of microtubules needed mainly for long-distance polarized transport of cellular materials (Horio and Oakley, 2005; McDaniel and Roberson, 1998; Mouriño-Perez et al., 2006; Roberson et al., 2011; Steinberg, 2007). If the interphase, parallel arrays of microtubules are indeed absent from Monoblepharidomycetes, the Monoblepharidomycetes must have an alternative mechanism for long-distance transport. In the two species that I examined, the vesicles concentrated at hyphal tips did not form a Spitzenkörper or even an apical crescent-shaped accumulation. The hyphal apex in the Monoblepharidomycetes did contain larger apical vesicles and smaller microvesicles with electron-lucent and electron-opaque cores. Other hyphal fungi have vesicles of similar size ranges at their apices, but arranged in a more compact cluster (Grove and Bracker, 1970; Howard, 1981; Roberson and Fuller, 1988; Roberson et al., 2011; Vargas et al., 1993). To speculate, the lack of a discrete Spitzenkörper may be linked to the absence of cortical microtubules. In other fungi, the Spitzenkörper cannot form if microtubules are disrupted (Steinberg, 2007). Monoblepharidomycetes had a multilayered, uneven, roughened cell wall unlike most other fungi, which have even, regular layers of cell wall material (Fuller and Clay, 1993(Riquelme et al., 2011; Roberson and Fuller, 1988). Interestingly, these cell walls have been shown to contain cellulose (Fuller and Clay, 1993) which is rare in fungal cell walls that are predominantly made of chitin (Alexopoulos et al., 1996). Whether the different layers or unusual composition of the cell wall plays any role in the biology of these organisms is unknown. Brett Couch (UBC !  -/!  Botany, personal communication) proposed that uneven wall construction could be a consequence of the lack of a Spitzenkörper. If the rate and direction of supply of vesicles to the plasma membrane varies over time, and if the release of wall building materials is irregular, an uneven cell wall may result. Monoblepharidalean Golgi bodies are arranged in stacks as in non-hyphal chytrids and animal cells (Alberts et al., 2002; Fuller and Clay, 1993; James et al., 2006b). No other hyphal fungus examined thus far has stacked Golgi cisternae. Instead, they possess irregular and tubular ‘Golgi equivalents’ (Grove and Bracker, 1970; Howard, 1981; Roberson and Fuller, 1988; Roberson et al., 2011; Vargas et al., 1993). Whether this difference in Golgi form has any significance is unclear, and it is difficult to conceive of an experimental approach to functional comparisons between the two forms. Having highlighted the differences between vegetative hyphae of the Monoblepharidomycetes and other fungi, I next point out similarities. Monoblepharidalean hyphae contained all the usual eukaryotic cellular equipment: nuclei, Golgi, ER, vesicles, vacuoles, mitochondria, ribosomes, microtubules and actin, all encased in cell wall, as in other fungal hyphae. Their nuclei, ER, ribosomes and mitochondria appear to be no different from those of any other fungus. Suggesting conserved function, the monoblepharidalean actin cytoskeleton is similar to that of other fungi. As in other fungi, actin in Monoblepharis and Gonapodya formed individual microfilaments, plaques, and cables in the cortical region of the hypha. Actin plaques were concentrated in the regions flanking the very apex of hyphal tips. This collection of actin patches is likely related to endocytotic membrane recycling at the hyphal tip to maintain polarity (Shaw et al., 2011). While cytoplasmic bulk flow could also be moving materials to the hyphal tip, I favour actin-based transport since it is the rule in yeast. In filamentous fungi, it is assumed that actin cables function in transport based on their activity in S. cerevisiae (Pruyne and Bretscher, 2000). Actin microfilaments, observable in the Spitzenkörper core of many fungi, are thought to ferry vesicles to the apical plasma membrane over a short distance (Schuchardt et al., 2005; Steinberg, 2007). Actin cables function in intracellular vesicle transport in yeast and actin plaque movement in Aspergillus (Berepiki et al., 2010; Pruyne and Bretscher, 2000; Taheri-Talesh et al., 2008). My observations support the idea that a robust network of actin and not the sparse, nucleus-associated microtubule  !  -(!  cytoskeleton is more likely responsible for intracellular transport in the Monoblepharidomycetes. Ancestral State Reconstruction My thesis provides the first formal ancestral state reconstruction for the origin of hyphae in fungi, and importantly, I was able to include the representatives of the earliest diverging hyphal fungi, of the Monoblepharidomycetes as well as the Ascomycota/Basidiomycota/zygomycete lineage. My analyses, along with phylogenies of yeasts (Dujon, 2010; Schmitz and Philippsen, 2011), imply that hyphae have arisen four times during the evolution of fungi. To date, the origins of hyphae have only been hinted at in the literature. In his recent review of hyphal evolution, Harris (2011) speculated that rhizoids could be the precursors of hyphae in the Chytridiomycota, a possibility that I had considered early in the development of this thesis project. Stajich et al. (2009) suggested that though there are truly mycelial chytrids, the filamentous form became prevalent in the