GENOMIC AND MOLECULAR GENETIC ANALYSES OF SECONDARY METABOLISM, TOXIN PRODUCTION, AND IRON HOMEOSTASIS IN COCHLIOBOLUS HETEROSTROPHUS A Dissertation Presented to the Faculty of the Graduate School of Cornell University In Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy by Bradford J Condon August 2013 © 2013 Bradford J Condon GENOMIC AND MOLECULAR GENETIC ANALYSES OF SECONDARY METABOLISM, TOXIN PRODUCTION, AND IRON HOMEOSTASIS IN COCHLIOBOLUS HETEROSTROPHUS Bradford J Condon, Ph.D. Cornell University 2013 Cochliobolus heterostrophus is a model necrotrophic maize pathogen. In 1970, a new and highly virulent race, race T, swept the US east coast, armed with a novel secondary metabolite Host Selective Toxin (HST), T-toxin. The genetic and molecular toolkit developed to characterize the genetics of T-toxin production is combined with genomic resources for C. heterostrophus and related species herein. Comparative genomics of lab and field strains (Tox+ race T and Tox- race O), and other Cochliobolus species, revealed that suites of secondary metabolism nonribosomal peptide synthetase (NRPS) and polyketide synthase (PKS) genes are astoundingly diverse among species but remarkably conserved among isolates of the same species. Strain unique NRPSs and PKSs may produce HSTs, such as PKS1 and PKS2 that biosynthesize T-toxin. These race T specific genes map to the genetically complex Tox1 locus, which is associated with a reciprocal translocation of race O chromosomes. Two candidate reciprocal translocation breakpoint locations were found by whole-genome alignment of race T and O assemblies. Phylogenetic analyses of all ten C. heterostrophus Tox1-affiliated proteins revealed that Didymella zeae-maydis and Leptosphaeria maculans each possess complete but slightly divergent Tox1 loci, while Talaromyces stipitatus possesses a distant ortholog of C. heterostrophus PKS1. Genes at the L. maculans Tox1 locus are clustered, in contrast to the disconnected and scattered nature of C. heterostrophus Tox1. In vitro T-toxin activity assays demonstrated that C. heterostrophus, D. zeae-maydis, and T. stipitatus all display T-toxin-like activity. Broadly conserved secondary metabolite genes produce metabolites with core biological functions. NPS2 and NPS6, encoding NRPSs producing iron-chelating siderophores, are key examples. In C. heterostrophus, deletion of these results in loss of extracellular siderophore biosynthesis, attenuated virulence, hypersensitivity to oxidative and iron-depletion stress, and defective sexual spore development. In addition to siderophores, fungi can also utilize high affinity Reductive Iron Assimilation (RIA) mechanisms to acquire iron. Characterization of genetic mutants found that RIA is dispensable for C. heterostrophus. When RIA/siderophore polymutants were created, however, basic and severe morphological defects occurred. The role of HapX, an iron-responsive transcription factor, was also investigated. HapX is required for full virulence, iron-depletion tolerance, and sexual development. BIOGRAPHICAL SKETCH Bradford Joseph Condon was born to Thomas and Lynn Condon on December 19, 1984 in Danbury, Connecticut. He was raised, along with his older sister Brett, in the town of Redding, CT. Bradford enjoyed a wide variety of activities growing up, including classical guitar, choir, basketball, cross country running, and skiing. His love of animals fostered an interest in biology, which eventually found him at Oberlin College majoring in biology, along with religion. He started at Cornell University in the Department of Plant Pathology and PlantMicrobe Biology in the fall of 2007, and joined Dr. B. Gillian Turgeon’s laboratory in the spring of 2008. His education in Ithaca has spanned many biological disciplines, and he is thankful for the support of his family, advisor, committee, and friends through tumultuous years of delayed adulthood. iii To Mom and Dad With love and gratitude iv ACKNOWLEDGMENTS This Dissertation was made possible thanks to the hard work and support of many people. First and foremost I am thankful to my advisor, Gillian Turgeon, for her guidance, support, patience, and friendship in my time here. I owe my success at Cornell to her fierce dedication and loyalty. Many members of the Turgeon lab group have provided me with training and friendship over the years. In particular I would like to thank Dongliang Wu, Jinyuan Liu, Kathryn Bushley, Ning Zhang, Nur Ain Izzati Mohd, Marisa Queiroz, Benjamin Horwitz, and Sung-Hwan Yun. I would also like to thank my minor committee members Kathie Hodge and Magdalen Lindeberg, for their input and support. Thanks to Derina Samuel for her hard work and support through the Center for Teaching Excellence. Thanks to Alan Collmer for his dedication as an instructor, for serving as a temporary committee member, and hosting my laboratory rotation. Thanks also to Maria Harrison for hosting my final laboratory rotation. Thank you to Barry Scott at Massey University for hosting my NSF-EAPSI summer in New Zealand. Thanks also to Michael Milgroom and Stewart Gray, who both served as DGS during my stay, and thanks to Margaret Haus, Dawn Dailey O’Brien, Jackie Armstrong, and Alicia Caswell who have served as GSR. Thanks to George Hudler, former department chair and instructor, and Bill Fry, for serving as department chair and providing generous support. For his excellent photographic work I am grateful to Kent Loeffler. Thanks to all of the administrative staff who have provided a friendly face and assistance on countless occasion, especially Carol Fisher, Tracy Holdridge, Jackie Armstrong, and Andrea Gilbert. While at Cornell I was supported by the Presidential Life Science Fellowship, NSF- EAPSI, departmental teaching assistantships, and Dr. Turgeon’s program, for which I am very grateful. v TABLE OF CONTENTS Page LIST OF FIGURES……………………………………………………………………………..viii LIST OF TABLES……………………………………………………………………………….xii Chapter I. Introduction………………………………………………………………………..1 Summary…………………………………………………………………………………14 References………………………………………………………………………………..15 Chapter II. Comparative genome structure, secondary metabolite, and effector coding capacity across Cochliobolus pathogens…………………………………………24 Abstract………………………………..…………………………………………………25 Introduction………………………………………………………………………………26 Results……………………………………………………………………………………32 Discussion………………………………………………………………………………..76 Materials and Methods…………………………………………………………………...87 References………………………………………………………………………………..98 Chapter III. Genomics-driven investigation of T-toxin production in C. heterostrophus…...113 Abstract………………………………..………………………………………………..114 Introduction…………………………..…………………………………………………115 Materials and Methods…………….....…………………………………………………130 Results and Discussion…………………………..……………………………………..143 Conclusions……………………..………………………………………………………184 References………………………………………………………………………………186 Chapter IV. The role of iron acquisition and homeostasis in C. heterostrophus cellular biology and pathobiology………………………………………………………………..198 Abstract…………………………………………………………………………………199 Introduction……………………………………………………………………………..200 Materials and Methods………………………………………………………………….218 vi Results and Discussion………………………………………………. ………………..231 References………………………………………………………………...…………….269 Appendix………………………………………………………………...……………………...287 Supplemental Figures…………………………………………………………………...288 Supplemental Methods………………...………………………………………………..338 References………………………………………………………………………………345 vii LIST OF FIGURES Figure Page II.1. C. heterostrophus RFLP sequences anchor sequenced scaffolds to the genetic map……………………………………………………………………………………….35 II.2. C. heterostrophus unique regions, secondary metabolite and small secreted protein encoding genes are distributed throughout the genome……….....……………………....42 II.3. Cartoon of cross-species phylogenomic analyses of individual AMP binding domains from NRPS proteins..…………………………………………………………………….47 II.4. NPS2 is an example of a highly conserved Dothideomycete NRPS..…………………...49 II.5. The NPS1/NPS3/NPS13 expansion group of NRPS AMP domains..…………………...52 II.6. NPS1, NPS3 and NPS13 are examples of NRPS proteins encoded by highly recombinogenic and expanded NPS genes…………………………………………........54 II.7. Genomic organization of the scaffold associated with the VHv1 locus conferring high virulence of pathotype 2 isolate ND90Pr to barley cv. Bowman compared to the corresponding region in pathotype 0, isolate ND93-1…………………………………...59 II.8. S. turcica has an ortholog of the C. carbonum NRPS HTS1 responsible for HC- toxin biosynthesis………………………………………………………………………………62 II.9. C. victoriae has an ortholog of A. fumigatus GliP responsible for gliotoxin production………………………………………………………………………………..64 II.10. Cartoon of cross-species phylogenomic analyses of individual ketosynthase domains from PKS proteins……………………………………………………………………….68 II.11. The two PKSs responsible for T-toxin production by race T of C. heterostrophus are unique to race T………………………………………………………………………….70 III.1. Tox1 maps to a reciprocal translocation between race O and race T chromosomes……122 III.2. Tox1 scaffolds and genes in all sequenced C. heterostrophus race T strains……….….145 III.3. Graphical representation of visual search strategies for race T unique sequence…........147 III.4. Candidate sequences for the location of race O chromosome 6 and 12 breakpoints.......152 viii III.5. A PCR product spanning the proposed Tox1 related breakpoint on chromosome 12 can be amplified from race O, but not race T, DNA……………………………………….......154 III.6. Phylogenetic relationship of ChPks1 and relatives……………………………………..156 III.7. Phylogenetic relationship of ChPks2 and relatives……………………………………..157 III.8. Talaromyces stipitatus displays T-toxin-like activity in an in vitro microbial assay…..164 III.9. L. maculans strains in the PPP-MB collection do not demonstrate T-toxin activity in in vitro assays……………………………………………………………………………...165 III.10. The decarboxylase Dec1 is unique to C. heterostrophus, D. zeae-maydis, and L. maculans………………………………………………………………………………..168 III.11. The 3-hydroxacyl-CoA dehydrogenase Lam1 is unique to C. heterostrophus, D. zeae- maydis, and L. maculans………………………………………………………………..168 III.12. The short chain reductase Oxi1 is found in many Tox1- taxa, and is duplicated in D. zeae- maydis…………………………………………………………………………………..169 III.13. The protein of unknown function Tox9 appears unique to C. heterostrophus, D. zeae- maydis, and L. maculans………………………………………………………………..170 III.14. The protein of unknown function Tox10 does not form a strong phylogenetic relationship across C. heterostrophus, D. zeae-maydis, and L. maculans…………………………...171 III.15. The reductase ChRed1 is found only in C. heterostrophus, D. zeae-maydis, and Macrophomina phaseolina.…………………… ………………………………………172 III.16. The ChRed2 and Red3 reductase family has members in L. maculans and D. zeae- maydis. …………………………………………………………………………………172 III.17. The full C. heterostrophus Tox1 locus can be found in D. zeae-maydis and L. maculans, but not T. stipitatus…………………..…………………………………………………174 III.18. The L. maculans strain JN3 genome contains all known C. heterostrophus Tox1 genes.…………………………………………………………………………………...177 III.19. C. heterostrophus abc18 mutants produce and export T-toxin. ………………………..179 III.20. C. heterostrophus tox10 mutants do not produce T-toxin.……………………………..181 ix III.21. C. heterostrophus race T WT and abc18 mutants, but not tox10 mutants, produce lesions typical of T-toxin production on T-cytoplasm corn. …………………………………...182 III.22. tox10 mutants are not reduced in virulence on N-cytoplasm corn. …………………….183 IV.1. Fungi possess three modes of high affinity iron acquisition.…………..……………….206 IV.2. Plating tetrads on different media allows rapid identification of desired polymutants...233 IV.3. Impaired growth of double and triple mutants on CMX….…………………………….236 IV.4 Impaired growth of double, triple, and quadruple mutants on minimal medium………238 IV.5. Observation of nps6ftr1 and nps2nps6ftr1 conidia in liquid culture shows abnormal colony growth…………………………………………………………………………..239 IV.6. Ferrous salt, ferric salt, and ferric citrate supplement nps2nps6ftr1 mutant growth…...241 IV.7. nps6ftr1 and nps2nps6ftr1 are impaired in asexual spore production………………….242 IV.8. nps2nps6ftr1 growth is restored by proximity to WT and nps2, but not nps6 or nps2nps6 cultures………………………………………………………………………………….244 IV.9. abc6 mutants overfeed nps2nps6ftr1 triple mutants in an NPS6-dependent manner…..246 IV.10. Iron mutants are sensitive to oxidative and iron stress…………………………………247 IV.11. Iron mutants are progressively sensitive to oxidative and iron stress…………………..248 IV.12. Virulence of iron mutants is progressively reduced compared to WT…………………250 IV.13. Iron mutants are progressively reduced in virulence with or without supplemental iron……………………………………………………………………………………...251 IV.14. DAB staining is reduced in maize leaves inoculated with iron mutants………………..253 IV.15. Iron mutants are progressively impaired in colonizing maize leaves…………………..255 IV.16. Host autoflorescence accompanies fungal entry………………………………………..256 IV.17. hapX mutants are sensitive to iron, but not oxidative, stress…………………………...258 IV.18. hapX mutants feed nps2nps6ftr1 siderophores as WT………………………………….260 IV.19. hapX mutants are impaired in ascospore development…………………………………261 IV.20. hapX mutant crosses do not produce complete tetrads…………………………………262 x IV.21. All hapX X WT progeny are WT (Hygromycin B sensitive) ………………………….263 IV.22. hapX mutants are male and female fertile in sexual crosses……………………………264 IV.23. hapX is reduced in virulence on maize…………………………………………………266 S1. Tox1 maps to a reciprocal translocation breakpoint on race O chromosomes 6 and 12………………………………………………………………………………………..288 S2. Generalized schematic for knockout transformation construct generation and confirmation of integration……………………………………………………………..289 S3. Genetic distance correlates with physical distance on the C. heterostrophus map…….290 S4. A C. heterostrophus dispensable chromosome is present in some but not all C. heterostrophus strains…………………………………………………………………..290 S5. C. sativus SSR sequences anchor sequenced scaffolds to the genetic map…………….291 S6. Analysis of the mating type region in Cochliobolus spp. and S. turcica……………….295 S7. Maximum likelihood tree of NRPS AMP-binding (AMP) domains identified using Augustus………………………………………………………………………………..296 S8. Maximum likelihood tree of PKS ketosynthase (KS) domains identified using Augustus………………………………………………………………………………..303 S9. Quantification of spot blotch disease induced by the C. sativus wild type and mutant (Δ115356) on barley cv. Bowman.……………… …………………………………….309 S10. Quantitative real-time PCR analysis of S. turcica PKS gene (protein ID 161586) during infection of maize cultivar W64A-N……………………………………………310 S11. Complete ChPks1 and ChPks2 phylogenetic tree………………………………………311 S12. Complete decarboxylase Dec1 phylogenetic tree………………………………………319 S13. Complete Lam1 phylogenetic tree……………………………………………………...322 S15. Complete Tox9 phylogenetic tree………………………………………………………328 S16. Complete reductase phylogenetic tree………………………………………………….331 xi LIST OF TABLES Table Page II.1. Cochliobolus and Setosphaeria-host interaction biology……………….……………….27 II.2. C. heterostrophus race O strain C5, race T strain C4, C. sativus and S. turcica genome statistics………….…………………….…………………….…………………….……..32 II.3. Statistics for short read re-sequenced Cochliobolus genomes………….………………..33 II.4. SNPs between C. heterostrophus (Ch) strain C5 and other strains and species…………37 II.5. SNPs between C. sativus strains ND90Pr and ND93-1………….………………………38 II.6. SNPs between C. carbonum and other species………….…………………….…………39 II.7. Conservation of C. heterostrophus strain C5 nonribosomal peptide synthetases in other Cochliobolus strains and species………….…………………….……………………….46 II.8. Total and unique Cochliobolus NRPS and PKSs………………………………………..57 II.9. Conservation of C. heterostrophus strain C5 polyketide synthases in other strains and species……………………………………………………………………………………67 II.10. Comparative small secreted protein candidate effector inventories……………………..73 II.11. C. heterostrophus strain C5 SSP candidate effector analysis……………………………74 III.1. Strains used in this chapter……………………………………………………………..130 III.2. Primers………………………………………………………………………………….136 III.3: Panseq subtracted race T scaffolds..……………………………………………………148 III.4. Species included in PKS KS domain tree………………………………………………158 III.5. PKS inventories and distribution patterns………………………………………………160 III.6. %GC content in PKS transcripts………………………………………………………..162 IV.1. C. heterostrophus strains used in this study……………………………………………219 IV.2. Primers used in this study……………………………………………………………....223 IV.3. Crosses and selfs set up in this study…………………………………………………...225 IV.4. Genes studied in this chapter…………………………………………………………...231 xii ! Chapter I Introduction !1 ! A. Introduction Fungi are beautiful, marvelous, adaptable organisms. They form a kingdom of alien shapeshifters capable of inhabiting any imaginable niche given time to work their evolutionary magic. The hallmark plasticity of fungi is reflected in each species’ genome, which births a cataclysm of biological possibilities. These traits feed their capacity for mischief and destruction as principal agents of plant disease. For my PhD thesis, I have focused primarily on the fungal genus Cochliobolus, with particular attention to Cochliobolus heterostrophus, the causal agent of Southern Corn Leaf Blight (SCLB). There are three chapters of experimental results: (II) Comparative genome structure, secondary metabolite, and effector coding capacity across Cochliobolus pathogens, (III) Genomics-driven investigation of T-toxin production in C. heterostrophus, and (IV) The role of iron acquisition and homeostasis in C. heterostrophus cellular biology and pathobiology. Each chapter addresses one particular facet of C. heterostrophus as a necrotrophic pathogen of corn. This introduction sets the stage, providing the historical and biological context of key members of the genus Cochliobolus, and the conceptual context for fungal phytopathogenesis in the world of molecular plant pathology. 