Blastocladiomycota and the Ascomycota/Basidiomycota/zygomycete lineage. In either case, there were no conclusions about whether or not the hyphae of the chytrid, Polychytrium aggregatum or those in the Monoblepharidomycetes were homologous to, for example, hyphae in Ascomycota. My ancestral state reconstructions show that hyphae arose convergently in the Monoblepharidomycetes and in other fungi. This conclusion informs attempts to reconstruct the niche of the fungal ancestor — if the fungal ancestor was not hyphal (James et al., 2006a), it was not capable of invasive growth or of, for example, forming mycorrhizae. I have shown that hyphae are not all the same any more than butterfly, budgie and bat wings are the same. While the process of convergence may have begun with conserved genetic characters in both fungal and animal examples, close examination of the end products reveals differences indicative of independent evolution. The details of hyphal construction in Monoblepharidomycetes such as absence of a typical Spitzenkörper and lack of cortical microtubules may reflect convergent hyphal origin. Intergeneric Relationships in the Class Monoblepharidomycetes Previously, the only analysis of phylogenetic relationships within the Monoblepharidomycetes was based solely on partial 28S rDNA sequence from twenty or so isolates (Chambers, 2003). That study suggested that Gonapodya was not monophyletic, thus, !  -%!  pseudoseptations arose twice, a finding that was contradicted in my thesis. Instead, from my inferred phylogeny using more sequences under maximum likelihood and Bayesian inference methods, I concluded that pseudosepta had a single origin in the ancestor of Gonapodya. Additionally, my work could inform a higher-order classification scheme within the Monoblepharidomycetes. Hyaloraphidium was formerly classified as an alga (Ustinova et al., 2000) but as sister to other Monoblepharidomycetes, it should now be classified in a fungal family. Oedogoniomyces belongs to its own monogeneric family (Barr, 1990) but it should be transferred to the Harpochytriaceae since it is nested among species of Harpochytrium.  3.2 Strengths and Limitations of this Research The fixation quality achieved here was excellent. Rapid freezing was facilitated by the small diameter of the hyphae (3-5µm). The images that I obtained from the hyphal Monoblepharidomycetes were largely free from artifacts such as ice crystals, tears, and wavy membranes. I was able to preserve delicate structures such as actin-like clusters of filaments. Hyphae truly appeared as if they were frozen instantly. Typically, during chemical fixation, the slow penetration of aldehydes through the cytoplasm as proteins become cross-linked kills cells slowly. Dying cells will self-digest leading to the destruction of the internal organization of the cell. One of the great strengths of this work was the use of rapid plunge-freezing for the initial fixation of the mycelia which prevented ice crystal formation and circumvents the problems of created during traditional chemical fixation. We obtained serial sections for interpreting the three dimensional structures in the cell. Serial sectioning was of utmost importance. With only single sections, I could not have interpreted structures such as the mysterious grid-like structure or the shape of vacuoles in the cytoplasm. Through serial sections, I observed several complete nuclei and their associated centrioles and microtubules, which confirmed light microscopic observations of single centrioles. Serial sections were necessary to conclude that the astral-like arrays did not simply represent a rearrangement of the usual parallel arrays of microtubules seen in other fungi. My molecular phylogeny of the fungi also benefited from the large number of sequences from the Monoblepharidomycetes. Chambers’ (2003) parsimony-based phylogeny of the !  -)!  Monoblepharidomycetes from 28S rDNA sequence data from 23 isolates was only able to group the Monoblepharidomycetes into two clades with Gonapodya isolates assorting into each of the two clades. Having more taxa, a second gene and using inference methods that account for variation in substitution rates, multiple substitutions, and posterior probabilities led to a well-supported phylogeny. I firmly resolved relationships between genera. Unfortunately, I was unable to contribute any new information on the placement of the Monoblepharidomycetes among other fungi because my analysis, like previous analyses, used rDNA sequence data. I had originally planned to sequence some protein coding genes, a possible useful direction for future research. The ancestral state reconstruction that I present here has the advantage of including two hyphal chytridiomycotan fungi. By combining my newly gathered information with data from other fungi (Dujon, 2010; Schmitz and Philippsen, 2011), I was able to count the number of convergent origins of hyphae in the fungi, which had not been done previously. My phylogeny was largely congruent with other phylogenies where taxon sampling overlapped, indicating that it was mostly robust. There are caveats in these types of analyses, however. Backbone relationships in my tree were not always well supported. Since my likelihood ancestral state reconstructions relied on branch lengths and branching patterns, errors in phylogenetic inference could result in errors in ancestral state reconstruction. Additionally, ancestor state reconstructions were based on a model of the relative probability of evolutionary transitions and I cannot be sure that the single parameter model I applied was realistic. However, I tested the robustness of the results using alternative two-parameter models with different rates of character evolution. Thus, even though the tree used in the reconstruction was not fully resolved and my model of character evolution may not hold true, my conclusions represent the most reasonable hypothesis to date for evolution of hyphae in the Kingdom Fungi.  