1. Cochliobolus heterostrophus and related species Cochliobolus (anamorph Bipolaris/Curvularia) spp. are young, closely related species (<20 MYA [1]) which makes them ideal for comparative studies. For example, to determine fundamental genome biology that defines a pathogenic lifestyle, or what determines hostspecificity, we take advantage of the fact that some Cochliobolus spp. are highly aggressive pathogenic species specific to their cereal hosts, while others are more general pathogens of !2 ! cereals, and still others are saprobes. To address mechanistic differences in reproductive lifestyle, we take advantage of the fact that some Cochliobolus spp. require a partner for sexual reproduction, some are capable of self-mating and thus do not require a partner, while still others have no known sexual stage. i. Basic biology The genus Cochliobolus divides phylogenetically into two groups, each associated with a distinct anamorphic stage. The first group, which encompasses the majority of known aggressive pathogenic species with significant impact on host crops, has a Bipolaris asexual stage while the second group has a Curvularia asexual stage [2]. Under the ‘one name one fungus’ initiative, the Bipolaris name may be adopted to comply with the International!Code!of!Nomenclature!for! Algae,!Fungi,!and!Plants [3]. Most contemporary genetic, molecular, and genomic research on virulence determinants and reproductive development of the group has employed this designation. The first group of species includes the necrotrophic corn pathogens Cochliobolus heterostrophus and Cochliobolus carbonum, the oat pathogen, Cochliobolus victoriae, the rice pathogen, Cochliobolus miyabeanus, the sorghum pathogen, Bipolaris sorghicola and the sugarcane pathogen, Bipolaris sacchari. Cochliobolus lunatus, also a pathogen of sorghum, falls in the second group. The only species with a known hemibiotrophic lifestyle is the generalized cereal and grass pathogen, Cochliobolus sativus which belongs to the first group. Some of these species, i.e., C. lunatus can act as opportunistic human pathogens. !3 ! ii. Historical importance The best-studied necrotrophic Cochliobolus spp. are notorious for their ability to evolve novel, highly virulent races producing Host Selective Toxins (HSTs) associated with the capacity of their producers to cause diseases on cereal crops that were bred, inadvertently, for susceptibility to the HST-producing pathogen [4,5]. For example, in 1970, race T, a novel race of C. heterostrophus (Bipolaris maydis) caused a major epidemic of Southern Corn Leaf Blight (SCLB) which destroyed more than 15% of the maize crop on the US eastern seaboard [6]. Race T is genetically distinct from race O in that it uniquely carries genes for biosynthesis of T-toxin, an HST essential for high virulence on Texas male sterile cytoplasm (Tcms) corn [5]. This story is related in its entirety in Chapter III of this work. C. victoriae (Bipolaris victoriae), causal agent of Victoria Blight, produces the chlorinated cyclic pentapeptide HST victorin, rendering it highly virulent to oats carrying the dominant Vb allele [7]. Varieties of this type were introduced into the US from Uruguay [7] because they carry the Pc-2 gene for resistance to crown rust caused by Puccinia coronata [8]. The Vb-associated trait, susceptibility to C. victoriae, and the Pc-2-associated trait, resistance to P. coronata, cannot be separated genetically [9]. C. victoriae caused widespread destruction in the 1940’s on Pc-2 oats, because, as is now well-documented, victorin elicits Pc-2-dependent Programmed Cell Death (PCD). Thus, like the C. heterostrophus T-toxin/Tcms case, the monoculture of Victoria oats carrying Pc-2 was the perfect milieu for attack by C. victoriae. Recent work with Arabidopsis revealed a NB-LRR-type resistance protein (LOV1), guarding a thioredoxin protein target (TRX-h5), that when activated confers susceptibility to C. victoriae and victorin [10,11]. Victorin thus acts by co-opting effector triggered defenses against the biotroph, P. coronata, to promote susceptibility to a necrotroph. !4 ! In contrast to the dominant plant host genes required for susceptibility to C. heterostrophus and C. victoriae, susceptibility to Northern Corn Leaf Spot caused by C. carbonum (Bipolaris zeicola) is conferred by a homozygous recessive maize gene(s) [12,13]. C. carbonum race 1 produces the cyclic-tetrapeptide HST, HC-toxin, which is specifically active, as is the fungus itself, against corn with the naturally occurring mutant genotype hmhm [5,14,15]. A mutation in the Hm1 gene is thought to have occurred prior to 1930 in a maize line in Kansas under low disease pressure [16]. When this germplasm was moved to the cornbelt in the 1930s, disease resulted. Hm1 (and Hm2) encode carbonyl reductases which inactivate the toxin [12]; hmhm lines cannot inactivate the toxin, and are therefore sensitive. The site of action of HCtoxin in susceptible corn is histone deacetylase; it is hypothesized that HC-toxin acts to promote infection of maize of genotype hm1hm1 by inhibiting this enzyme, resulting in accumulation of hyperacetylated core histones. This then alters expression of genes encoding regulatory proteins involved in plant defense [17,18]. C. carbonum races 2 and 3 do not produce the toxin. C. sativus (Bipolaris sorokiniana), a hemibiotroph and less specialized cereal pathogen, causes diseases of roots (Common Root Rot), leaves (Spot Blotch), and spikes (Black Point or Kernel Blight) of cereals (mainly barley and wheat) [19,20], but also attacks grasses, including switchgrass (Panicum virgatum L.) [21-23]. Three C. sativus pathotypes (0, 1, and 2) have been described [24] based on differential virulence patterns on three barley genotypes (ND5883, Bowman, and NDB112). Pathotype 0 isolates show low virulence on all three barley genotypes. Pathotype 1 isolates show high virulence on ND5883 but low virulence on other barley genotypes. Pathotype 2 isolates show high virulence on Bowman but low virulence on ND5883 and NDB112. Genetic analysis and molecular mapping indicates that a single locus, VHv1, controls high virulence of the pathotype 2 isolate ND90Pr on Bowman [24,25]. The VHv1 locus !5 ! is unique to pathotype 2 and encodes two nonribosomal peptide synthetases (NRPSs), one of which when deleted, drastically reduces virulence of pathotype 2 on cultivar Bowman [26]. This is one of only a few examples of a secondary metabolite, not a small secreted protein, acting as an effector on a specific cultivar of the producer’s host. Performance of the mutant on other cultivars has not been tested, nor has the corresponding NPS gene been introduced to other races, thus it is unclear at this point whether this is clearly a hemibiotrophic type of interaction or is an HST-type interaction. C. miyabeanus (Bipolaris oryzae) is the causal agent of Brown Spot of rice which contributed, along with a cyclone and tidal waves, to the Bengal rice famine of 1942/1943 that resulted in starvation of more than two million people [27]. The interaction between rice and C. miyabeanus is inadequately understood from the perspective of genetic and molecular mechanisms, although it has been reported that, like other Cochliobolus species, the fungus utilizes phytotoxins to trigger host cell death [28]. No HST has ever been reported. iii. Resources developed for genetic and molecular manipulation of Cochliobolus Cochliobolus spp. are easily grown in culture, produce abundant asexual spores and can be stored for long periods of time in glycerol or silica gels [29]. They have an efficient sexual stage readily produced in the laboratory in three weeks [30], and are easily transformed [31]. Targeted gene deletion using PCR fragments is highly efficient [31-33]. Targeted gene deletion using protoplast suspensions yields properly integrated transforming DNA in transformants at close to 100% efficiency [31]. Chromosomes can be resolved using pulsed-field gel electrophoresis [34,35]. !6 ! 2. C. heterostrophus as a necrotroph A biotroph is a pathogen that relies on the vitality of its host to acquire nutrients, while a necrotroph actively thrives on dead and dying host cells [36,37]. We now understand that many pathogens are hemibiotrophs, having an initial biotrophic phase, with a distinct shift towards necrotrophy late in infection [37,38]. C. heterostrophus, C. carbonum, C. miyabeanus, and C. victoriae are all textbook necrotrophs, with no indication of a biotrophic phase. C. sativus, on the other hand, is now classified as a hemibiotroph [26]. The current understanding of hostnecrotrophic defense is reviewed below to provide necessary background for subsequent chapters dealing with C. heterostrophus in its interactions with maize. i. Overview of plant host defenses Plant defense responses against any pathogen falls roughly into two categories- innate immunity/PAMP (Pathogen Associated Molecular Pattern)-Triggered Immunity (PTI), and Effector-Triggered Immunity (ETI) [39]. PTI and ETI are effective against biotrophs and hemibiotrophs, while defense against necrotrophs falls under innate immunity/PTI. The role of ETI for necrotrophs is uncertain at this point. Sharing a defense pathway with non-pathogens, and lacking the intricate gene for gene interactions of ETI, has lead to the belief that necrotrophs are less sophisticated pathogens than biotrophs. This view is slowly changing as researchers acknowledge that necrotroph-host interactions are more complex than previously thought [40]. Signaling for these pathways is overseen by a complex network of hormones. Against a necrotroph, ethylene (ET) and jasmonic acid (JA) are the dominant hormones promoting PTIbased defenses [41]. Many PTI-based immune responses require JA and ET, and exogenous preapplication of high levels of JA or ET can confer resistance to necrotrophs, while ABA can !7 ! increase susceptibility to (some, but not all) necrotrophs [42,43]. Arabidopsis hormone mutants, such as the ethylene insensitive ein2, tell a similar tale of resistance and susceptibility [44]. In fact, ET, JA, SA, ABA work cohesively, along with other hormones such as auxin and gibberellins, to facilitate PTI or ETI [41]. Levels of ET, JA, SA, and ABA will all increase in response to Botrytis cinerea infection, and some pathogens may disrupt the balance by producing their own hormones or hormone analogs [45]. A final important note is that much of our knowledge regarding plant defenses comes studies with Arabidopsis. It follows that these models are generalizations based on interactions between Arabidopsis and a small set of fungal necrotrophs, particularly B. cinerea, but also Sclerotinia sclerotium, Alternaria brassicicola, and Leptosphaeria maculans (which is now classified as a hemibiotroph) [36]. The interactions between maize and C. heterostrophus may in some cases be quite different. ii. Innate immunity There are many layers to innate immunity, and a brief synopsis is provided here. The structural integrity of the plant (both the cuticle and cell wall) must first be compromised [46,47]. After attaching and germinating, fungal spores gain entry into the plant by developing appressoria (penetrating infection structures) which generate mechanical pressure and/or by secreting hydrolytic enzymes [46,47]. The cell wall (composed of cellulose, hemicellulose, and pectin [48,49]) is degraded by the secretion of Cell Wall Degrading Enzymes (CWDEs), including oxidases, cutinases, xylanases, lipases, laccases, cellulases, proteases, and pectinases (such as polygalacturonases and endopolygalacturonases) [14,36,50]. Oxalic acid may also be secreted, to acidify the enzymatic working environment and increase enzyme efficacy [51]. Necrotrophs tend to have larger repertoires of CWDEs than biotrophic pathogens, and these !8 ! expanded inventories grant CWDEs functional redundancy [52,53]. This makes demonstrating clear roles for these enzymes in pathogenesis through molecular/genetic studies difficult. Plants can fortify the cell wall by cross-linking glycoproteins, or depositing callose and lignin [37]. Polygalacturonase inhibiting proteins (PGIPs) are also embedded within the cell wall to counteract fungal pectolytic enzymes [50,54]. PGIPs block cell wall degradation directly, and also enhance accumulation of oligogalacturonides [50,55], one of several types of Damage-Associated Molecular Patterns (DAMPs). DAMPs are host molecules generated in the wake of an invading pathogen that the plant recognizes, spurring various defenses (see below) [56,57]. A plant’s immune response is not all defensive: they also have offensive mechanisms to destroy invaders. The plant can produce Pathogenesis-Related (PR) proteins, a hallmark of ET/JA mediated PTI [36,58]. There are 17 PR families, including antifungal peptides, protease inhibitors, defensins, and other small peptides [59,60]. Similarly, plants produce metabolites with antimicrobial properties called phytoalexins (or phytoanticipins if preformed) [36,61]. Successful pathogens may, in turn, detoxify or export these phytotoxic compounds, as is the case with Nectria haematococca and the phytoalexin pisatin [62,63]. It must be mentioned that reactive oxygen species (ROS) are an essential component of initial plant defenses [64,65]. There is a biphasic oxidative burst in the apoplast upon successful recognition of plant pathogens that is mediated by plant NADPH oxidases (also known as respiratory burst oxidases in plants) [64-67]. This oxidative burst is essential for mediating defense signaling, and also eliciting the hypersensitive response (HR) (see below). ROS also can have direct antimicrobial effects, and they can lignify the cell wall, and cross-link cell wall !9 ! glycoproteins [65,66]. The role of ROS in plant-microbe interactions is discussed at length in Chapter IV. The above defenses that are not pre-formed must be triggered, released, or otherwise expressed in response to the pathogen. For this to happen, the plant must recognize the presence of a pathogen. Plant perception of pathogen-altered homeostasis can trigger innate immune responses, mediated by shifts in cellular status or intracellular signaling [57,68]. Host-derived DAMPs released by PGIPs and other mechanisms may also alert the host. Finally, the plant may recognize components of the pathogen itself in the form of PAMPs or Microbe Associated Molecular patterns (MAMPs), which in the case of fungi is often chitin. The Stagonospora nodorum and Pyrenophora tritici-repentis proteinaceous toxin ToxA is enough to stimulate production of phytoalexins in wheat, suggesting specific necrotrophic virulence factors may induce resistance responses [69]. iii. Effectors and host selective toxins The defenses described are extremely effective, and indeed, “Most plants are resistant to most plant pathogens” [70]. Bacterial and fungal pathogens, particularly hemibiotrophs and biotrophs, have therefore evolved effector proteins to overcome PTI [39,71]. These small, secreted proteins are typified by those transferred by the type-III secretion system in Pseudomonas syringae to the host cell to act on specific plant cellular targets [72,73]. Biotrophic and hemibiotrophic fungi, too, transfer small proteins that interact with intracellular host targets, although the delivery mechanism is not consistent across fungal species (and still hotly debated) [74,75]. Plants, in turn, possess a retaliatory defense system, effector triggered immunity (ETI), to detect effectors and their handiwork, and stop infection by triggering the hypersensitive response (HR), killing the cell resulting in complete, qualitative resistance (hence the original ! 10 ! ‘avirulence’ nomenclature for effectors) [76]. Biotrophs then evolve more effectors to prevent HR, starting a perpetual back and forth between plant and pathogen typical of the so-called “zigzag” model of susceptibility and resistance [39]. In contrast to the qualitative resistance of HR and ETI, quantitative resistance provides small boosts to resistance, and is functionally very different, and tied towards a more effective PTI/innate immunity [77]. There are no reports of R genes conferring qualitative resistance against necrotrophs, although qualitative resistance pathways still operate in the presence of HSTs [45]. This is because ETI relies on HR, which is not thought to be an effective strategy against necrotrophs [37,39]. In fact, necrotrophs seem to manipulate host encoded R proteins to trigger HR, to the benefit of the pathogen. For example, the product of the Arabidopsis RPW8 gene is a resistance protein against powdery mildew [78], but mediates susceptibility against both B. cinerea and A. brassicicola [79]. Similarly, the Arabidopsis LOV1 gene encodes a typical CC-NBS-LRR R protein [9,10], and its presumed ortholog in oats, PC2, encodes a protein conferring resistance to Puccinia coronata while simultaneously conferring susceptibility to C. victoriae [80]. This is because C. victoriae produces the HST victorin, which specifically targets LOV1/PC2, which triggers HR and promotes disease [9]. HSTs, such as C. heterostrophus’s T-toxin and C. carbonum’s HC-toxin, do not target R proteins. Furthermore, not all toxins are secondary metabolites. S. nodorum and P. triticirepentis produce the HST ToxA, which promotes disease in the presence of the product of the dominant susceptibility gene Tsn1 in wheat. ToxA is not a secondary metabolite, but a ribosomally encoded protein [81-83]. The argument has been made that HSTs and nonselective toxins produced by necrotrophs are truly effectors, as they are molecules which interact with host molecular targets to promote ! 11 ! disease [81]. Should the definition of an effector be limited to small secreted proteins that act as virulence effectors in the host cell, or should it include any fungal molecule, including toxins or pectolytic enzymes, acting outside the fungus to promote disease? The answer depends on if the true identify of a classical ‘avirulence’ protein effector stems from its composition (a small and cysteine-rich secreted protein), its activity (promoting virulence, but in the presence of a corresponding R protein resulting in avirulence), or its localization (inside the host cell). While it is true that effectors are always assumed to be playing a positive role in virulence (when they are not “caught” by an R-protein), much like an HST, broad range toxin, or pectolytic enzyme, they also are a signature of a very specific battle, conforming to the rules of the “zig-zag” model. Necrotrophic “effectors”, such as ToxA in S. nodorum, result in susceptibility in the presence of a host susceptibility factor, and the interaction is termed an inverse gene-for-gene interaction [69,84]. The term effector replaced avirulence protein precisely because they were thought to promote disease in the absence of an R gene: why, then, include proteins which promote disease only in the presence of a host factor? Does labeling necrotrophic virulence factors and toxins as effectors bring together to two types of molecules with very different consequences for disease? It is therefore my opinion that effectors are most effective and meaningful as a label when reserved for secreted proteins that play a role in ETI. A biotrophic effector in the presence of an R gene product will result in resistance. That same effector, secreted by a necrotroph, targeting the same R protein, will result in disease rather than resistance, and is therefore adequately described as a HST. These terms are structured so that we can understand how a host and a pathogen interact. How can effector be a useful term if it describes anything secreted by a fungus, that may positively or negatively impact disease? Without a tight definition, “effector” ! 12 ! will likely go the way of “virulence factor”, describing anything that, when removed or disrupted, reduces an observed in planta phenotype, even if the reduced phenotype is not related to plant-microbe interactions at all (for example genes required for polar hyphal growth). If necrotrophic HSTs and small cysteine-rich secreted proteins are to be termed ‘effector’, than a defining characteristic that separates them from the many other secreted factors must be agreed upon. Whatever terminology is used to describe them, it is clear that necrotrophs utilize both secondary metabolites and proteins to promote disease in both a specific and generalized manner. The offensive strategy of a necrotroph has become more refined over the years, and more subtleties will likely be revealed in the future. Combining our understanding of plant innate immunity and necrotrophic defense may ultimately be misleading. ! 13 ! B. Summary To bypass host defenses, a necrotrophic foliar pathogen must break into the leaf, survive pre-formed and responsive defenses, and trigger the death of its host. It must also survive the highly cytotoxic environment generated during plant defense responses and cell death. Finally, it must acquire nutrients and propagate to new hosts (typically by asexual or sexual production). HSTs are a key component of the Cochliobolus offensive strategy, and they are therefore a primary focus of the analyses presented in Chapters II and III. 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Manning VA, Chu AL, Steeves JE, Wolpert TJ, Ciuffetti LM (2009) A host-selective toxin of Pyrenophora tritici-repentis, Ptr ToxA, induces photosystem changes and reactive oxygen species accumulation in sensitive wheat. Mol Plant Microbe In 22: 665– 676. 83. Ciuffetti LM, Manning VA, Pandelova I, Betts MF, Martinez JP (2010) Host-selective toxins, Ptr ToxA and Ptr ToxB, as necrotrophic effectors in the Pyrenophora triticirepentis-wheat interaction. New Phytol 187: 911–919. 84. Oliver RP, Solomon PS (2010) New developments in pathogenicity and virulence of necrotrophs. Curr Opin Plant Biol 13: 415–419. ! 23 Chapter II Comparative genome structure, secondary metabolite and effector coding capacity across Cochliobolus pathogens* *This chapter is reprinted under a creative commons license from Condon, B. J., Leng, Y., Wu, D., Bushley, K. E., Ohm, R. A., Otillar, R., et al. 2013. Comparative Genome Structure, Secondary Metabolite, and Effector Coding Capacity across Cochliobolus Pathogens. PLoS Genet. 9:e1003233 24 Abstract The genomes of five Cochliobolus heterostrophus strains, two Cochliobolus sativus strains, three additional Cochliobolus species (Cochliobolus victoriae, Cochliobolus carbonum, Cochliobolus miyabeanus) and closely related Setosphaeria turcica were sequenced at the Joint Genome Institute (JGI). The datasets were used to identify SNPs between strains and species, unique genomic regions, core secondary metabolism genes, and small secreted protein (SSP) candidate effector encoding genes with a view towards pinpointing structural elements and gene content associated with specificity of these closely related fungi to different cereal hosts. Whole genome alignment shows that three to five percent of each genome differs between strains of the same species, while a quarter of each genome differs between species. On average, SNP counts among field isolates of the same C. heterostrophus species are more than 25X higher than those between inbred lines and 50X lower than SNPs between Cochliobolus species. The suites of nonribosomal peptide synthetase (NRPS), polyketide synthase (PKS), and SSP-encoding genes are astoundingly diverse among species but remarkably conserved among isolates of the same species, whether inbred or field strains, except for defining examples that map to unique genomic regions. Functional analysis of several strain-unique PKSs and NRPSs reveal a strong correlation with a role in virulence. 25 A. Introduction The filamentous ascomycete genus Cochliobolus (anamorph Bipolaris/Curvularia) is comprised of more than forty closely related, often highly aggressive, pathogenic species with particular specificity to their host plants. All members of the genus known to cause serious crop diseases fall in a tight phylogenetic group suggesting that a progenitor within the genus gave rise, over a relatively short period of time (<20 MYA, Ohm et al., [1]) to the series of distinct species [2], each distinguished by unique pathogenic capability to individual types of cereal (Table II.1). Aggressive members include the necrotrophic corn pathogens Cochliobolus heterostrophus and Cochliobolus carbonum, the oat pathogen, Cochliobolus victoriae, the rice pathogen, Cochliobolus miyabeanus, the sorghum pathogen, Bipolaris sorghicola, the sugarcane pathogen, Bipolaris sacchari, and the hemibiotrophic generalized cereal and grass pathogen, Cochliobolus sativus. All of the known Cochliobolus pathogens are classified as necrotrophs, except for C. sativus, which, although previously classified as such, has more recently been described as a hemibiotroph [3]. Many necrotrophic Cochliobolus spp. and related taxa (e.g., Pyrenophora tritici-repentis, Stagonospora nodorum) are notorious for their ability to evolve novel, highly virulent, races producing Host Selective Toxins (HSTs) and their concomitant capacity to cause diseases on cereal crops that were bred, inadvertently, for susceptibility to the HST-producing pathogen [4,5]. For example, in 1970, race T, a previously unseen race of C. heterostrophus (Bipolaris maydis) caused a major epidemic [Southern Corn Leaf Blight (SCLB)], destroying more than 15% of the maize crop [6]. Before 1970, C. heterostrophus was known as an endemic 26 Table II.1. Cochliobolus and Setosphaeria-host interaction biology Speciesa Host/tissue Disease HST? HST/Effector Pathogen Target Lifestyle Ch race O corn/leaves Southern Corn Leaf ? - necrotroph (strains C5, Blight Hm540) Ch race T corn with Southern Corn Leaf T-toxin URF13 protein necrotroph (strains C4, Tcmsb/leaves Blight Hm338, PR1x412) Cc race 1 (strain hm1hm1c corn/ Northern Leaf Spot HC-toxin histone necrotroph 26-R-13) leaves deacetylase Cv (strain FI3) Vbd oats/crown Victoria Blight victorin LOV1 necrotroph Cm (strain rice/leaves Brown Spot ? - necrotroph WK1C) Cs (strain barley, wheat, Spot Blotch, ? - hemibiotroph ND90Pr) cereals/ leaves Common root rot St (strain 28A) corn/leaves Northern Leaf ? Ht1, Ht2, Ht3, hemibiotroph Blight HtNe a Ch = C. heterostrophus, Cc = C. carbonum, Cv = C. victoriae, Cm = C. miyabeanus, Cs = C. sativus, St = S. turcica b Tcms = cytoplasmic male sterility, c hm1hm1 = homozygous recessive for carbonyl reductase, d Vb = presumed to be the same as the LOV1e (Pc-2) gene for resistance to P. coronata e Ht1, Ht2, Ht3, HtN are defined as resistance genes based on differential resistance/susceptibility to a set of S. turcica races pathogen (race O) of minor economic importance, first described in 1925 [7]. Subsequent research over the ensuing four decades since 1970 has demonstrated that the epidemic was triggered by the unfortunate confluence of complex DNA recombination events in both the fungal pathogen and the plant host. Race T is genetically distinct from race O in that it possesses an extra 1.2 Mb of DNA [8,9] encoding genes for biosynthesis of the polyketide secondary metabolite, T-toxin, an HST essential for high virulence [4]. These genes are missing in race O. On the plant side, the presence in Texas male sterile cytoplasm (Tcms) maize, of a hybrid mitochondrial gene called T-urf13, composed of segments of two mitochondrial and one chloroplast gene, is essential for susceptibility. Tcms corn does not need to be detasseled to prevent self-crossing because it is male sterile, a desirable trait for breeders producing hybrid 27 seed. The resulting popularity of Tcms maize was disastrous from the perspective of pathogen attack, however, as it served as a monoculture of susceptible germplasm [10,11]. Similarly, C. victoriae (Bipolaris victoriae), causal agent of Victoria Blight, produces the chlorinated cyclic pentapeptide HST, victorin, rendering it highly virulent to oats carrying the dominant Vb allele [12]. The fungus caused widespread destruction (20 states) in the 1940’s on oat varieties containing the recently introduced Pc-2 gene for resistance to crown rust caused by Puccinia coronata [13]. Like the C. heterostrophus T-toxin/Tcms case, the monoculture of Victoria oats carrying Pc-2 was the perfect milieu for attack by C. victoriae producing victorin, which elicits Pc-2-dependent Programmed Cell Death (PCD). Recent work with Arabidopsis identified a resistance-like protein responsible for susceptibility to C. victoriae and victorin [14,15]. This work is seminal in demonstrating fungal HSTs can target resistance proteins to promote disease. In contrast to the dominant plant host genes required for susceptibility to C. heterostrophus and C. victoriae, susceptibility to Northern Corn Leaf Spot caused by C. carbonum (Bipolaris zeicola) is conferred by a homozygous recessive maize gene(s) [16,17]. C. carbonum race 1 produces the cyclic-tetrapeptide HST, HC-toxin, which is specifically active against corn with the genotype hmhm, as is the fungus itself [4,18,19]. Hm1 and Hm2 encode carbonyl reductases which inactivate the toxin [16]; hmhm lines, cannot inactivate the toxin, and are therefore sensitive. The site of action of HC-toxin in susceptible corn is histone deacetylase; it is hypothesized that HC-toxin acts to promote infection of maize of genotype hm1hm1 by inhibiting this enzyme, resulting in accumulation of hyperacetylated core (nucleosomal) histones. This then alters expression of genes encoding regulatory proteins involved in plant defense [20,21]. C. carbonum races 2 and 3 do not produce the toxin. 28 C. miyabeanus (Bipolaris oryzae) is the causal agent of Brown Spot of rice which contributed, along with a cyclone and tidal waves, to the Bengal rice famine of 1942/1943 that resulted in starvation of more than two million people [22]. To date, no HST has been associated with virulence, although C. miyabeanus culture filtrates can suppress plant phenol production [23]. C. sativus (Bipolaris sorokiniana) causes diseases of roots (Common Root Rot), leaves (Spot Blotch), and spikes (known as black point or kernel blight) of cereals (mainly barley and wheat) [24,25] , but also attacks many grasses, including switch grass (Panicum virgatum L.) [3], [26,27] and Brachypodium distachyon (S. Zhong, unpublished). Three C. sativus pathotypes (0, 1 and 2) have been described [28], based on differential virulence patterns on three barley genotypes (ND5883, Bowman, and NDB112). Pathotype 0 isolates show low virulence on all three barley genotypes. Pathotype 1 isolates show high virulence on ND5883 but low virulence on other barley genotypes. Pathotype 2 isolates show high virulence on Bowman but low virulence on ND5883 and NDB112. Genetic analysis and molecular mapping indicates that a single locus, VHv1, controls high virulence of the pathotype 2 isolate ND90Pr on Bowman [29,30], however, the exact nature of the gene(s) was unknown before this study (see Results). More recently, a new pathotype, highly virulent on NDB112, the most durable spot blotch resistance source in barley [31], has been found in North Dakota [32] and Canada [33]. Setosphaeria turcica (Exserohlium turcicum, Helminthosporium turcicum), a hemibiotrophic vascular leaf pathogen, is a member of the closest genus to Cochliobolus (see Fig. 1 in Ohm et al., [1]), and causes Northern Leaf Blight (NLB), a major disease of maize and sorghum in the US and internationally [34]. To date, at least four races of S. turcica have been identified based on their differential virulence performance on maize carrying resistance genes 29 known as Ht [35,36]. In recent work, Martin et al. [34], identified new resistance genes (named St) in both maize and sorghum. The connections between the Ht and St resistance genes are unclear at this point. Until recently, it was assumed that necrotrophs use brute force methods (e.g., arsenals of cell wall degrading enzymes, HSTs) to invade and kill host tissues and do not subtly manipulate the host during infection, as do biotrophs with their arsenal of effectors [37]. Several lines of evidence, from studies of the Dothideomycete, necrotrophic wheat pathogens, Pyrenophora tritici-repentis [38] and Stagonospora nodorum, [39] indicate that, like biotrophs, these pathogens secrete protein effectors (in this case also called HSTs) that interact with host targets in a gene-for-gene manner. Unlike biotrophs, however, interaction of the fungal effector and host protein results in susceptibility, rather than resistance. The above-mentioned research on C. victoriae, further indicates mechanistic overlap of pathogenic lifestyle and has major implications regarding the challenge plants face in defending themselves against both necrotrophs and biotrophs [37,40]. Recent studies involving Arabidopsis have revealed that victorin-induced PCD requires a host NB-LRR-type resistance protein [15,41]. Thus a protein with canonical resistance protein structure is required for susceptibility. These observations point toward victorin, subverting effector triggered defenses against biotrophs, such as P. coronata (see above), to promote susceptibility to a necrotroph. Here we provide a comparative analysis of genome similarities and differences among Cochliobolus and Setosphaeria pathogens, with particular emphasis on strain and species-unique sequences, secondary metabolism genes, and genes encoding small secreted proteins. Identification of these key structural genomic and molecular differences is the first step in understanding species-specificity and how closely related necrotrophic and hemibiotrophic 30 pathogens cause disease. As proof of concept, we offer several examples of how comparative approaches pinpoint virulence associated, species-specific regions of interest. The Joint Genome Institute (JGI) has sequenced five strains of C. heterostrophus (three race T and two race O strains) and four additional species of Cochliobolus, including C. carbonum, C. victoriae, C. miyabeanus, and C. sativus and one strain of S. turcica. Add to this that host genome sequences (corn, rice, barley and B. distachyon) for six of these pathogens are available and one has the information base for dissecting both sides of the interaction mechanism going forward. 31 B. Results 1. Genome statistics Five strains of C. heterostrophus, one strain of C. sativus, and one strain each of C. victoriae, C. carbonum, C. miyabeanus, and S. turcica were sequenced by JGI (Tables II.2, II.3). Two C. heterostrophus strains and one strain each of C. sativus and S. turcica were fully sequenced as described in Materials and Methods, while the remaining genomes were sequenced using Illumina and assembled de novo using Velvet, as described in Materials and Methods. The highly inbred race O lab strain C5 was used as the reference sequence for all comparisons, as it is the most complete, consisting of only 68 scaffolds. Table II.2. C. heterostrophus race O strain C5, race T strain C4, C. sativus and S. turcica genome statistics Genome Characteristic Genome sequence total (Mb) Contig sequence total (Mb) C. heterostrophus strain C5 36.46 36.32 Sequenced strain C. heterostrophus C. sativus S. turcica strain strain C4 strain ND90Pr 28A 32.93 34.42 43.01 32.09 33.22 38.26 Genome scaffold count 68 207 157 407 Genome contig count 88 586 478 1951 Scaffold N/L50 7/1.84 Mb 13/0.96 Mb 7/1.79 Mb 8/2.14 Mb Contig N/L50 12/1.17 Mb 55/0.18 Mb 43/0.24 Mb 210/0.05 Mb % genome covered by repeats 9% 1% 6% 12.96% # predicted genes 13,336 12,720 12,250 11,702 Overall sequence assembly and annotation statistics are presented in Tables II.2 and II.3. All Cochliobolus genomes are in the 32-38 Mb range with an estimated gene content of 11,70013,200. The S. turcica genome is ~43 Mb with ~11,700 genes. Overall gene content and genome organization are highly similar within this group of fungi. In contrast, comparative analysis of C. heterotrophus, C. sativus and S. turcica in the context of 14 more distantly related Dothideomycetes genomes described elsewhere (Ohm et. al., [1]) revealed significant variation. 32 Table II.3. Statistics for short read re-sequenced Cochliobolus genomesa Strain Sequencing # Reads % reads % paired Read length # Assembled depth mapped (bp) nodes Velvet assembly size (bp) Ch Hm540 35.2 37,796,798 95 94 35 4,387 32,575,369 Ch Hm338 79.55 37,923,292 97 98 76 5,227 33,353,785 Ch PR1x412 33.58 36,292,318 94 95 35 7,229 32,661,792 Cv FI3 30.41 81,216,986 19 92 76 4,525 33,638,943 Cc 26-R-13 33.05 83,423,746 20 90 76 3,671 32,298,417 Cm WK1C ATCC 44560 27.39 81,110,386 17 90 76 a Using Illumina technology Ch =C. heterostrophus, Cv = c. victoriae, Cc = C. carbonum, Cm = C. miyabeanus 3,553 32,695,608 2. Mapping scaffolds to the genetic maps C. heterostrophus: A genetic map with 125 RFLP markers was constructed previously [8] using C. heterostrophus race O field strain Hm540 (sequenced herein) and race T C-strain B30.A3.R.45 (same backcross series as strains C5 and C4, sequenced herein [42,43]) as parents. RFLP sequences were used to refine the C. heterostrophus race O strain C5 physical assembly and link it to the genetic RFLP map (Fig. II.1). The interconnected genetic and physical maps allowed comparisons of physical and genetic distance, which was found to be ~13 kb/cM on average (Fig. S1). Correlations were also made between previously estimated chromosome sizes based on CHEF gel analysis [9], and physical size based on sequence assemblies (Tables S1, S2). 33 34 Figure II.1. C. heterostrophus RFLP sequences anchor sequenced scaffolds to the genetic map. Genetic linkage groups determined by Tzeng et al. [8], are in light blue (linkage group numbers on the left). Assembled scaffolds from the reference C5 strain that could be anchored to each linkage group are in light green; numbers above are internal JGI identifiers (Table S2). Black bars indicate the relative locations of RFLPs Tzeng et al. [8] or feature, linked by dotted lines. The relative scale of genetic/physical distance was determined by calculating the average genetic/physical distance between consecutively placed RFLP markers (Fig. S1). Scaffold ends marked with a T contain telomeric sequence. Unplaced scaffolds are shown in pink. Maps are to scale and shown as centiMorgans (cM) or kilobases (kb). 35 Based on RFLP map data, parental field isolate Hm540 was reported to lack the dispensable chromosome present in the other parental strain (B30.A3.R.45) used to build the map [8]. Comparisons of all five sequenced C. heterostrophus strains, using the Mauve alignment tool [44], supported this observation and revealed that all sequenced strains carried the previously recognized ‘B’ or dispensable chromosome (which corresponds to strain C5 scaffold 16) (Fig. S2). Race O chromosomes 6 and 12 are of interest because counterparts are reciprocally translocated in race T and the high-virulence conferring Tox1 locus, encoding genes for biosynthesis of T-toxin, maps genetically to both breakpoints [9]. Comparison of the race O and race T assemblies provided some clues as to the physical locations of these breakpoints, but the exact positions remain elusive, due to structural complexity associated with these regions [45]. Additional details regarding linkage of the C. heterostrophus physical assembly to the genetic map are available in Supplemental Methods (Text S1). C. sativus: Before mapping sequenced scaffolds to the previously constructed C. sativus genetic map [30] 121 polymorphic simple sequence repeat (SSR) markers were identified in the assembly sequences of the ND90Pr and ND93-1 parents, as described in Materials and Methods. Then, sequences of the SSRs and other markers were used to assign sequenced scaffolds to the updated map. Thirty of these linkage groups contained SSR markers and were found to be associated with 16 scaffolds, summing to 29.32 Mb. Seven linkage groups were unassigned (Fig. S3). The two AFLP marker sequences (E-AG/M-CG-121 and E-AG/M-CA-207), cosegregating with the VHv1-associated high virulence of C. sativus pathotype 2 on cultivar Bowman [30], were used as blast queries against the ND90Pr genome assembly. E-AG/M-CG-121 mapped to scaffold 5, while E-AG/M-CA-207 mapped to scaffold 40 (Fig. S3). Details of the construction, linking, and analysis of the C. sativus map are available in Supplemental Methods (Text S1). 36 Table II.4. SNPs between C. heterostrophus (Ch) strain C5 and other strains and species Isolate ChC4 ChHm338 ChHm540 ChPR1x412 C. victoriae C. carbonum C. miyabeanus C. sativus # Scaffolds 207 2,711 4,387 7,229 1,207 3,671 3,553 157 # Aligned 173 1,988 1,954 4,269 522 1096 891 70 Total bases Aligned bases 32,929,167 31,789,127 33,353,785 32,483,575 32,575,369 30,971,859 32,661,792 31,041,698 33,659,279 24,967,357 32,298,417 24,784,636 32,878,767 24,421,161 34,417,436 25,730,439 Total SNPs 1,584 30,624 50,864 33,552 2,083,899 2,059,993 2,110,786 1,981,616 Aligned bp/SNP 20,068.9 1,060.7 608.9 925.2 12 12 11.6 13 S. turcica 407 179 43,014,577 6,932,055 749,006 9.3 3. Single nucleotide polymorphisms (SNPs) between the reference C. heterostrophus C5 strain and other strains i. SNPs in C. heterostrophus strains Genome assemblies were aligned, pairwise, to the C. heterostrophus race O C5 reference using MUMmer [46] to analyze SNP frequencies between strains and species and to infer wholegenome similarity. C. heterostrophus race T strain C4, which belongs to the same inbred laboratory strain series as the C5 reference [42] contained 1,584 SNPs (Tables II.4, S3). In contrast, the three C. heterostrophus field strains, race O strain Hm540, race T strains Hm338, and PR1x412 had 50,864, 30,624, and 33,552 SNPs, respectively, (~20-30X more than strain C4) (Tables II.4, S3). More SNPs were identified for Hm540, the only race O field isolate, than for the race T field strains Hm338 and PR1x412, despite the reference C5 strain being race O. This supports previous RFLP data which indicated that race O field strain Hm540 was more diverse than any field isolate examined [8] and the hypothesis that race T arose once and more recently than race O [9,47,48]. For all C. heterostrophus strains, 30-31.5 Mb (95-97%) of the assembled basepairs (bp) could be aligned, although only 40-80% of the scaffolds could be 37 aligned. Thus 3-5% of each genome could not be aligned and corresponding sequences were located on small, difficult to assemble contigs. ii. SNPs in C. sativus strains For the two C. sativus genomes, ND90Pr and ND93-1, 86.6% of the higher quality ND90Pr genome could be aligned to 96.6% of the ND93-1 genome, yielding 60,448 SNPs (Table II.5). The relative similarity between these two strains is comparable to that seen between C. heterostrophus strains. TABLE II.5. SNPs between C. sativus strains ND90Pr and ND93-1 Isolate # Scaffolds # Aligned Total bases Aligned bases ND90Pr ND93-1 157 18,112 156 17,418 34,417,436 30,342,250 29,889,643 29,309,584 Aligned bp/SNP 494 485 iii. SNPs in Cochliobolus species C. victoriae, C. carbonum, C. miyabeanus, and C. sativus species had 2,083,899, 2,059,993, 2,110,786, and 1,981,616 SNPs, respectively, when aligned to C. heterostrophus reference strain C5, ~54X fold more than comparisons within species (Tables 4, S3). In each case, 24-25 Mb (75%) of assembled bps could be aligned to the reference, indicating that, in addition to the much higher quantity of SNPs than between C. heterostrophus strains, a full quarter of each genome was not present in C. heterostrophus. Although C. sativus has a wider host range than any of the other species, there is no compelling evidence that it is less related to C. heterostrophus than the other species. All species are estimated to have arisen relatively recently (< 20MYA ago, see Fig. 1 in Ohm et al., [1]). Analysis of the mating type (MAT) regions (Text S1, Fig. S4, Table S4) yielded similar patterns, except that within the C. heterostrophus species, the most similar MAT flanking regions were those of the same mating type, regardless of whether the strain was race O or race T, or an inbred or field strain. C. 38 carbonum and C. victoriae are capable of crossing with each other [48] and thus would be predicted to be closely related and to have fewer SNPs between them than between either of them and other species. Indeed, when C. victoriae was aligned to C. carbonum, 292,216 SNPs were identified, roughly 10-fold fewer than when Cochliobolus species are compared to the C. heterostrophus reference, yet 10-fold more than are present when C. heterostrophus field strains are compared to the C. heterostrophus C5 reference strain (Tables II.6, S3). Thus, the SNP data are in line with the notion that C. victoriae and C. carbonum are closely related and that C. victoriae may have arisen from a C. carbonum isolate [49]. For comparison, aligning C. miyabeanus or C. sativus to the C. carbonum genome yields 2,078,277 and 2,022,068 SNPs, respectively (Tables II.6, S3), in the range seen when comparisons are made with C. miyabeanus, C. carbonum, C. victoriae, or C. sativus to C. heterostrophus C5 (Tables II.4, S3). Table II.6. SNPs between C. carbonum and other species Isolate # Scaffolds # Aligned Total bases Aligned bases C. victoriae 1,207 565 33,659,279 30,153,232 C. miyabeanus 3,553 1,008 32,878,767 25,518,991 C. sativus 157 96 34,417,436 26,024,810 ChC5 68 67 36,456,735 27,650,456 S. turcica 407 329 43,014,577 7,595,377 Total SNPs 292,216 2,078,277 2,022,068 2,094,062 747,875 Aligned bp/SNP 115.2 12.3 12.9 13.2 10.2 SNP calls generated in all genome comparisons were inflated for A to G, T to C, C to T, and G to A transitions, the most common type of mutation. In all comparisons made, including C5 to C4, these transitions were 5-10 times more abundant than other changes (Table S3). iv. SNPs between C. heterostrophus and S. turcica We could align only 16.12% (6.9 Mb) of the S. turcica genome to the C. heterostrophus reference genome, as indicated in Table II.4. Full details are available in Table S3. 