3.3 Applications of These Findings and Future Research Directions Ancestral state reconstructions are only as good as the phylogenies that they are based on. My phylogeny of the fungi did not fully resolve evolutionary relationships between basal fungal groups. Therefore, in future, I would like to repeat my analysis on trees with different !  -*!  topologies recovered in other phylogenetic studies to further test the reliability of my conclusions about the evolution of hyphae. However, evolutionary relationships among the deeper clades in the Kingdom Fungi are still largely undefined. I hope that phylogenomics will yield a more robust and comprehensive phylogeny of the Kingdom Fungi in the next few years. An ancestral state reconstruction based on such a phylogeny, would be ideal. Assuming my results are correct, I suspect that hyphae have evolved repeatedly due to the presence of homologous genes in non-hyphal fungi that have repeatedly been co-opted for hyphal growth. Fungi with very different morphologies can share very high genomic similarity as demonstrated by hyphal Ashbya gossypii and the bread yeast, Saccharomyces cerevisiae (Phillipsen et al. 2005). Moreover, budding yeast cells and the rhizoids that anchor chytrid sporangia exhibit polarized growth. I hope to test hypotheses about repeated evolution of hyphae in the future by searching for homologs of genes implicated in hyphal growth in the genomes of early-diverging fungal lineages such as chitin synthases, which form chitin microfilaments in fungi (Alexopoulos et al., 1996; Larson et al., 2011) and septins (Lindsey et al., 2010). Alternatively, hyphae may have arisen several times simply because very little molecular information was needed to develop the machinery underlying sustained polarized growth de novo. After all, hyphae also originated independently in distantly related colorless brown algae, the oomycetes and even plants, animals and bacteria can also grow long, tubular cells (Harris, 2011). At the moment, nearly 210 fungal genomes, not counting those from different isolates of the same species, have already been sequenced or are in progress (http://fungalgenomes.org/wiki/Fungal_Genome_Links). Seven of these sequences represent chytrids including the monoblepharidomycete, Gonapodya prolifera. Now that I have resolved the intergeneric relationships in the Monoblepharidomycetes, one possible future direction would be to examine species-level relationships and develop monoblepharidomycete-specific primers for surveying their distribution and diversity in the environment. Since some Monoblepharidomycetes are soil dwelling, I would be interested to know how well they disperse in soil. The distribution of chytrid fungi in diverse lineages in the fungal tree of life suggests that the earliest fungi to colonize land may have been zoosporic (Sekimoto et al. manuscript under review). Insights into land-based dispersal of flagellated chytrids could again inform studies probing the niches of extinct, ancient fungi.  !  -&!  Given the unusual form of the microtubular cytoskeleton in monoblepharidalean hyphae, I would like to determine the relative importance of microtubules and actin in tip growth. Using microtubule and actin depolymerizing agents on living mycelia, I would compare the extension rates and shape of healthy and chemically treated hyphae. The very intricate system of actin cables in the cytoplasmic periphery of Monoblepharis and Gonapodya suggests that actin could play a much more significant role in growth than in other fungi. As I discussed previously, the tubular shape of fungal cells is largely dependent on actin and hyphae can continue to grow even when they cannot form their usual extensive, parallel microtubular arrays (Steinberg, 2007). However, since microtubules are involved in the transport of endosomes, nuclei, vesicles and mRNA in other fungi (Inoue et al., 1998; Steinberg, 2007), I wish to ascertain whether this is still the case in fungi with an unorthodox arrangement of filamentous microtubules. Also, I would like to confirm whether actin is mostly responsible for vesicle trafficking or if cytoplasmic bulk flow is capable of transporting materials for growth. In addition, this atypical cytoskeleton poses other questions in the field of comparative genomics: Do Monoblepharidomycetes have more proteins with myosin domains? How do the genetic complements of motor proteins in evolutionarily distant fungi compare? Do Monoblepharidomycetes have the specific motor proteins such as kinesin-1 and myosin-5 that co-localize to the Spitzenkörper in filamentous fungi (Steinberg, 2007)? 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