39 v. Summary Inbred C. heterostrophus C4 strain has far fewer SNPs (1,584), than C. heterostrophus field strains, when compared to the C. heterostrophus C5 reference. The numbers of SNPs in the C. heterostrophus field strains are comparable to each other, with Hm540, a race O isolate, containing the most (50,864) when compared to race O C. heterostrophus C5. The largest numbers of SNPs are between C. heterostrophus C5 and the additional Cochliobolus species (1.9-2.1 million) and these are ~54 fold higher than within species SNPs (Tables II.4, S3). The C. sativus strains are as similar to each other as different C. heterostrophus strains are to one another, while C. carbonum and C. victoriae demonstrate a level of relatedness in between that seen among isolates within a species and across species. 4. Identification of strain- and species-specific genomic regions To begin to identify regions in the C. heterostrophus C5 assembly not represented in other strains, we first mapped gaps in the reference assembly (thick vertical black bars, Fig. II.2). A single gap was present in the assembly of 9 of 16 chromosomes and we speculate that these gaps correspond to centromeric regions. We then mapped sequence reads of all Cochliobolus genomes in this study to the C. heterostrophus C5 reference, identifying regions in the C5 reference that were not present in the query genome (Table S5). The sets of C5 genomic regions that were absent in a given query were combined to determine genomic regions unique and/or conserved at different taxonomic levels. 40 41 Figure II.2. C. heterostrophus unique regions, secondary metabolite and small secreted protein encoding genes are distributed throughout the genome. Reads from each C. heterostrophus or Cochliobolus species isolate were aligned to the C. heterostrophus C5 reference genome and regions of low coverage mapped; orange lines indicate no match in all non C. heterostrophus species but conserved in C. heterostrophus isolates (C. heterostrophus unique), green lines indicate low coverage of C. heterostrophus strain C4 reads, blue lines indicate low coverage of all C. heterostrophus field isolates. Gaps in the assembly are indicated as black vertical bars. The locations of all C. heterostrophus strain C5 NPS (red flags), PKS (yellow flags), and SSP (green flags) genes were also mapped to the C5 reference assembly. Note that one or more NPS or PKS genes map to most linkage groups and that several map to unplaced scaffolds. SSPs map to every chromosome/linkage group and most unplaced scaffolds. Same scale as Fig. II.1. 42 Individual reference-unique region counts were recorded for each of the C. heterostrophus strains. There were 609 areas of the C5 assembly unique (C4 reads did not map there) to the C5 genome when C4 reads were mapped to it. For C. heterostrophus strains Hm540, PR1x412, and Hm338, there were 3,279, 4,383 and 1,480 such regions, respectively. When only regions greater than 5,000 bp were considered, there were 19 when C4 was used as the query, and 33, 75, and 30, when PR1x412, Hm540, and Hm338 were used as the queries, respectively. Many of the gaps associated with Hm540 mapped to reference C5 scaffold 16 corresponding to the dispensable B chromosome, which is absent in Hm540 (Fig. S2). The C5 reference-unique regions were then combined and filtered to identify conserved genomic regions at the strain or species level, where regions were unique in one type of comparison, but not in others (Table S5). We designated “inbred C strain”-specific regions as gaps found when all field strains were aligned to C5 but not when C4 was aligned to C5, race O specific regions as gaps found when all race T strains were aligned to C5 but not when race O strain Hm540 was aligned to race O strain C5, and C. heterostrophus-specific regions as gaps found when all Cochliobolus species were aligned to C5, but not when any C. heterostrophus strain was aligned. A total of 28,556 bp is missing from all C. heterostrophus field strains, yet present in the C4 assembly (Table S5). This ~30 kb of DNA represents sequence uniquely conserved in the inbred C strains. There are at least six fungal-specific Zn2Cys6 transcription factors (ID# 1019013, 1020538, 1021058, 1021066, 1100349, 1100899) present in this C strain unique cache that may signify “early action” strain diversification. Zn2Cys6 transcription factors were among the most abundant predicted domains in the C. heterostrophus gene catalogue [1]. 43 There were almost no race O specific regions identified (sequence found only in C5 and Hm540); only 11 regions, summing to 4,309 bp, were identified (Table S5). Our working hypothesis is that the essential difference between race O and race T is the 1.2 Mb of Tox1 race T DNA (not in race O C5 and therefore not able to be aligned). Both race O only regions (scaffold 12, 732532-734880, and scaffold 19, 441888-443521) contain a single protein each (ID# 59063, 34937) with no conserved domains or predicted function. Most significantly, at the species level, a total of 11.76 Mb DNA was missing from all non-C. heterostrophus Cochliobolus genomes analyzed, yet found in all field strains of C. heterostrophus. Only 1.6 Mb of this was found in pieces larger than 5 kb. Most of the sequence that separates C. heterostrophus from other species, therefore, is not the result of large wholesale insertions or deletions of DNA, but from a more piecewise gain and loss. 5. Secondary metabolism: nonribosomal peptide synthetases Nonribosomal peptide synthetases (NRPSs), found in fungi and bacteria, are multimodular megasynthases that catalyze biosynthesis of small bioactive peptides (NRPs), including virulence determinants, such as HSTs, via a thiotemplate mechanism independent of ribosomes [50,51,52,53,54]. NRPSs synthesize peptides using sets of core domains, known as modules, which consists of three domains: 1) an adenylation (AMP) domain which recognizes and activates the substrate via adenylation with ATP, 2) a thiolation (T) or peptidyl carrier protein (PCP) domain which binds the activated substrate to a 4’- phosphopantetheine (PP) cofactor via a thioester bond and transfers the substrate to 3) a condensation (C) domain which catalyzes peptide bond formation between adjacent substrates on the megasynthase complex. 44 NRPSs can be mono-, bi-, or multi-modular and core domains in any particular multimodular enzyme may be most closely related to one another or to a domain from a different NRPS. The suites of NRPS encoding genes (NPS) in the C. heterostrophus C4 and C5 genomes were identified and annotated previously [55,56,57]. To address degree of conservation and evolutionary relationships of NRPS proteins in our subject species in order to make inferences about function, we used the fungal AMP-binding (AMP) domain Hidden Markov Model (HMM) developed by Bushley and Turgeon [57] to identify individual AMP domains in the additional strains of C. heterostrophus and other Cochliobolus species, plus S. turcica. Phylogenetic trees were built to develop a comparative NRPS AMP domain inventory and included the known C. heterostrophus C5 AMP domains as a reference (Table S6). i. Conservation of known C. heterostrophus NRPSs First, using the highly curated set of C. heterostrophus NRPSs (Table II.7, left column), we determined if all 28 AMP domains of all 14 C. heterostrophus C5 reference NRPS proteins were present in each genome analyzed. For the C. heterostrophus genomes, all C5 reference AMP domains and thus all complete NRPS proteins were present (Table II.7, Fig. II.3, for full phylogenetic trees see Figs. S5A, B, and for master inventories, see Table S6), with the single exception of bimodular NPS5, which was absent from race T strain PR1x412. Thus, there is almost complete conservation of NRPSs at the species level. At the genus level, seven of the 14 NRPS proteins (and thus genes) in C. heterostrophus reference strain C5 were conserved across all species (Table II.7, Fig. II.3, for full phylogenetic trees see Figs. S5A, B). When conservation of NRPSs was considered across all Cochliobolus spp. and the related maize pathogen S. turcica, six of the 14 NRPS proteins in C. heterostrophus reference strain C5 were completely conserved across all species (NPS2, 4, 6, 19, and 12, and 45 12-like, Table II.7). The phylogenetic profile (Fig. II.4) of the highly conserved NPS2 protein is an example of complete conservation. Note that all four NPS2 AMP domains are found in all species. NPS2 biosynthesizes the hexa-peptide siderophore, ferricrocin, and an evolutionary mechanism by which a four AMP domain NRPS can generate a six component metabolite has been proposed by us [58,59]. In cross genome evaluation of conservation of NRPSs (see Table S18 in [1]), this was one of only two NRPSs conserved across 18 Dothideomycete genomes. Table II.7. Conservation of C. heterostrophus strain C5 nonribosomal peptide synthetases in other Cochliobolus strains and species C5 C4 Genbank JGI ID In other C. In other Cochliobolus In S. turcica? name accession heterostrophus spp.? strains? NPS1.1 AAX09983 1101207 + Cv, Cc - NPS1.2 AAX09983 1101207 + Cv, Cc - NPS1.3 AAX09983 1101207 + Cv, Cc - NPS2.1 AAX09984 128084 + ++ NPS2.2 AAX09984 128084 + ++ NPS2.3 AAX09984 128084 + ++ NPS2.4 AAX09984 128084 + ++ NPS3.1 AAX09985 128098 + Cm, Cs - NPS3.2 AAX09985 128098 + Cm, Cs + NPS3.3 AAX09985 128098 + Cm, Cs + NPS3.4 AAX09985 128098 + Cm, Cs + NPS4.1 AAX09986 1091637 + ++ NPS4.2 AAX09986 1091637 + ++ NPS4.3 AAX09986 1091637 + ++ NPS4.4 AAX09986 1091637 + ++ NPS5.1 AAX09987 1095362 not in PR1x412 - - NPS5.2 AAX09987 1095362 not in PR1x412 - - NPS6.1 AAX09988 128080 + ++ NPS7.1 AAX09989 1209664 + +- NPS8.1 AAX09990 1227314 + -- NPS8.2 AAX09990 1227314 + Cs - NPS9.1 AAX09991 1159473 + -- NPS9.2 AAX09991 1159473 + -- NPS10.1 AAX09992 1175121 + ++ NPS11.1 AAX09993 1104551 + Cs - NPS12.1 AAX09994 116719 + ++ (116719) NPS12.1 - 1223123 + ++ (118012) NPS13 AY884198 1215193 + -- + = present, - = absent; 2x = two copies Strains Ch = C. heterostrophus, Hm540, PR1x412 are C. heterostrophus strains, Cs = C. sativus, Cv = C. victoriae, Cc = C. carbonum, Cm = C. miyabeanus, St = S. turcica 46 Figure II.3. Cartoon of cross-species phylogenomic analyses of individual AMP binding domains from NRPS proteins. NRPS AMP domains were extracted from all five C. heterostrophus and from the C. victoriae, C. carbonum, C. miyabeanus, C. sativus, and S. turcica genomes. Members of the reference set of previously annotated C. heterostrophus NRPS AMP domains [57] were used as benchmarks for branches. Branches of the full Augustus phylogenetic tree (Fig. S5A) are 47 collapsed according to clustering with the reference set of C. heterostrophus AMP domains. Presence in each of the five C. heterostrophus strains (ChC5, ChC4, Ch540, Ch338, ChPR1), Cochliobolus species [C. victoriae (Cv), C. carbonum (Cc), C. miyabeanus (Cm), C. sativus (Cs)], or S. turcica (St) is noted in parentheses. Designations such as ‘St x2’ indicate that two AMP domains from S. turcica are present. AMP domains not grouping with the previously annotated C. heterostrophus set are labeled as ‘New_1 through _8’. Groups are color-coded according to their distribution in the genomes examined: green = present in all genomes, blue = absent from S. turcica only, orange = exclusive to C. heterostrophus and present in all C. heterostrophus strains, purple = present discontinuously in some, but not all Cochliobolus genomes, red = absent from the C. heterostrophus C5 reference, and black = absent from all included genomes. The latter includes outgroups and NRPS AMPs in species other than Cochliobolus or Setosphaeria producing some well-known metabolites [e.g., Alternaria alternata AMT producing AM-toxin, (AMT)]. Bootstrap values 75% and above are indicated on the branches. Branch lengths represent the average number of substitutions per site. Asterisks indicate unique C. sativus NRPSs discussed in text (Table II.8, Fig. II.7). 48 Figure II.4. NPS2 is an example of a highly conserved Dothideomycete NRPS. NPS2 consists of four AMP domains (cartoon above partial tree) that produce the hexapeptide intracellular siderophore, ferricrocin, responsible for iron storage within cells. NPS2 is present in all five strains of C. heterostrophus, all other Cochliobolus species and Setosphaeria. Note four branches, which cluster together on the thumbnail of the full tree (Fig. S5A), each corresponding to one of the four NPS2 AMP domains (gray AMP in NRPS cartoon to the right). Each fungal race or strain is color coded as indicated. Reference C5 AMP indicated as e.g., ChC5 77609 NPS2 AMP1 4; 77607 is the Genbank protein ID, NPS2 is the NRPS designation, AMP1 4 denotes the 1st AMP domain of a total of 4 AMPs in the NRPS. JGI protein IDs are given for S. turcica, C. sativus, and C. heterostrophus strain C5. All other AMPs indicated by NODE number and Augustus gene call number (e.g., Cvict NODE 1785 g7786 t1 1 is a C. victoriae AMP on NODE 1785 in the Velvet assembly carrying Augustus called gene 7786; t1 1 indicates the order in which the AMP domain was predicted by HMMER). C. heterostrophus strain C4 AMP domains are indicated by scaffold and gene call. Bootstrap values 80% and above are indicated on the branches. 49 ii. Discontinuously distributed and expanded NRPSs Of those NRPS proteins for which all C. heterostrophus AMP domains are not conserved across all species (Table II.7), three, NPS1, NPS3, and NPS13 are of particular note as they are expanded discontinuously, with multiple homologs for some, but not all AMP domains of these proteins in different species (Figs. II.5, II.6). We hypothesize that this group of NRPSs is a spawning ground for AMP domain diversity. On the whole protein level, the complete C. heterostrophus NPS1 (trimodular) and NPS3 (tetramodular) domain sets are either present or absent in other species. NPS1 is intact in C. victoriae and C. carbonum, while NPS3 is intact in C. miyabeanus and C. sativus, but absent from the other genomes (Fig. II.5, Table II.7). Monomodular C. heterostrophus NPS13 is found only in C. heterostrophus (Fig. II.5, Table II.7). NPS1, NPS3 and NPS13 protein AMP domains, as noted above, are expanded discontinuously resulting in a suite of novel proteins we call ‘NPS1/NPS3/NPS13 expanded’ (Fig. II.5). These new proteins may be mono- or multi-modular. C. victoriae has three such proteins, C. carbonum and C. miyabeanus have one each, and C. sativus has seven (Table 8, Fig. II.5). A C. heterostrophus NPS13-related module is present in tetramodular form in C. carbonum, C. victoriae, and C. miyabeanus, and in trimodular form in C. sativus. Both the monomodular NPS13, and tetramodular NPS13 related protein, are found in the C. heterostrophus Hm540 strain. 50 51 Figure II.5. The NPS1/NPS3/NPS13 expansion group of NRPS AMP domains. NPS1 and NPS3 AMP domains are discontinuously distributed and expanded across the Cochliobolus isolates sequenced (Fig. II.6 and [57]). NPS1 and NPS3 AMP 2 and 4 domains (color coded green) group at the top of the phylogenetic tree (thumbnail to the left, full tree Fig. S5A), while NPS1, NPS3 and NPS13 AMP domains 1 and 3 (color coded red or blue) group near the bottom of the tree. Each C. heterostrophus strain has NPS1 NPS3 and NPS13. The other Cochliobolus species possess either a complete C. heterostrophus NPS1 or NPS3 ortholog, but not both and none has NPS13. All species, however, have one or more additional NPS proteins consisting of NPS1/NPS3 or NPS1/NPS3/NPS13-related domains, that are absent from all C. heterostrophus genomes except Hm540. The C. heterostrophus Hm540 genome includes all four corresponding genes: NPS1, NPS3, NPS13, and the additional NPS1/NPS3/NPS13 gene. The pattern of AMP domain expansion/loss/recombination is complicated and the additional NRPSs can be mono-, bi-, tri-, and tetra-modular proteins. C. sativus has 7 expanded NRPSs whose AMP domains group with NPS1/NPS3/NPS13 proteins, (Fig. II.6). One of these is C. sativus ID 115356 (double asterisk) which maps to a unique region of the genome in pathotype 2 strain ND90Pr (Fig. II.7), associated with high virulence on barley cultivar Bowman. Gene/AMP nomenclature as described in Fig. II.4. 52 53 Figure II.6. NPS1, NPS3 and NPS13 are examples of NRPS proteins encoded by highly recombinogenic and expanded NPS genes. The reference NPS1, NPS3 and NPS13 proteins are cartooned bottom left and AMP domains are color coded as in Fig. II.5. AMP domains corresponding to these proteins are completely conserved in the five strains of C. heterostrophus, but show discontinuous presence in all other Cochliobolus species (Fig. II.5) and Setosphaeria. Thumbnail of full AMP tree (Fig. S5A) is shown at left. Note some AMP domains from NPS1 and NPS3 group at the top of the tree (AMPs 2 and 4, green), while the rest group at the bottom of the tree (AMPs 1 and 3, red); NPS13 AMP1 (blue) also groups at the bottom of the tree. Branches correspond to individual AMP domains which group together and the particular corresponding AMP domain is colored coded on the right of the diagram. Branches not in the original reference set of C. heterostrophus AMPs [57] are labeled as ‘expanded’ (Fig. II.5). Gene/AMP nomenclature and bootstrap values as described in Fig. II.4. Double red asterisk indicates C. sativus protein ID 115356 discussed in Fig. II.7). 54 As shown in Fig. II.6, C. heterostrophus NPS1 AMP2 groups with C. heterostrophus NPS3 AMP2 and AMP4 orthologs, with 99% bootstrap support. The NPS1 and NPS3 AMP2 and AMP4 expanded group (New_1, 2, 3, Fig. II.3) is most closely related to NPS3 AMP2 (100% bootstrap support). C. heterostrophus NPS1 AMP1 and AMP3 group with C. heterostrophus NPS3 AMP 1 and AMP3 orthologs (NPS1 AMP1, NPS3 AMP 1 and AMP3 group together with 93% bootstrap support). The NPS1 and NPS3 AMP1 and AMP3 expanded group (New_7, Fig. II.3) is nested within, but without bootstrap support. S. turcica has seven total NPS1/3/13 associated AMP domains. Four cluster with NPS3 AMP3, two with NPS3 AMP2, and one with NPS3 AMP4 (Fig. II.6, Table S6). These expanded S. turcica domains belong to four different NRPSs (ID# 51661, 36641, 65284, 48467), none of which corresponds to Cochliobolus NRPSs. Monomodular NPS13, as noted, is found only in C. heterostrophus, but there is an NPS13 expanded group of AMP domains, sister to (95% bootstrap support) C. heterostrophus NPS13. C. sativus possesses additional NPS13 expanded domains, always co-occurring with NPS1 and NPS3 expanded domains (Figs. II.5, II.6). iii. Species-unique NRPSs In initiating our survey, we hypothesized that this category of NRPS would be most likely to include candidates associated with virulence functions, given the unique or spotty distribution signatures of HSTs. We define an NRPS as unique when no other Cochliobolus species has all of the orthologous AMP domains in identical whole-protein organization. The reference, C. heterostrophus, has 14 NRPSs, three of which (NPS5, NPS8 and NPS9) are unique to this species but found in both race O and T (Tables II.7, II.8, S6). None of these three has an obvious role in virulence [55]. Of the other Cochliobolus species, C. miyabeanus has the fewest 55 total (11) NRPSs and no unique ones, while C. sativus has the most (25), 14 of which are unique (Table II.8) and include seven belonging to the NPS1/NPS3/NPS13 expanded group (and an eighth unique NRPS, ID# 358216) (Table II.8, Fig. II.5). When the AMP domains from the 25 NRPSs identified in C. sativus isolate ND90Pr (pathotype 2) were used as blast queries to identify orthologs in C. sativus isolate ND93-1 (pathotype 0), five (ID# 130053, 140513, 104448, 115356, and 350779) were not present in the latter and thus are unique to ND90Pr. C. carbonum and C. victoriae have 20 and 18 total NRPSs, respectively. Six are unique to C. carbonum (Table II.8). None of these is in the NPS1/NPS3/NPS13 expanded group. One of the C. carbonum unique NRPSs is HTS1, responsible for HC-toxin biosynthesis. It has long been recognized that HTS1 is only found in race 1 of this species and not in any other Cochliobolus species [60] and our genome survey confirms this. Our NRPS survey, however, identified an ortholog in S. turcica (see section below). Four other novel C. carbonum NRPS AMP domains group in the 11 AMP domain Tolypocladium inflatum SimA clade for biosynthesis of cyclosporin, suggesting C. carbonum as a possible source of a cyclosporin-type molecule (Table II.8, Fig. S5). C. victoriae has five unique NRPSs, including two from the NPS1/NPS3/NPS13 expanded group (Fig. II.5, Table II.8). One of the C. victoriae unique NRPSs is on n1179 (g7087) and is an ortholog of the gene for gliotoxin. Another C. victoriae unique AMP domain (on node 572, Fig. S5A) is incomplete but has a match to a bimodular S. turcica NRPS (ID #97841). The fifth unique AMP, on node 3108 (Fig. S5A), groups with the NRPS for aminoadipate reductase (lysine biosynthesis), but without bootstrap support. For S. turcica, eight (ID# 36641, 48467, 51661, 65284, 99043, 155102, and 54477, 97841, 99181 Table S6) NRPSs were unique, not occurring in any Cochliobolus genome. One of these, 99181, clusters with varying support with PesA, a Metarhizium anisopliae NRPS 56 producing an unknown product. Four of S. turcica’s unique NRPSs (ID# 36641, 48467, 51661, 65284) contained NPS1/NPS3/NPS13 expansion AMP domains as noted above (Table S6). Table II.8. Total and unique Cochliobolus NRPS and PKSs Species NRPS PKS Total Unique ID NPS1/NPS3/ ID Total Unique ID NPS13 Ch race 14 0 NAa, b associated 3 AAX09983, 25 2 ABB08104, T AAX09985, ABB76806 AY884198 Ch race 14 0 NA 3(4) 1101207, 23 0 NA O 128098, 1215193, (n1164, g9443)c Cv 18 5 n3108, 4 n99, g1225; 21 1 n4, g34 g9998; n244, n244, g2706; n37, g2706; n37, g303; n1179, g303; g7087; n572, n1179, g5163 g7076 Cc 20 6 n105 g794; 2 n841, 27 2 n2423, n2409 g6648; g3976; n82, g6650; n3710, g629 n189, g1086 g7564; n559, g3150; n120, g870; n464, g2585 Cm 11 0 NA 2 n281, 21 0 NA g1518; n8391, g8904 Cs 25 14 54465, 8 54465, 18 0 NA 121744, 121744, 129982, 129982, 358670, 358670, 129976, 129976, 351207, 351207, 115356, 115356, 130053, 358216, 140513, 104448, 49884, 103953, 350779, 25865 Strains Ch = C. heterostrophus, Cv = C. victoriae, Cc = C. carbonum, Cm = C. miyabeanus, Cs = C. sativus; a NA, not applicable; b for Ch C5 and Cs, JGI protein IDs are given; for Ch C4, GenBank protein IDs are given; all others are Augustus gene call numbers; c in Hm540 only, See Figs. S5A and S6A, Tables S6 and S10. 57 iv. NRPSs showcasing the value of the phylogenomic approach in pinpointing candidate virulence determinants Case Study 1: NRPSs unique to C. sativus pathotype 2, isolate ND90Pr determine virulence on barley cultivar Bowman. Previous genetic studies have indicated that a single locus (VHv1) in C. sativus pathotype 2 isolate ND90Pr controls high virulence on barley cv. Bowman [30]. Two AFLP markers E-AG/M-CG-121 and E-AG/M-CA-207 that co-segregate with the VHv1 locus in ND90Pr [30] mapped to scaffold 5 (Fig. II.7A) and 40 (Fig. S3), respectively, in the genome assembly. The VHv1 region (distal end of scaffold 5) carrying the E-AG/M-CG-121 marker includes 43 predicted genes (Table S7), plus many repetitive elements (Fig. II.7A, Table S8). None of the 43 genes was found in the C. sativus pathotype 0 isolate ND93-1 genome. Two of the genes in the region encode NRPS (ID# 115356, 140513), mentioned above as unique to isolate ND09Pr. 115356 is on a branch by itself in the NPS1/NPS3/NPS13 expansion group on the NRPS AMP tree (Figs. II.6, II.5, double asterisks, Table II.8) and is not found in any of the other Cochliobolus species, in S. turcica, or in Genbank. NRPS ID# 140513, also unique to the VHv1 region (Fig. II.7), maps to a C. sativus-specific clade we call ‘New_8’, consisting of seven AMP domains (Fig. II.3, asterisk) corresponding to three additional C. sativus- unique NRPSs. One of these is ID# 130053 with three AMPs on scaffold 25. Based on proximity in the genome assembly, we believe a second unique AMP, in protein ID# 49884, also on scaffold 25, is actually a fourth AMP domain of 130053. The remaining NRPS grouping in the New_8 clade is ID# 104448. 58 Figure II.7. Genomic organization of the scaffold associated with the VHv1 locus conferring high virulence of pathotype 2 isolate ND90Pr to barley cv. Bowman compared to the corresponding region in pathotype 0, isolate ND93-1. A. Mauve alignment [44] of ND93-1 scaffolds to ND90Pr. Colored blocks [Locally Collinear Blocks (LCB)] indicate matches between the two genomes, and vertical block shading corresponds with % similarity. Note very few colored blocks at right end of top row. The ~ 133 kb VHv1 locus, which maps to distal end of scaffold 5 (2.18 Mb), is unique to isolate ND90Pr, as indicated by absence of colored blocks at right end of the top row and the second row which is the same region at higher resolution. Below this Mauve alignment segment of the VHv1 region on scaffold 5 (from position 2,044,422 to 2,177,878) is the JGI browser view of the same region, displaying gene models and repeats. There are forty-three predicted genes (blue) in this region, only a fraction of which have KOG or GO descriptions (Table S7). Two NRPSs (ID # 115356 and 140513, shown in red) map to this region and are unique to the ND90Pr isolate and also not found in any of the genomes examined in this manuscript (Figs. II.3, II.5 Table II.8). E-AG/MCG-121 is one of two AFLP markers (the other is E-AG/M-CA-207 on scaffold 40) are linked to the virulence locus, VHv1 (Fig. S3). B. Quantitative real-time PCR analysis of five NPS genes (protein ID 49884, 350779, 130053, 115356, 140513) during infection of barley cv. Bowman. Gene expression was normalized based on the expression of the β-Actin gene, and the values are the relative expression levels in comparison with M96, a mixture of mycelia harvested, at 96 hours after 59 culture set up, from different media including PDA, MM, V8PDA, and water agar. B6, B12, B24, B48, B72, and B96 are samples collected at 6, 12, 24, 48, 72, and 96 hours after inoculation. Primers are shown in Table S9. The error bars indicate the minimum and maximum values of relative expression of the gene. C. Inoculation of barley cv. Bowman with wild type (ND90Pr) and a mutant lacking the gene corresponding to protein ID 115356. Images taken 7 days after inoculation. Virulence on Bowman is significantly reduced compared to that of plants inoculated with the wild-type strain (Fig. S7). Differences in NPS gene content and pathogenicity phenotype of closely related C. sativus strains allowed us to identify candidates for functional analyses. We hypothesized that, since the VHv1 region is unique to isolate ND90Pr and contains two NRPSs (ID#140513, 115356) unique to C. sativus, one or both of these might be responsible for high virulence on barley. We conducted real time PCR (Table S9) on infected barley leaves and demonstrated that expression levels of the genes corresponding to 140513 and 115356 in the VHv1 region, and also of the genes corresponding to unique proteins 130053 and 49884 described above were up- regulated 12 hours post inoculation (Fig. II.7B), while the gene corresponding to protein ID# 350779, which maps in the Gliotoxin clade (Fig. S5A, Table S9), was not. Deletion of the gene corresponding to protein 115356 indicates that, indeed, it is involved in the high virulence of ND90Pr on cv. Bowman, as the mutant is significantly reduced in virulence to the host, compared to the wild-type strain (Figs. II7C, S7). Thus, our comparative approach to analyzing secondary metabolite core proteins (NRPSs) led to the identification of a unique genomic region in C. sativus pathotype 2 isolate ND90Pr associated with high virulence on barley cv. Bowman that carries NRPSs which, when functionally manipulated, impacted virulence. Case Study 2: Among Cochliobolus species, the NRPS HTS1, which biosynthesizes the tetrapeptide HST HC-toxin, is unique to C. carbonum race 1 and has been demonstrated previously to be required for pathogenicity to hmhm maize [4,18,19]. HTS1, however, is present 60 in S. turcica and other fungi. Given the thorough documentation of HTS1 as a pathogenicity determinant, functional analyses were not necessary to cement the connection between its unique signature within the genus and its role as a virulence determinant. With the wider genomic resources reported here, we found orthologs of all four HTS1 AMP domains in S. turcica (ID# 29755) (Fig. II.8A). Manning et al., [61] also report orthologs in P. tritici repentis (ID# 12015) and Wight and Walton have found an ortholog in Alternaria jesenkae and demonstrated, furthermore, that the isolate makes HC-toxin [62]. In addition, there are HTS1 orthologs (APS1, Acc#: ACZ66258) in the Sordariomycete, Fusarium incarnatum/semitectum, that biosynthesize a different metabolite, apicidin [63]. In C. carbonum race 1 strain, SB111, the original strain in which the HTS1 locus was described, the structural organization of the cluster of genes encoding enzymes for HC-toxin production is complex and includes two copies of most genes, in two clusters residing in an ~ 600 kb region. The organizations of the S. turcica and P. tritici-repentis clusters are similar to each other, but different from that described for C. carbonum (Fig. II.8). Firstly, there is no evidence that the S. turcica and P. tritici-repentis genes are duplicated. Secondly, only orthologs of C. carbonum HTS1, ToxA, ToxE, and ToxF (C. carbonum HTS1 cluster nomenclature) proteins are clustered in S. turcica and P. tritici-repentis; orthologs of C. carbonum ToxC, ToxD, and ToxG proteins are found in both genomes, but on separate scaffolds in each genome (Fig. II.8). HTS1, ToxA, ToxE, and ToxF orthologs are present in the F. semitectum APS1 cluster, however there is no ToxC, ToxD, and ToxG in the sequenced cluster and, as genome sequence is not available for this species, we were unable to search for these genes. 61 ! Figure II.8. S. turcica has an ortholog of the C. carbonum NRPS HTS1 responsible for HCtoxin biosynthesis. A. Gene annotation and comparisons of the S. turcica, P. tritici-repentis, and F. semitectum regions carrying orthologs of the HC-toxin locus genes in C. carbonum strain SB111. Protein designations (color coded) correspond to C. carbonum and F. semitectum (APS) nomenclature. HTS1 is an NRPS, ToxA, E, F correspond to efflux pump, DNA-binding, and branched chain amino acid transaminase, proteins, respectively, and FAS α is a fatty acid synthase alpha subunit. Tox C (FAS beta subunit), ToxD (dehydrogenase), and ToxG (alanine racemase) in the cluster in C. carbonum, are not clustered in the other species but map to different scaffolds in the S. turcica and P. tritici-repentis assemblies. In C. carbonum, all of the known genes required for HC-toxin production are multicopy, in two linked, but separated clusters in a 600 kb region in isolate SB111; the genes are absent from toxin non-producing C. carbonum isolates that have been examined [122]. B. Portions of the full phylogenetic tree (Fig. S5A) showing placement of the HTS1 AMPs, extracted from tree to the left. HTS1 has four AMP domains cartooned bottom left. 62 Each C. carbonum AMP domain (red), groups, with high bootstrap support, with S. turcica protein 29755, a four AMP domain NRPS, (red), except for AMP4. In each of these matches, C. carbonum is represented twice, once by the SB111 AMP domain of the deposited sequence #AAA33023, and once as extracted by Augustus from our Illumina Velvet assembly of strain 26R-13. Note all HTS1 AMP domains group separately one from another and AMP2 is distant from the others. ! Thus the phylogenetic approach, when conducted with the suite of Cochliobolus genomes, pinpointed the NRPS, HTS1, as unique to C. carbonum race 1 and functional analyses done previously, prove that the metabolite is an HST required for pathogenicity. The inclusion of a genome from a genus sister to Cochliobolus, however, identified an ortholog in S. turcica. This, combined with reports of HTS1 orthologs in other phylogenetically scattered groups, indicates a complex genetic history. Case Study 3: C. victoriae has a NRPS that groups with high support with the A. fumigatus NRPS for Gliotoxin biosynthesis. In addition to the NRPSs extracted from our sequenced genomes, our phylogenetic trees included NRPSs producing known products, such as the two AMP domain A. fumigatus NRPS, GliP, for gliotoxin biosynthesis and the Leptosphaeria maculans NRPS, SirP, for sirodesmin production, both epipolythiodioxopiperazine (ETP) toxins. Two C. victoriae AMP domains on node 1179 clustered with 99-100% bootstrap support with A. fumigatus GliP AMP1 and AMP2 (Fig. II.9, Table II.8). Both were evolutionarily closer than the corresponding SirP AMP domains. Thus, our objective to determine if any of the unknown NRPSs grouped with NRPSs with characterized products yielded a C. victoriae candidate for production of gliotoxin or a related metabolite. Furthermore, examination of the neighborhood surrounding the C. victoriae bimodular NRPS, indicates that all of the genes in the A. fumigatus gene cluster [64] are present in a cluster in C. victoriae (Fig. II.9A). 63 Figure II.9. C. victoriae has an ortholog of A. fumigatus GliP responsible for gliotoxin production. A. Gene annotation and comparisons of the C. victoriae and A. fumigatus regions carrying orthologs of the Gliotoxin biosynthetic proteins. Protein designations (color coded) correspond to A. fumigatus nomenclature [64]. In A. fumigatus, GliP is an NRPS, GliT, F, N, A, G, M, C, J, I and Z correspond to oxidase, cytochrome P450, methyl transferase, transporter, glutathione S-transferase, O-methyltransferase, cytochrome P450, dipeptidase, aminotransferase and Zn finger proteins, respectively. GliK is of unknown function. In C. victoriae, ‘ORF’ = unknown function. B. The bimodular (2 AMP domains) C. victoriae NODE 1179 NRPS (Augustus gene call g7087) is an example of the phenomenon of spotty conservation of NRPS AMP domains across fungi. L. maculans also has an ortholog (SirP, producing sirodesmin), however, this is not as closely related as the C. victoriae ortholog. Of the two AMP domains that comprise the C. victoriae GliP ortholog, AMP2 is present in C. carbonum (NODE 464 g2585) and AMP1 is 64 found in C. sativus (ID 103953). Neither possesses both, although additional, related sister domains are found in other Cochliobolus species. Cartoon (bottom) shows the A. fumigatus, C. victoriae and L. maculans NRPS orthologs color coded as to AMP domain. Branches carrying GliP orthologs extracted from full phylogenetic tree (Fig. S5A) to left. Gene/AMP nomenclature and bootstrap values as described in Fig. II.4. v. Summary To thoroughly understand the evolutionary history of multimodular NRPSs, AMP domains were analyzed as individuals using a combination of amino acid alignments and phylogenetic tree building. Results show that, within a species, NRPSs are highly conserved, but conservation dissipates as comparisons are made across the genus. Thus, diversity of these genes, their encoded proteins and corresponding metabolite potential, is truly enormous. Strainunique NRPSs are primary suspects for producing small molecules conferring high virulence or host specificity. A robust example of this are the species- and strain-unique C. sativus ND90Pr NRPS proteins 115356 and 140153 which map to the unique VHv1 high virulence conferring region, for which gene deletion confirms a role in cultivar specific virulence. The phylogenetic structure of NPS1 and NPS3 enzymes suggest that corresponding genes undergo rapid duplication and expansion and could act as a cauldron for the formation of new NPS genes. 6. Secondary metabolism: polyketide synthases Polyketide synthases (PKSs), like NRPSs, are large multidomain enzymes that produce small molecules (polyketides) with functions that include HSTs. The suites of PKS encoding genes (PKS) in the C. heterostrophus C4 and C5 genomes were identified and annotated previously [65]. To address degree of conservation and evolutionary relationships of PKSs in our subject species in order to make inferences about function, we used the PFAM ketosynthase 65 domain (KS) HMM as a query to search for orthologs in the additional strains of C. heterostrophus and other species, and the related maize pathogen, S. turcica. i. Conservation of known reference strain C. heterostrophus C5 polyketide synthases Most C. heterostrophus PKSs are conserved across all C. heterostrophus strains, although PKS16 is absent from the genome of strain Hm338 and PKS25 is absent from strain Hm540 (Table II.9, Fig. II.10, for the full phylogenetic trees see Fig. S6, and Table S10 for master inventories). PKS13 is a pseudogene found only in strain C5 (and C4). As with the NRPSs, conservation of PKSs across the Cochliobolus genus is not as high as within C. heterostrophus species and is even less when the related genus, S. turcica is considered. Seven out of the 23 PKSs in reference strain C5 are conserved in all Cochliobolus species and S. turcica (Table II.9, Fig. II.10); the only known product of these is melanin (produced by C. heterostrophus PKS18). Otherwise, the products of conserved PKSs are unknown. Three PKSs are present in all Cochliobolus genomes, but not S. turcica (Table II.9). Two PKSs are unique to C. heterostrophus, while nine are present discontinuously throughout the species examined. Blast searches using the predicted protein sequences of C. sativus isolate ND90Pr PKSs as queries against the genome sequences of the isolate ND93-1 (a pathotype 0 isolate) indicated that all PKSs predicted in ND90Pr were found in ND93-1, except one (ID# 184740). ii. Species-unique PKSs As for the NRPSs, we anticipated that species-unique PKSs (PKSs possessing KS domains lacking orthologous KS domains with bootstrap support in other species) would be the likeliest candidates for virulence functions, given the well-documented roles of HSTs in virulence and their corresponding unique or spotty distribution patterns in related strains. C. heterostrophus race O has 23 PKSs and no unique ones, while C. heterostrophus race T has 25 66 PKSs, two of which are unique. C. victoriae, C. miyabeanus, and C. sativus have 21, 21, and 18 PKSs, respectively, and no unique ones; C. carbonum has 27 PKSs including two unique ones, and S. turcica has 27 PKSs, including 13 unique ones (Table S10). Table 5. Table II.9. Conservation of C. heterostrophus strain C5 polyketide synthases in other strains and species ChGene Genbank C4 JGI C5.V3 In all C. In all In S. turcica? protein ID Protein ID heterostrophus Cochliobolus strains? spp? PKS1 ABB08104 Race T - - PKS2 ABB76806 Race T - - PKS3 AAR90258 1098212 + + - PKS4 AAR90259 96868 + Cs - PKS5 AAR90260 1103337 + ++ PKS6 AAR90261 1168707, + Cv, Cc, Cm + 1229323 PKS7 AAR90262 1118456 + Cv, Cc, Cm - PKS8 AAR90263 28817 + Cc, Cm, Cs + PKS9 AAR90264 1104189 + ++ PKS10 AAR90265 96669 + Cv, Cc + PKS11 AAR90266 1112706 + Cc, Cs - PKS12 AAR90267 1216295 + +, 2x Cc + PKS13 AY495653 75987 - PKS14 AAR90268 67271 + +, 2x Cm + PKS15 AAR90269 108173 + + - PKS16 AAR90270 33896 not in Hm338 - + x2 PKS17 AAR90271 1105179 + - - PKS18 AAR90272 34478 + ++ PKS19 AAR90273 81477 + ++ PKS20 AAR90274 1108855 + - - PKS21 AAR90275 1029307 + Cv, Cc, Cm, Cs - PKS22 AAR90276 105287 + Cm + PKS23 AAR90277 77059 + ++ PKS24 AAR90278 1209664 + + - PKS25 AAR90279 1034546 not in Hm540 - - + = present, - = absent; 2x = two copies; Strains Ch = C. heterostrophus, C4, Hm338, Hm540, PR1x412 are C. heterostrophus strains, Cs = C. sativus, Cv = C. victoriae, Cc = C. carbonum, Cm = C. miyabeanus, St = S. turcica, for Ch C4 Genbank protein IDs are given, for Ch C5 JGI protein IDs are given See Fig. S6A, and Table S10 67 Figure II.10. Cartoon of cross-species phylogenomic analyses of individual ketosynthase domains from PKS proteins. The ketosynthase (KS) domains were extracted from all five C. heterostrophus and from the C. victoriae, C. carbonum, C. miyabeanus, C. sativus and S. turcica genomes. See Fig. II.3 for species designations, color codes and format. KS domains colored black and therefore absent in analyzed genomes include outgroups and KS domains in animal fatty acid synthases (FAS). KS domains not grouping with the previously annotated C. heterostrophus set are labeled as ‘New _1 through _10’. Gene/KS nomenclature and bootstrap values as described in Figs. II.3 and II.4 for AMP domains. 68 iii. Discontinuously distributed and expanded PKSs There were ten clusters (designated ‘New’ 1-10) of PKS genes that did not have an ortholog in C. heterostrophus (Figs. II.10, Table S10). Some of these clusters, such as ‘New_2 or New_5’, had representatives of only a few species. Others such as ‘New_3 or New_6’ contained representatives of all species except C. heterostrophus. Two clusters (New_1 and New_8) were not sister to a group with a C. heterostrophus reference PKS, but contained an ortholog from a C. heterostrophus field strain (Fig. II.10). The only expanded group of PKSs from the C. heterostrophus set was PKS14, which had two orthologs in C. miyabeanus (Table II.9, Fig. II.10). Otherwise, C. heterostrophus PKSs were either conserved as single copies, discontinuously present in single copy, or C. heterostrophus unique. Expansion did not seem to occur centered around a certain set of ‘birthing reservoir’ genes, as for NPS1 and NPS3 AMP domains (Fig. II.6). iv. PKSs showcasing the value of the phylogenomic approach in pinpointing candidate virulence determinants Case Study 1: All race T strains of C. heterostrophus have two PKSs not found in race O strains or any other known species. PKS1 and PKS2, genes required for biosynthesis of T-toxin, are present in all C. heterostrophus race T strains but absent from all C. heterostrophus race O strains and all Cochliobolus species examined to date (Figs. II.10, II.11A, Table II.9). In previous work, we demonstrated that strains deleted for either of these two PKS genes fail to make T-toxin and are much reduced in virulence on T cytoplasm corn [45,48,66,67]. This racespecific PKS example mirrors the aforementioned NRPS examples, i.e., the C. sativus ND90Pr region carrying the NRPSs 115356 and 140513 (Fig. II.7) and the C. carbonum region encoding HTS1 (Fig. II.8). 69 Figure II.11. The two PKSs responsible for T-toxin production by race T of C. heterostrophus are unique to race T. A. Mauve alignment [44] of the sequences of three race T (C4, Hm338, PR1x412) and two race O (C5, Hm540) strains of C. heterostrophus. Sequences in common across all five genomes are colored ‘mauve’. The three orange blocks correspond to regions found only in race T genomes and carry the PKS1 gene essential for T-toxin biosynthesis and high virulence on cytoplasmic male sterile corn. 70 B. PKS1 and PKS2 are found in race T strains only and are unrelated by phylogeny. In C. heterostrophus, PKS1 is most closely related to PKS7 (high bootstrap support), which is found in both race T and race O strains, as well as C. victoriae, C. carbonum, and C. miyabeanus, but not in C. sativus or S. turcica. PKS2 groups in a different location with PKS3, but without bootstrap support, and is found in all strains examined except S. turcica. Full phylogenetic tree is to left (Fig. S6A). The reference set of C. heterostrophus strain C4 PKSs are shown by their Genbank numbers [e.g., Ch AAB08104, which is PKS1 (Table 9)] and their Augustus gene call (C4 scaffold 74 g10456) and thus are in the tree in duplicate. C5 proteins are indicated as JGI protein ID (e.g., CocheC5 1118456). For more information on evolutionary relationships of PKS1 and PKS2 see [45]. Case Study 2: A S. turcica-specific PKSs is up-regulated in planta. Three S. turcica PKSs grouped together in a S. turcica- specific clade, labeled ‘New_10’ (Fig. II.10, Table S10). Using real time PCR, we examined in planta expression of one of the S. turcica unique genes (ID 161586) and found that expression was increased 560-fold at three days post inoculation (Fig. S8), then dipped and rose again to the same level at seven days post inoculation. Although we haven’t yet deleted this gene, based on other test cases, it is tempting to couple the in planta expression pattern with a possible role in virulence. v. Summary Like NRPS proteins, the PKSs examined were either highly conserved, partially conserved, or strain unique. Some orthologs had duplicated members for some species, but this expansion did not orbit a particular set of genes such as NPS1 and NPS3. PKSs identified as strain or species-unique include characterized, as well as unknown, candidate virulence factors. The race T unique C. heterostrophus PKS genes PKS1 and PKS2 are examples of characterized unique, highly specific virulence factors. Further characterization of strain-unique PKSs, such as S. turcica ID 161586, which is highly expressed in planta (Fig. S8) could reveal novel virulence factors. 71 7. Location of C. heterostrophus NPS and PKS genes Several publications demonstrate that species/strain unique sequences tend to reside in variable regions of the genome such as in subtelomeric locations [68,69] and dispensable chromosomes [70]. All C. heterostrophus reference strain C5 NPS and PKS genes were mapped to the assembled linkage groups (Fig. II.2). For NPSs, 13 of the 14 total could be mapped to one of the 16 linkage groups/chromosomes, and 6 of the 13 were <200 kb from the end of the linkage group. Two (NPS 5, 9) of the six are unique to C. heterostrophus and two (NPS1, 11) have limited distribution in Cochliobolus spp.. For PKSs, 22 of the 25 total could be mapped to one of the 16 linkage groups, 9 of the 22 were <200 kb from the end of the corresponding linkage group and one (PKS25) mapped to the B chromosome. Five (PKS 13, 16, 17, 20, 25) of the ten are unique to C. heterostrophus and one more (PKS11) has limited distribution in Cochliobolus spp.. In sum, approximately half of the NPSs and PKSs map to scaffold ends, in some cases with mapped telomeres (Fig. II.1). As chromosome ends are notoriously variable, this placement could indicate a mechanism for patchy phylogenetic distribution of these genes. Note that the two PKSs involved in T-toxin production by race T are absent in race O strain C5, but map (genetically) to the breakpoints of race T chromosomes 12;6 and 6;12 which are reciprocally translocated with respect to chromosomes 6 and 12 in race O C5 [9]. Note also that PKS3, which has a phylogenetic relationship, but without bootstrap support, to PKS2 (Figs. II.10, II.11B), maps internally to race O chromosome 6 (Fig. II.2), and that PKS7, which is the closest (with bootstrap support) C. heterostrophus PKS to PKS1 (Fig. II.11B), maps to the end of unplaced scaffold 20 (Fig. II.2). 72 8. Small Secreted Proteins (SSPs) To identify candidate effector proteins, we searched the gene catalog of each species for proteins that were cysteine rich (> 2% cysteine), small (<200 amino acids), predicted to be secreted (using Phobius [71]), and without transmembrane domains. Between 141 and 289 SSPs per genome (Table II.10) were identified with C. sativus ND90Pr containing the most and C. heterostrophus Hm338 the fewest. We next conducted an all versus all blast analysis to determine if SSPs were strain or species-unique, using an 80% blast cutoff. Very few C. heterostrophus SSPs were unique to any particular strain as most could be found in at least one other C. heterostrophus field or lab strain. Using this approach, we identified between one and 21 unique SSPs (Table II.10, master inventory Table S11). We found more strain-unique SSPs in the other Cochliobolus genomes, as our analysis included five C. heterostrophus strains. S. turcica and C. sativus had the most isolate-unique SSPs, containing 191 and 167 candidates, respectively. As these are the two strains thought to act as hemibiotrophs, it is interesting that they contain more SSPs, and more unique SSPs, than the necrotrophic isolates, although this is only a correlation at this point. Table II.10. Comparative small secreted protein candidate effector inventories Strain Total Strain unique SSPs total 1753a 506 Ch C5 180b 14 Ch C4 171 21 Ch 338 141 1 Ch Hm540 151 4 Ch PR1x412 151 8 Cc 26R13 153 24 Cv FI3 160 25 Cm WK1C 143 51 Cs ND90Pr 289 167 St 28A 214 191 a Total SSPs for all strains examined b Total for each strain (e.g., Ch C5) 73 The C. heterostrophus C5 assembly has 180 predicted secreted proteins matching the criteria. We examined each of these in the JGI browser with respect to EST support, SNPs, and predicted functional domains. Seventy-two of these calls had absolutely no EST support, while 24 calls had incomplete EST support (ESTs matching some portion, but not all, of the gene call), leaving 84 with complete (spanning the entire gene model) EST support (Table II.11). Genes in the no-EST support category are of special interest, as they may be specifically expressed in planta and thus not expressed under conditions used for preparing our EST libraries (fungus grown in vitro on a variety of complete (CM) and minimal (MM) media, and mixed into CM or MM pools). Lack of strong EST support may also suggest an erroneous gene call. Table II.11. C. heterostrophus strain C5 SSP candidate effector analysis Expression data In complete medium 84 In minimal medium 24 No expression data 72 Predicted Domains and Conservation None None, but present in other orders PFAM domain predicted 120 23 37 SNPs SNPs in at least one Cochliobolus species no SNPs 101 79 As is typical with candidate effectors, functional domain predictions were lacking, with only 37 candidates having some predicted function, generally involved in cell wall or extracellular matrix function (Tables II.11, S11). An additional 23 candidates were conserved in other fungi outside of the Dothideomycetes. The remaining 120 calls were featureless and seemingly unique to the Dothideomycetes. C. heterostrophus strain C5 SSP calls were rich in 74 SNP calls to other Cochliobolus genomes: 101 candidate SSPs had SNPs with at least one other Cochliobolus genome. In our all versus all blast analysis, only 6 of the 180 C. heterostrophus C5 SSPs were found in all 10 strains examined and 14 were unique to strain C5 (Table II.11). The presence or absence of most SSPs did not fall into easily categorized bins such as C. heterostrophus-specific, or maize-pathogens only. Instead, SSPs were present and absent in no particular pattern across the genomes. 115 SSPs were present in at least one other species (C. victoriae, C. miyabeanus, C. carbonum, or S. turcica), with seven found in all species, and 27 in all Cochliobolus species. SSPs mapped to all scaffolds larger than S26 (Fig. II.2). Unlike those in some phytopathogens, such as Leptosphaeria maculans [72], SSP encoding genes did not occur in clusters; candidates seldom were located within 10 kb of each other (Fig. II.2). These genes were, however, often located in or near regions we identified as C. heterostrophus species unique (Fig. II.2). 75 C. Discussion The genomes of five C. heterostrophus strains, two C. sativus strains, three additional Cochliobolus species (C. victoriae, C. carbonum, C. miyabeanus) and S. turcica, a member of a close sister genus, were sequenced and compared, to identify unique genomic regions and to inventory secondary metabolism and SSP encoding genes. This dataset is distinctive in that it allows us to contrast genomes that are very closely related, yet differ in several key ways. First, our dataset includes highly related pathogens of several different host plants (corn, wheat, rice, barley, Brachypodium). Second, this set includes pathogens that are highly virulent on specific cultivars of a particular host (i.e., C. heterostrophus race T on Tcms maize, C. victoriae on Vb oats, C. carbonum on hmhm maize), as well as more generalist pathogens such as C. sativus that can cause disease on multiple hosts (barley, wheat, Brachypodium). Third, the group includes two hemibiotrophs, S. turcica and C. sativus, while the rest are necrotrophs. Fourth, we can make graduated comparisons at different levels of parental and phylogenetic relatedness as we progress from genomes of the same inbred line, species, genus, and family. We have used this final point as a probe to attempt to understand the significant genomic differences between strains that shape host choice, specificity, and lifestyle. 1. Structural differences across strains and species Our whole-genome alignment data support graduated degrees of similarity at the highly inbred strain, field strain, species and genus levels. C. heterostrophus strains C4 and C5, offspring of successive backcrosses [42], were highly similar to one another, with 20 fold fewer SNPs than pairwise comparisons of reference strain C5 to C. heterostrophus field strains. This 76 remarkably low number of SNPs highlights the power of selective inbreeding in establishing uniformity across the genome. The two C. sativus field strains, when aligned to each other, had a comparable number of SNPs to those of C. heterostrophus field strains aligned to reference C. heterostrophus strain C5. Other Cochliobolus genomes had roughly 50 fold more SNPs than C. heterostrophus field strains when aligned to the C. heterostrophus C5 reference. This level of similarity was seen when comparing any two Cochliobolus species to one another, with the exception of comparing C. victoriae to C. carbonum. These two species are capable of successfully mating, although progeny of crosses are unable to cross to each other or to their parents (Turgeon lab, unpublished). We have hypothesized that C. victoriae may have evolved from a C. carbonum strain [49]. This similarity is seen at the whole genome level, as C. victoriae and C. carbonum share an intermediary number of SNPs compared to C. heterostrophus inter- and intra-species comparisons. Our SNP data show that approximately one quarter of the genome differs between Cochliobolus species and that only about one tenth of this is found in segments larger than 5 kb. We and others [72,73,74] have recently introduced the term mesosyteny [1] to describe organizational conservation between species. Genetic content is conserved across chromosomes, but not co-linearly. It seems possible that our findings here, showing that many small, scattered differences summing to significant quantitative differences (i.e., 25% dissimilar), could be the product of the same mechanisms that result in mesosyntenic patterns. Pathogens of the same host (e.g., C. carbonum and C. heterostrophus on maize) or lifestyle were not more similar to each other than those of different hosts; instead overarching genetic patterns followed phylogenetic lines. As it is estimated that the Pleosporaceae arose as a group less than 20 MYA 77 (see Ohm et al.,[1]) and the genus Cochliobolus is young in the group, genome comparisons provide us with an overall picture of a timeline of how genome diversity varies with speciation. Our intra-species SNP tallies are comparable to SNP tallies found when strains of other species are examined. For example, there were 10,495 SNPs called between two Fusarium graminearum strains [75], and a range of 13,274-188,346 SNPs called for 18 Neurospora crassa classical genetic mutants [76]. Both dataset tallies are in the same range as our C. heterostrophus field strain comparisons. With respect to SNPs in different species of the same genus, it is unusual that we were able to perform a whole-genome SNP analysis at all, without limiting our scope to coding sequence. We owe this to the very close phylogenetic relationship of these species. 2. Strain-, species-, and genus-specific genes for secondary metabolism Although individual functional domains of NPS/PKS proteins can be identified bioinformatically, attempting to predict their corresponding metabolite product is challenging. The genes encoding these proteins evolve rapidly and through complex mechanisms [57,65] and whole gene alignment methods provide misleading or unclear results when determining presence or absence of a particular NPS or PKS gene. Here, we extracted individual conserved signature catalytic domains, i.e., the AMP-binding domains from mono- or multi-modular NRPSs or the ketosynthase (KS) domain from multidomain PKSs using customized HMM models, then built alignments and phylogenetic trees with these individual units to determine the presence or absence of whole or partial NRPS and PKS proteins, and their evolutionary relationships. In our opinion this is a necessary first step towards understanding evolutionary history of the corresponding genes and the possible small molecules produced by these highly diverse proteins. 78 We found that within a Dothideomycete genus, in this case Cochliobolus, approximately half of the NPS and a third of the PKS genes are well conserved (present in all strains). When related S. turcica was considered these numbers dropped to a third and a fifth, respectively. The rest were found to be poorly conserved or species-unique when the highly curated C. heterostrophus gene sets were used as reference. The small molecules produced by the corresponding non-conserved proteins are largely uncharacterized, but the differences between strains and species imply that the potential for production of biochemically unique molecules is large and considerably beyond that expected for closely related strains (housekeeping genes share ~95% identity). These findings refine our understanding of NPS and PKS genes, as very few are conserved. For example, only two NPS genes and one PKS gene were found when 18 Dothideomycete genomes were analyzed [1]. Broadly conserved secondary metabolism genes, where they have been characterized, produce small molecules that serve basic cellular functions (ferricrocin, melanin [58,77,78,79]). Poorly conserved NPS and PKS genes, while still largely uncharacterized, can include those involved in host-specific high virulence. 3. NPS1, NPS3, and NPS13 embody the intriguing genetic origins of NRPS proteins NPS1 and NPS3 AMP domains are discontinuously distributed and expanded across the Cochliobolus and Setosphaeria isolates sequenced and are sources of much of the NRPS diversity (Figs. II.5, II.6 and [57]). The individual AMP modules do not cluster by protein, but instead, NPS1, NPS3 and NPS13 AMP domains occur in two distinct, and mixed, groups (Fig. II.6). Strikingly, each Cochliobolus species possesses either a complete C. heterostrophus NPS1 or NPS3 ortholog, but never both. Furthermore, all species have one or more additional NRPS proteins consisting of NPS1/NPS3 related domains, and a NPS13 related domain that is absent 79 from all C. heterostrophus genomes except Hm540 (Fig. II.5). The C. heterostrophus Hm540 genome includes all four corresponding genes: NPS1, NPS3, NPS13, and the additional NPS1/NPS3/NPS13 gene (Fig. II.5). The pattern of duplication and loss appears to have been very rapid to account for this distribution, and is further complicated by the presence of additional bi-, mono-, tri-, and tetra-modular proteins, particularly in C. sativus, whose AMP domains group with NPS1/NPS3/NPS13 proteins, (Figs. II.5, II.7, S5). 4. Evolutionary origin of NPS and PKS genes The origin of strain or species unique secondary metabolism genes is of great interest and horizontal gene transfer is a common, but not the only, explanation for their appearance [65,80,81,82]. The volatility of the NPS1, NPS3 and NPS13 family raises the possibility that genes we presume are horizontally transmitted could have vertical histories obfuscated by species and strain sequence depth. We speculate that partial or whole genes encoding individual domains or whole proteins recombine and expand quickly, and, when they confer high virulence, as in the case of HSTs, can spread rapidly throughout a population. When the susceptible host allele is not present in the population, the gene is lost or not conserved in the majority, but not the entirety, of the population, as is the case for C. heterostrophus race T and genes for T-toxin production; race T is difficult to find in the field currently [43]. As we sequence more and more isolates, we might find that the T-toxin genes are present in strains of many more Dothideomycetes than we originally expected. This is certainly the case with the HC-toxin genes which is not so surprising, given that the Hm alleles are present in most plants. 80 5. Pinpointing virulence-associated secondary metabolite genes Identifying secondary metabolites that function as virulence factors (such as HSTs) is a primary goal when studying a pathogen’s genome. The impact of HSTs was realized early on because they render the producing fungi pathogenic or highly virulent to principal crops. Thus, most were characterized physiologically and genetically decades ago [4,18,19,40,83,84,85]. The pivotal point of our comparative analyses is the strikingly obvious observation that secondary metabolite genes, when unique to a species or strain, are likely to encode a virulence determinant. We provide several examples. The first example is the C. heterostrophus PKS1 and PKS2 genes involved in production of the HST T-toxin. These genes reside in 1.2 Mb of DNA, not found in race O and located at the breakpoints of two race T chromosomes (12;6, 6;12), reciprocally translocated with respect to race O counterparts (chromosomes 6, 12). The T-toxin genes are not in race O or any other Cochliobolus species. Deletion of either PKS eliminates T-toxin production and drastically reduces virulence of the fungus on Tcms maize, as reported earlier [45,67]. Known Tox1 genes, such as PKS1 are on very small scaffolds (~25 kb) in race T strains C4, Hm338, and PR1x412 (Fig. II.11), which cannot be further assembled due to the repetitive and AT-rich nature of the locus. Thus the physical structure of the Tox1 locus remains elusive, but its association with a unique genomic region, however complex, is clear-cut. Although T-toxin is unique to C. heterostrophus, a closely related fungus, Didymella zeae maydis (formerly, Phyllosticta maydis, Mycosphaerella zeae maydis), produces a polyketide HST, PM toxin, with the same biological specificity as T-toxin. The central PKS for PM-toxin is the closest PKS to C. heterostrophus PKS1, but still only ~ 60% identical at the amino acid level and organization of the cluster of 81 genes required for toxin production differs; in D. zeae-maydis, the genes are present in a single tight cluster [86,87]. The second example, examined here, is the NRPS, HTS1, for HC-toxin production. The genes for HC-toxin produced by C. carbonum, were identified two decades ago in a tour de force molecular manipulation exercise [88,89]. A strong genomic signature attends these genes as they reside in an ~ 600 kb region not found in other races of C. carbonum, or in any of the additional Cochliobolus genomes examined then or here. Two copies of a cluster of HC-toxin genes are located in this region and both copies of the core NRPS, HTS1, had to be deleted to demonstrate elimination of toxin production and reduction of virulence [88,89]. In current investigations, we, Manning et al., [61] and Wight and Walton [62] have discovered that HCtoxin like genes are present in S. turcica (Fig. II.8), P. tritici-repentis and A. jesenskae, respectively. These genes are also apparent orthologs of the genes for apicidin (APS1) production by some Fusarium species [90]. Thus, genes for HC-toxin or HC-toxin-like metabolites are more broadly distributed than previously thought. In terms of amino acid identity, the S. turcica and P. tritici-repentis NRPS HTS1 proteins are 79% identical at the amino acid level, but identity drops to 39-43% when these are compared to the C. carbonum HTS1 or APS1 proteins. The S. turcica and P. tritici-repentis HTS1 orthologs lack the C-terminal condensation domain found in C. carbonum HTS1 and F. semitectum APS1, suggesting S. turcica and P. tritici-repentis make a different product. Whether or not S. turcica and P. triticirepentis are capable of producing HC-toxin is unknown, however, it has been reported [62] that A. jesenskae does. That HC-toxin producing capability might be found in pathogens other than C. carbonum is not unreasonable, considering the maize defense gene Hm1, necessary to detoxify the toxin, is found in all grasses [91]. 82 The third example concerns C. sativus. Two of the NPS genes unique to C. sativus ND90Pr (IDs 115356 and 140513, Fig. II.7) are present at the VHv1 locus associated with high virulence on cultivar Bowman. The entire VHv1 locus is absent in the low virulence isolate, ND93-1, and the two NPSs are not found in any other genomes examined here, or in Genbank. Our data show that these genes are up-regulated 12 hrs after inoculation, and that deletion of one of them, (115356), significantly reduces virulence on barley cultivar Bowman (Fig. II.7). Recent work on deletion of the gene encoding 4'-phosphopantetheinyl transferase provided indirect evidence that a secondary metabolite is involved in the biosynthesis of the virulence factor in ND90Pr [92]; our current work directly confirms this. The phylogenetic location of these VHv1 NPS genes is revealing in that they are either in branches with no close sister members (ID# 140513, Fig. S5A) or in the NPS1/NPS3/NPS13 expansion clade (ID# 115356, Figs. II.5, II.6). In addition to these two genes, we have evidence based on real time expression data on RNA from inoculated barley, that the C. sativus genes corresponding to protein ID#s 49884 and 130053 are also up-regulated at 12 hrs post inoculation. These genes are found in the same C. sativus specific clade (New_8, Figs. II.3, S5A) as protein ID# 140513, and although we have not yet made mutants, our prediction is that these will also contribute to virulence. Our analyses of the hemibiotroph, S. turcica, is in its infancy, however, as noted in the Results we have identified 13 unique PKSs, three of which (ID# 161586, 30113, 34554) grouped together in a S. turcica- specific clade, called ‘New_10’ (Figs. II.10, S8, Table S10). As preliminary support for the importance of unique PKSs, we used real time PCR, to examine in planta expression of one of these S. turcica unique genes (161586) and found that expression was indeed increased (>500 fold) by three days post inoculation. Although we haven’t yet 83 deleted this gene, it is tempting to predict that the in planta expression pattern is indicative of a role in virulence. One of the species-unique NRPSs in C. victoriae (on node 1179, gene #7087) is a NRPSs with two AMP domains clustering with 99-100% bootstrap support to AMP domains from the bimodular A. fumigatus NRPS, GliP, which produces the ETP toxin, Gliotoxin. Related to these NRPSs is the L. maculans NRPS, SirP, which produces sirodesmin. Candidate orthologs of these NRPSs have been reported in Chaetomium globosum, Magnaporthe oryzae, and Fusarium graminearum [93]. Gliotoxin is associated with virulence of A. fumigatus to immunecompromised patients [94]. Functional characterization of the newly discovered C. victoriae counterpart is necessary to determine the type of ETP produced and whether or not it might play a role in virulence, as Gliotoxin does. Note the entire Gliotoxin gene cluster [64] is present in C. victoriae (Fig. II.9). Gene knockout and screening for alteration in virulence to oats, due to victorin production, indicates no change from that of wild type (Wu, Turgeon, unpublished). This C. victoriae NRPS is not found in other Cochliobolus genomes, yet it clusters with A. fumigatus GliP, exemplifying the patchy distribution signature of most members of the NPS family of genes. 6. Effectors and lifestyle Effectors are pathogen produced small secreted proteins (SSPs)/small molecules that interact with the host plant to promote disease. Effectors historically were called avirulence proteins (as their discovery hinged on association with a corresponding plant resistance gene), but we now recognize that effectors are virulence factors that aid the pathogen by specifically targeting aspects of host cell defense and recognition. Evading detection is a necessary strategy 84 for (hemi)biotrophs, where triggering the host hypersensitive response curtails disease. Necrotrophs, on the other hand, benefit from the death of host cells, and have evolved molecules such as HSTs, like victorin which subverts function of an R gene (Lov1/Pc-2) to trigger susceptibility and plant cell death intentionally [14,15,95]. Protein HSTs such as P. triticirepentis and Stagonospora nodorum ToxA are clear examples of secreted, necrotrophic, proteinaceous, host-selective virulence factors acting to effect virulence in host cells, like any other effector, but which, in the presence of a R-protein look-alike, is necessary for susceptibility [96]. The lingering question is whether or not necrotrophs utilize SSP effectors in the traditional (and difficult to identify) sense of micromanipulation of the host environment, or, instead, use effectors to trigger host cell death through abuse of (hemi)biotrophic defenses. In this regard, given our clear discovery that at least one NRPS metabolite (ID# 115356), when deleted has a much reduced phenotype reminiscent of a necrotrophic HST phenotype, we question whether C. sativus should truly be considered a hemibiotroph or a necrotroph. On the other hand, our SSP analysis shows that C. sativus and S. turcica have an expanded SSP repertoire compared to the other species examined, which is consistent with a hemibiotroph strategy, i.e., arsenals of effectors are used to evade host detection. The repertoire of candidate effectors in necrotrophs, nevertheless, is quite large. If only a small subset of these is involved in virulence, it would mean that Cochliobolus, and perhaps other necrotrophs, use effectors more expansively than is recognized. This is a difficult question to address, and our in silico analysis requires experimental confirmation of in planta expression and secretion before we can be sure Cochliobolus species utilize protein effectors. Perhaps a strategy prioritizing species or strain unique regions would aid characterization attempts. Bearing on this point, the SSP catalogue differed markedly from secondary metabolites in their conservation across, and 85 within, species. Only six of the 180 (3%) C. heterostrophus C5 SSPs were identified in all genomes examined (including S. turcica), unlike the 7/25 (28%) PKSs and 6/14 (43%) NRPSs. Considering C. heterostrophus genomes only, 27 of these 180 SSPs were present in each (15%), again, far fewer than the 21/25 (84%) PKS and 13/14 (93%) NRPS C. heterostrophus C5 genes conserved throughout all C. heterostrophus genomes. This indicates that, more so than secondary metabolite genes, SSP encoding genes are extraordinarily volatile in the evolutionary history of the genus. 7. Final thoughts The stories of the SCLB and Victoria blight epidemics are dramatic examples of interactions between crops, whose ‘evolution’ is driven by human intervention (breeders) and their pathogens, which evolve naturally to exploit new genetic susceptibilities. Both the Tcms and Pc-2 genes were introduced into maize and oats, respectively, by breeders fewer than 30 years before the epidemic outbreaks. Specifically, Tcms was discovered in the 1940’s, incorporated into elite corn inbred lines increasingly throughout the 1960’s, and was present in almost all of the hybrid corn in the US by 1970. The vast monoculture of Tcms maize was the perfect host for the previously unknown race. Species of Cochliobolus spp. clearly have proven their ability to cause extraordinary crop losses. As we begin to understand the intimidating capacity for diverse production and evolution of new HSTs, we must also look for ways to apply this knowledge to our disease response strategies. 86 D. Materials and Methods 1. Strains C. heterostrophus strains sequenced by JGI included inbred strains C5 (ATCC 48332, race O, MAT1-1, Tox1-) and C4 (ATCC 48331, race T, MAT1-2, Tox1+) and field strains Hm540 (geographical origin North Carolina, race O, MAT1-1, Tox1-), Hm338 (New York, race T, MAT1-2, Tox1+, ATCC 48317), and PR1x412, (a progeny of a cross between PR1C from Poza Rica, Mexico and strain 412, unknown geographical origin , race T, MAT1-1, Tox1+). In addition, the genomes of C. victoriae strain FI3 (unknown geographical origin, MAT1-2, victorin+), C. carbonum strain 26-R-13 [MAT1-1, HC-toxin+, a progeny of a cross between C. carbonum strains 2-R-6 (alb2; MAT1-1) and Five Points (unknown geographical origin, MAT12) performed by Dr. Steve Briggs]. C. miyabeanus strain WK1C (Wuankuei, Yulan county China, MAT1-2), C. sativus isolate ND90Pr (North Dakota, ATCC 201652, MAT1-2, pathotype 2 on barley cv. Bowman) and S. turcica strain St28A (New York, race 2,3,N, MAT1-1) were sequenced by JGI. C. sativus isolate ND93-1 (North Dakota, ATCC 201653, MAT1-1, pathotype 0 on barley cv. Bowman) was sequenced at the University of Hawaii. 2. Genomic resources The highly inbred C. heterostrophus reference race O lab strain C5 was sequenced using the Sanger whole genome shotgun approach, with paired end reads and improved by manual finishing and fosmid clone sequencing (http://www.jgi.doe.gov/sequencing/protocols/prots_production.html). Four different sized libraries were sequenced: 3.1kb, 6.8kb, and two fosmid libraries (32.3kb and 35.3kb), to a total 87 coverage of 9.95x. ESTs were generated by growing strains in complete and minimal medium under many conditions [on complete and minimal medium, on sexual reproduction plates, stress medium (-N, -Fe, etc)] and pooled as complete or minimal samples for sequencing and support of gene annotation. The genome of isogenic C. heterostrophus race T strain C4 was sequenced using Illumina technology (300bp insert size, 2x76bp reads to a nominal depth of 200x), assembled using Velvet [97] and AllPathsLG [98] and annotated using ESTs from C. heterostrophus strain C5. The genome of C. sativus pathotype 2 isolate ND90Pr was sequenced using a hybrid approach, which included 40kb fosmid Sanger reads, shredded consensus from Velvet assembled Illumina data (300bp insert size, 2x76bp reads), Roche (454) standard and Roche (454) 4kb insert paired ends, all assembled using Newbler [99] and annotated using C. sativus ND90Pr ESTs as described below. The genome of the second C. sativus isolate, ND93-1, was sequenced at the University of Hawaii by paired end 454 runs and assembled using Newbler. S. turcica strain 28A was sequenced using Roche (454), Sanger fosmids, and shredded consensus from Velvet assembled Illumina data; EST libraries were prepared from S. turcica strains using conditions described above for C. heterostrophus. The JGI annotation pipeline was used to annotate C. heterostrophus strains C5 and C4, C. sativus ND90Pr, and S. turcica. For this, the assembled genomic scaffolds were masked using RepeatMasker [100]with the RepBase fungal library of 234 fungal repeats [101] and genomespecific libraries derived using [102]. Multiple sets of gene models were predicted for each assembly, and automated filtering based on homology and EST support was applied to produce a final non-redundant GeneCatalog representing the best gene model found at each genomic locus. The gene-prediction methods were: EST-based predictions with EST map (http://softberry.com) using raw ESTs and assembled EST contigs for each genome; homology-based predictions with 88 Fgenesh+ [103] and Genewise [104] , with homology seeded by BLASTx alignments of the GenBank non-redundant sequence database (NR: http://www.ncbi.nlm.nih.gov/BLAST/) to the genomic scaffolds; and ab initio predictions using Fgenesh [103]) and GeneMark [105]. Genewise models were extended to include 5’ start and/or 3’ stop codons when possible. Additional EST-extended sets were generated using BLAT-aligned [106] EST data to add 5’ UTRs, 3’UTRs, and CDS regions that were supported by ESTs but had been omitted by the initial prediction methods. All genome annotations can be interactively accessed through MycoCosm [107], http://jgi.doe.gov/fungi. 3. Resequencing additional C. heterostrophus race T and race O strains and other Cochliobolus spp. Because the subject genomes are all closely related to the fully sequenced reference C. heterostrophus strain C5, additional C. heterostrophus field strains Hm540, Hm338 and PR1x412, C. victoriae, C. carbonum, and C. miyabeanus, were (re)-sequenced using using Illumina technology. DNA was randomly sheared into ~200bp fragments using Covaris E210 according to the manufacturer's recommendation and the resulting fragments were used to create an Illumina library. 2x76 bp reads were assembled using Velvet (version 1.1.04), [97]. There are no ESTs available for these organisms. Assembled contigs were mapped to the reference C. heterostrophus C5 for analysis of genome variation and rearrangements. Assembled reads are called ‘nodes’ (scaffolds). Overall sequence assembly and annotation statistics are presented in Table II.3. 89 The genomes without JGI annotation pipeline gene predictions (ChHm540, ChHm338, ChPR1x412, C. carbonum, C. miyabeanus, C. victoriae) were annotated using Augustus [108]. 4. Mapping genomes to the C. heterostrophus strain C5 assembly Assembled genomes were mapped individually to the C5 reference strain using the nucmer program of MUMmer v 3.22 [46], a program that finds unique, exact matches to build whole genome alignments. SNPs were called and analyzed using the dnadiff wrapper on the filtered MUMmer delta files. Unassembled reads were also aligned to the C. heterostrophus C5 reference genome for low coverage analysis using maq-0.7.1. Regions of the reference genome under a depth of three aligned reads were considered “low coverage” for our analyses. Adjacent low coverage regions were merged if they were separated by less than 100 bp in order to minimize noise from mis-mapping of occasional low quality reads. After low coverage regions were identified for pairwise comparisons to C5, regions were identified that were shared as low coverage in multiple genomes: i.e., C. carbonum, C. victoriae, C. miyabeanus, and C. sativus for C. heterostrophus specific regions; C. heterostrophus Hm540, Hm338, and PR1x412 for C strain specific regions; and C4, Hm338, and PR1x412 for race O specific regions. Cross genome comparisons were visualized using Mauve [44], a multiple genome alignment tool that visualizes localized collinear blocks (LCB) between genomes. 5. Mapping reference genome C. heterostrophus strain C5 sequenced scaffolds onto the genetic map Cloned RFLP markers [8] were sequenced and sequences used in blast queries against the C. heterostrophus C5 assembly. Top hits were filtered using Bioperl and manually confirmed to 90 span the entire RFLP with very high stringency to rule out markers that might exist as more than one copy. Physical and genetic distances of adjacent RFLPs mapping to the same scaffold were plotted and used to calculate an average ratio of physical to genetic distance (Fig. S1, Table S2). Relative RFLP location was used to orient scaffolds along the linkage group when possible. 6. Mapping of sequenced scaffolds to the C. sativus genetic map A genetic linkage map was generated previously using the mapping population derived from a cross between parental isolates ND93-1 and ND90Pr [30] and amplified fragment length polymorphism (AFLP) and RFLP markers. To add simple sequence repeat (SSR) markers to the map, the draft sequence assembly of the C. sativus isolate ND93-1 was screened for SSR loci with di- and tri-nucleotide units tandemly repeated six or more times using a Perl script (provided by Zheng Jin Tu at the University of Minnesota, St. Paul). The SSR-containing sequences from ND93-1 were aligned to the draft genome sequence of isolate (ND90Pr) of C. sativus. Only those sequences that were polymorphic between the two C. sativus parents (ND931 and ND90Pr) were used for primer design and tested for segregation in the mapping population used previously [30]. PCR conditions and detection of SSR markers were as previously described [109]. Map construction was performed by using MAPMAKER version 2.0[110]. A minimum LOD value of 4.0 and a maximum theta of 0.3 were used to group all SSR markers with previously mapped AFLP, RFLP and PCR markers [30]. The Kosambi mapping function was used to calculate the map distance. To associate linkage groups to the sequenced scaffolds of C. sativus, the sequences of mapped SSR markers were used as queries to blast against the draft genome assembly of C. 91 sativus isolate ND90Pr and the coordinates of each SSR marker were recorded for the associated scaffold. 7. Identification and phylogenomic characterization of nonribosomal peptide synthetases and polyketide synthases NRPS and PKS proteins were identified using our custom fungal AMP domain model [57] for the former and an HMM model build from C. heterostrophus KS domains plus sequences from the C-terminal and N-terminal ketosynthase (KS) Pfam domains for the latter (PF00109 and PF02801). Proteins were identified in two ways. In the first case, genome nucleotide sequences were searched using Genewise [111] and sequences extracted and concatenated by a Perl script utilizing Bioperl’s searchIO system [112]. In the second case, Augustus protein models (see above) and JGI protein models (C. heterostrophus, C. sativus, S. turcica) were searched with the PKS KS and NRPS AMP HMM using HMMER 3.0 [113], and the sequences extracted and concatenated using a Perl script with Bioperl’s searchIO. AMP and KS domains were aligned, separately, with MAFFT (http://mafft.cbrc.jp/alignment/software/) and manually inspected to remove columns of poor alignment. ProtTest [114] was run on both alignments and identified the RTREVF model as the best fit for the AMP domains and the WAGF model as the best fit for the KS domain alignments, respectively. RAxML [115] using the RTREVF and WAGF models with a gamma distribution was used to infer maximum likelihood trees and bootstrap support was determined using the fastbootstrap method with 1000 bootstrap replicates [116]. The CIPRES portal (http://www.phylo.org/sub_sections/portal/) was used for inference of phylogenetic trees. 92 C. sativus ND90Pr NRPS AMP domains were used as blast queries to identify AMP domain orthologs in ND93-1 with the methods above. An ND93-1 AMP was considered orthologous if it was at least 95% identical to the ND90Pr query. 8. Small secreted protein identification Candidate small secreted proteins (SSP) were identified by screening the gene catalogue of each genome. Proteins smaller than 200 amino acids and containing more than 2% cysteines were searched for transmembrane domains and secretion tags using Phobius [71]. Those without transmembrane domains were retained. EST support and domain prediction for C. heterostrophus C5 SSPs was performed using the JGI portal. Cross-genome comparisons were made based on all vs. all reciprocal best hit analysis with an 80% similarity cutoff. 9. C. sativus EST library construction Three EST libraries were constructed. A mycelia-only library was constructed by harvesting mycelia grown on different media including Potato Dextrose Agar (PDA), minimal medium (MM) [117], V8PDA (150 ml V8 juice, 850 ml H2O, 10 g PDA, 10 g Agar and 3 g CaCO3), and water agar (15g agar, 1000 ml water). Mycelia were harvested after 12, 24, 48, 76 and 96 hours of growth, and time points from different media were mixed together for RNA extraction. Equal amounts of extracted RNA from each of the 5 time points was bulked to construct the mycelia only library To construct the in planta cDNA libraries, two week old barley cv. Bowman and 4 week old Brachypodium distachyon line Bd21were inoculated with conidia of ND90Pr at a concentration of 5 x 103/ml [118]. Inoculated plants were incubated in a humid chamber for 24 hours and moved to the greenhouse. Leaves were harvested at 6, 12, 24, 93 48, 72, and 96 hours after inoculation and total RNA extracted from each sample. The final in planta cDNA libraries were constructed by mixing the equal amounts of total RNA from different time points. Total RNA was isolated from all samples using the PureLink RNA Mini Kit (Invitrogen, Carlsbad, CA) and purified by treatment with DNase I (Invitrogen, Carlsbad, CA). These three libraries were sequenced by JGI. 10. Quantitative real time PCR For C. sativus, total RNA extracted as described above at six time points (6, 12, 24, 48, 72, and 96 hours) after inoculation was used for RT-PCR. The reverse transcription reaction was performed on 2µg of total RNA using the SuperScript III First-Strand Synthesis System (Invitrogen, Carlsbad, CA). cDNA was diluted 20 times and used as the template for quantitative RT-PCR, which was performed with the AB7500 real time PCR system (Applied Biosystems, Foster, CA) (Table S9). For each cDNA sample, three replications were performed. Each reaction mixture (20 µl) contained 5µl of the cDNA template, 10µl of SYBR Green PCR Master Mix (Applied Biosystems, Foster, CA) and 0.3µl of each primer (10mM). All samples were normalized using RT-Actin-F and RT-Actin-R primers as a control, and values were expressed as the change in the increase/decrease of the relative levels of the control sample (M96, which is the mixture of mycelia harvested from different media including PDA, MM, V8PDA, and water agar). For S. turcica, leaf samples with lesions were collected at five time points (3, 5, 6, 7 and 8 days) after inoculation with 9 x 104 spores per plant (three weeks old) and total RNA was extracted and qPCR done as described [119]. The actin gene was used as internal control using ATC1 primers [119]. The S. turcica gene primers corresponding to protein ID 161586 are listed 94 in Table S9. Expression level was expressed as fold change versus mycelial samples harvested on Lactose Casein Agar (LCA) plates. 11. Transformation and gene deletion Fungal transformation and molecular characterization of gene knockout mutants were conducted according to the methods of [120]. The split marker system [121] was used for gene deletion. The 5' and 3’ flanking sequences of the NPS gene encoding protein ID 115356 were amplified from ND90Pr DNA using primer pairs GTCGACTGCCATCTGGAAAC/CACTGGCCGTCGTTTTACAACGTCCACTCGACAGGTC CGTAGGT and TCATGGTCATAGCTGTTTCCTGTGGTATCCACAAAGCCACAGCA/GACGAACCAGA GATGCATGA) respectively. To verify deletion of the gene corresponding to protein ID 115356, primers CAN1F3: AGTTGTTGGGGAGTTGTTGG and CAN1-F4: TGAGCCGTTGTCATGTATCG matching the deleted portion of the gene were used. The expected PCR product was obtained from WT DNA, but not when DNA of the deletion mutant was used as template. To further confirm that the hygromycin resistance gene replaced the target gene at the native locus, PCR was conducted using a primer located outside the 3' flank used for gene deletion (CsNPS1F0: GTCCTACGGCAATTGTGGAC) and a second primer (HY: GGATGCCTCCGCTCGAAGTA) located in the hygromycin resistance gene. No amplification occurred when WT DNA was used, while the expected amplicon was observed when DNA of the mutants was used as template. 95 12. Plant inoculation For C. sativus, virulence of the mutant (ID# 115356, Fig. II.7C) and wild type strains was tested on barley cv. Bowman by spray inoculation using 2×103 conidia/ml. Inoculated plants were incubated in a humid chamber for 18-24 hours, and then transferred to a greenhouse room (20+/-2ºC). For S. turcica, three week old W64A maize plants were sprayed with 9 x 104 spores per plant, and plants grown under conditions described previously for C. heterostrophus [119]. 13. Mating type locus comparisons MAT1-1 and MAT1-2 mating type regions were identified by blasting the corresponding known C. heterostrophus MAT sequences (MAT1-1: accession CAA48465, MAT1-2: accession CAA48464) against each genome. Regions immediately 10 kb upstream and downstream were aligned pairwise to C. heterostrophus C5 (MAT1-1) or C4 (MAT1-2) MAT regions using ProgressiveMauve [44] to generate SNP data, and as a group (with and without S. turcica for MAT1-1) for visualizing the alignment. 14. Data Access Genome assemblies and annotations are available via JGI Genome Portal MycoCosm (http://jgi.doe.gov/fungi, [107] and DDBJ/EMBL/GenBank under the following accessions Cochliobolus heterostrophus ATCC 48331 (race T, strain C4): AIHU00000000, Cochliobolus heterostrophus ATCC 48332 (race O, strain C5): AIDY00000000, Cochliobolus sativus ND90Pr: AEIN00000000, Setosphaeria turcica Et28A: AIHT00000000, C. sativus ND93- 96 1:PRJNA87041, Cochliobolus carbonum 26-R-13: AMCN00000000, Cochliobolus miyabeanus ATCC 44560: AMCO00000000, Cochliobolus victoriae FI3: AMCY00000000. 15. Acknowledgments SZ and YL thank Dr. Shaukat Ali and Joe Mullins for assistance in C. sativus inoculation on barley and Drs. Justin D. Faris and Maricelis Acevedo for providing equipment and facilities for the RT-PCR experiments. 97 E. References 1. Ohm R, Feau N, Henrissat B, Schoch CL, Horwitz BA, et al. 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The Supplemental text S1 and all Supplemental Figures are reproduced in the Appendix of this dissertation. 112 ! Chapter III Genomics-driven investigation of T-toxin production in Cochliobolus heterostrophus ! 113 ! Abstract In 1970, a new and highly virulent race of Cochliobolus heterostrophus, race T, swept the US east coast, destroying more than 15% of the US maize crop. The sudden emergence of race T, and the genetic complexity of the Tox1 locus required for biosynthesis of the linear polyketide T-toxin and concomitant high virulence to maize, have made the origins of Tox1 and race T a formidable and compelling mystery. Tox1 is not found in previously known race O. The pursuit to identify the complete set of genes required for T-toxin production and to determine the evolutionary origins of Tox1 continues here, leveraging the genomic resources afforded by multiple sequenced race O and race T C. heterostrophus strains (Chapter II), and an everexpanding set of related fungal genomes. In silico subtraction of race O unique DNA from race T identified 85kb of Tox1 candidate DNA, 48kb of which corresponds to known Tox1 sequence. Aligning race O and race T strains also yielded two candidate locations in the race O genome for the reciprocal translocation breakpoint where Tox1 maps in race T. Phylogenetic analyses of all ten C. heterostrophus Tox1-affiliated proteins revealed that Didymella zeae-maydis and Leptosphaeria maculans each possess complete but slightly divergent Tox1 loci, while the Eurotiomycete Talaromyces stipitatus possesses a distant ortholog of one of two C. heterostrophus Tox1 polyketide synthases (PKS). Genes at the L. maculans Tox1 locus are clustered, in contrast to the unlinked and scattered nature of genes at the C. heterostrophus Tox1 locus. The L. maculans locus lead to the discovery of a new gene required for T-toxin biosynthesis, TOX10. In vitro T-toxin activity assays demonstrated that C. heterostrophus, D. zeae-maydis, and T. stipitatus all display T-toxin-like activity. Results with L. maculans were inconclusive. ! 114 ! A. Introduction 1. Southern Corn Leaf Blight, Yellow Leaf Blight, and Blackleg i. Cochliobolus heterostrophus and the SCLB epidemic Cochliobolus heterostrophus, the causal agent of Southern Corn Leaf Blight (SCLB) was first described as a pathogen of maize in 1925 [1]. C. heterostrophus favors warm, wet climates, and prior to the SCLB epidemic of 1970, was restricted to US southeastern states [2]. In 1970, however, a new and highly virulent race of C. heterostrophus, race T, swept the US east coast, destroying more than 15% of the US maize crop [3,4]. Texas male sterile (T-cms) corn was widely planted at the time, as its trait of male sterility allowed for easier cross fertilization for hybrid vigor [3,5]. This same trait confers susceptibility to C. heterostrophus race T, although this was unknown at the time. We now understand that race T isolates produce a novel host selective toxin (HST), T-toxin, not found in the less virulent ‘race O’ isolates that were previously known. By 1970, 85% of the hybrid maize planted in the U.S. was T-cms [5-7]. When the previously unknown race T emerged, the impact was devastating; more than 30 states reported serious damage to corn, and the estimated damage was valued, at the time, at over one billion dollars [8,9]. There is evidence that C. heterostrophus race T was present in the field prior to 1970. In the Philippines, where T-cms corn was introduced in 1957 [2,10], C. heterostrophus was noted to be highly aggressive on T-cms corn as early as 1961 [11,12]. In the U.S., severe SCLB symptoms were reported prior to the epidemic in a number of Midwestern and central states [5,13]. ! 115 ! ii. Didymella zeae-maydis Coincident with the SCLB epidemic was the emergence of a new disease, Yellow Leaf Blight caused by a fungus then called Phyllosticta maydis (Arny & R.R. Nelson) [14]. The current most commonly invoked name (including herein) is Didymella zeae-maydis [9,15]. D. zeae-maydis has also been named Mycosphaerella zeae-maydis [16], Phoma zeae-maydis [17], and most recently with the one fungus one name initiative, Peyronellaea zeae-maydis [15]. Like C. heterostrophus (and Leptosphaeria maculans, below), D. zeae-maydis is a Dothideomycete in the order Pleosporales [18]. Unlike C. heterostrophus, it had not been reported as a pathogen of corn prior to its emergence, likely dating to 1967 in Wisconsin and Ontario [19,20]. All isolates of D. zeae-maydis are highly virulent on T-cms corn: there is no race O equivalent. Like C. heterostrophus race T, this high virulence is owed to the production of PM-toxin, a suite of polyketide toxins with a chemical structure and biological activity similar to that of T-toxin, produced by all known D. zeae-maydis isolates [21]. Genetic, molecular genetic, and genomic work with D. zeae-maydis has been scant, and appears limited to an abandoned random mutant library [22], chemical characterization of PMtoxin [21], and molecular genetic endeavors to identify the PM-toxin locus by graduate student Sung-Hwan Yun in the Yoder/Turgeon laboratory in the late 1990’s [23,24]. This work is instrumental to our understanding of T-toxin and PM-toxin, and is discussed in Section A.4.iv below. Otherwise, the pathobiology of D. zeae-maydis remains unexplored. iii. Leptosphaeria maculans Leptosphaeria maculans ‘brassicae’ (Desm.) Ces. et de Not [anamorph = Phoma lingam (Tode: Fr.) Desm.] causes ‘blackleg’ or stem canker of canola (Brassica napus), and is the most damaging disease of canola worldwide [25,26]. The L. maculans designation originally included ! 116 ! the less aggressive Leptosphaeria biglobosa strains, but was split into separate species [27,28]. Like D. zeae-maydis (and thousands of other misplaced fungi [15]), L. maculans was once classified based on its anamorph as a Phoma species, a genus which has since been heavily restructured thanks to molecular phylogenetics [15]. C. heterostrophus, D. zeae-maydis, and L. maculans, therefore, are all Dothideomycetes in the order Pleosporales, each in a distinct family (the Pleosporaceae, Didymellaceae, and Leptosphaeriaceae, respectively) [15,18]. While blackleg does not directly bear on the SCLB epidemic of 1970, the genome sequence of L. maculans [25] holds the key to understanding the origins of T-toxin as described in Section C.9, and thus its biology will be discussed briefly here. L. maculans is classified as a hemibiotroph, meaning it has distinct biotrophic and necrotrophic phases of infection. The primary inoculum is ascospores, which land on canola cotyledons and leaves, invading through the stomata. Initial growth is limited to the spaces between mesophyll cells, and lesions become visible after several days [29]. The fungus then invades the xylem and grows systemically [30]. Complete death of the host plant occurs once growth strangles the stem cortex, resulting in a black canker at the base of the stem (hence the name “blackleg”). After harvest L. maculans survives on stubble as a saprophyte and undergoes sexual reproduction, producing windborne ascospores that infect canola crops sown the following year [29]. As a hemibiotroph, a gene-for-gene interaction defines the stem canker pathosystem. There are at least eleven different [29,31] named effector genes (AvrLm1-AvrLm11 [32]) which result in avirulence in the presence of a corresponding host factor [31,33], some of which (eg., AvrLm1, AvrLm2, AvrLm4-7, AvrLm6) have been characterized at the molecular level [34-36]. AvrLm1 is an interesting case study, as it activates both biotrophic (salicylic acid) and ! 117 ! necrotrophic (ethylene) defensive marker genes in late (after 5 days) infection [37]. The two pathways are thought to be antagonistic (as described in Chapter I) [38,39], and their simultaneous activation speaks to the hybrid nature of hemibiotrophs. Cloning of the AvrLm genes led to the startling discovery that they were located in topographically defined regions called isochores: long, AT-rich stretches of DNA [40]. Indeed, the entire genome of L. maculans is structured in alternating AT-rich and GC-equilibrated regions, the former housing effectors and the latter the housekeeping genes, in general [25]. AvrLm1, for example, was located as a solo gene in an AT-rich region consisting mainly of LTRretrotransposons, between small (20-30kb) GC-equilibrated regions containing housekeeping genes. AvrLm6 also occurs as a solo-gene within a 133-kb, LTR-retrotransposon rich, noncoding region [34]. What were identified genetically as ‘clusters’ were physically distant genes separated by noncoding isochores [35]. Genetic recombination is apparently suppressed at these loci, resulting in a distorted physical/genetic distance. One effector, AvrLm11, is not only located in a large (321kb), AT-rich region, but is also found on a conditionally dispensable chromosome [32]. This AT- and repeat-rich quality will be of great interest later to our understanding of Tox1 in C. heterostrophus. While there is no question that L. maculans utilizes effector proteins and invades the plant as a hemibiotroph, this disease strategy was not always well understood. Although L. maculans clearly has a long, symptomless biotrophic phase in B. napus, the end phase is necrotic, and the latter was a major research focus [41,42]. Secondary metabolites, therefore, were a major focus for understanding L. maculans, especially in relation to the less virulent L. biglobosa strains. The best characterized secondary metabolite from L. maculans is sirodesmin PL, ! 118 ! synthesized by a nonribosomal peptide synthetase (NRPS) [43]. Sirodesmin PL is a member of the epipolythiodioxopiperazine (ETP) class of fungal secondary metabolites and is characterized by a disulphide bridge across a diketopiperazine ring [41]. The Aspergillus fumigatus toxin gliotoxin is the first, and best described, ETP-type toxin [44]. Sirodesmin PL (and other ETPs) is toxic to a variety of organisms, including animals, plants, and fungi; it does not act as a hostselective toxin (HST) [41,44]. Although Sirodesmin PL is only produced by virulent L. maculans strains, and not by the less virulent L. biglobosa isolates [45], its role in pathogenicity is unclear. Purified sirodesmin causes chlorotic (yellow) lesions on plant leaves, and has antibacterial and antiviral properties [41,46]. Mutants unable to produce Sirodesmin PL form smaller stem lesions on B. napus than wild-type isolates, but they are like wild type (WT) in cotyledon assays [47]. In addition to NRPS produced secondary metabolites, L. maculans also produces polyketides. The genome contains 15 PKS genes [48], and the chemical structure of several polyketides has been described, although none of these appears to be linear, like T-toxin and PM-toxin (see below) [49]. LmPKS2, for example, is responsible for the production of Phomenoic acid, which has antibiotic and antifungal properties [48]. iv. Three pathogens C. heterostrophus and D. zeae-maydis share many commonalities: a common host and pathogenic lifestyle, a history as emergent pathogens, and even the same susceptible host genotype. L. maculans, on the other hand, has a different host plant (a dicot, not a monocot), a different pathogenic lifestyle, and no known HSTs. Despite these differences, each organism’s genomic PKS inventory has a common thread that may explain the emergence of the race T SCLB epidemic. ! 119 ! 2. Identification of T-toxin and URF13 The high virulence of C. heterostrophus race T on T-cms corn was soon attributed to production of a host-selective toxin (HST), called T-toxin. Seedling root growth assays and leaf injection tests confirmed that T-toxin caused the same symptoms on T-cytoplasm corn as the fungus itself [6,50,51]. T-toxin was identified as a mixture of linear polyketols, ranging in length from C35 to C49, although primarily (60-90%) composed of C39 and C41 [52,53]. T-toxin analogs of shorter chain lengths (C15 to C26) could be synthesized in the laboratory, but were not as toxic as the natural longer products [54]. It was later determined that chain length appears to directly affect toxicity [55]. It was further deduced that T-toxin targets the host mitochondria, specifically the 13-kDa inner mitochondrial membrane protein T-urf13 [56]. Much later, a rapid and reliable in vitro assay for T-toxin was developed which takes advantage of the observation that transgenic E. coli cells expressing the URF13 gene are sensitive to T-toxin [57]. This assay is the basis for T-toxin and T-toxin-like activity tests in this Chapter. D. zeae-maydis is also highly virulent specifically on T-cms corn, and the discovery of Ttoxin prompted researchers to screen for a molecule with the same specificity. Two years later, the polyketide PM-toxin was discovered in culture filtrates [58-61]. PM-toxin has a similar structure to T-toxin, however the average polyketide chain length is 33-35 carbons long [21], whereas T-toxin is 35-49 carbons [52,53]. Unlike C. heterostrophus, PM-toxin is produced by all known isolates of D. zeae-maydis. T-toxin’s identity, its chemical structure, and the specific host protein it targets were all in hand by the end of the 1970’s. The chemical structure of PM-toxin was also solved, and the ! 120 ! molecule shown to have the same specificity. Understanding the genetic basis for T-toxin production, on the other hand, has proven to be a much greater challenge. ! 3. The Tox1 locus: classical genetics i. Odd ratios Despite the initial swift progress in identifying the HSTs and their corresponding host factor, the underlying genetics of the C. heterostrophus T-toxin locus, Tox1, remains a complex challenge. Initial characterization of progeny of a cross between C. heterostrophus race O and race T was consistent with a monogenic trait [6,7]. Ability to produce T-toxin, then, was thought to be the product of a single gene [8]. Indeed, randomly collected progeny of a cross between race O and race T parents showed that 42/82 (indicating a 1:1 ratio of race T to race O) were race T in their virulence and ability to produce T-toxin [10]. In this same study, however, only 16/120 (1:7 ratio race T:race O) progeny were race T when paired with a different race O parent. This 1:7 ratio was later attributed to the presence of a spore killer gene in 50% of race O field isolates [11]. It was not until 1988 that the defining genetic character of the Tox1 locus was inferred. Crosses between TOX+ (Tox1+) and TOX- (Tox1-) isolates showed a high frequency of nonrandom ascospore abortion. The Tox1 segregation pattern observed suggested that race T and race O differed by a chromosomal rearrangement, such as a reciprocal translocation [13]. This hypothesis was later directly supported by the RFLP map [62] (see Appendix). The Tox1 locus is genetically linked to the both breakpoints of hybrid chromosomes 6;12, and 12;6 in race T, which are the counterparts of a reciprocal translocation between race O ! 121 ! chromosomes 6 and 12 (Fig. III.1). Progeny of a cross between race T and race O strains that did not receive a functional pair (either the race T or the race O pair) of chromosomes were not viable, masking the true genetics of the Tox1 locus. 6 12 12;6 6;12 12;6 12 OT Tox1A Tox1B Tox1A TO Tox1 OT Tox1B race O race T OT 6 6;12 Figure III.1. Tox1 is associated with a reciprocal translocation. Chromosomes 6 and 12 of race O are hypothesized to have undergone a reciprocal translocation which created a hybrid pair in race T, that also carry DNA (black blobs) not found in race O (left). Tox1 consists of two loci in race T, Tox1A on chromosome 12;6 and Tox1B on chromosome 6;12. These loci are map genetically to the breakpoints (arrows) of the reciprocal translocation (right). We have since understood that Tox1 is actually two separate loci, Tox1A and Tox1B, each mapping genetically to the breakpoint on hybrid chromosomes 6;12, and 12;6 in race T. Support for this hypothesis could not be obtained until Tox+ strains were rendered Tox- by mutagenesis [63]. Different mutants in an originally race T genetic background could be crossed ! 122 ! together:some crosses yielded 25% TOX+ progeny, which indicated the two unlinked mutations [9,64]. When electrophoretic karyotypes of WT and tox1 mutants were compared, it was confirmed that mutants mapped to two loci on two separate chromosomes [64]. ii. Insertion of DNA in race T Electrophoretic karyotyping and the RFLP map provided more information regarding Tox1 than simply its location. They also helped to characterize the locus as 1.2Mb of highly repetitive, AT-rich DNA. While constructing the RFLP map, Tzeng at al. noted that 50% of the RFLP probes mapping within 4 cM Tox1 were highly repetitive, as compared to only 4% in the remainder of the genome [62,64,65]. Electrophoretic karyotyping demonstrated that chromosomes 6;12 and 12;6 did not sum to the size of race O chromosomes 6 and 12. Instead, the race T chromosomes 6;12 and 12;6 summed to about 1.2Mb more DNA than their race O counterparts [62]. The strains in this analysis were near-isogenic lines, backcrossed 6-12 times and differing only at Tox1: the additional DNA, therefore, was predicted to be located at or near Tox1 [62,65,66]. Cloning of genes from the Tox1 loci confirmed that they are associated with highly repeated AT-rich sequence. Tox1 genes, when identified were found on small scaffolds, which could not be linked or assembled together. The Tox1 locus, therefore, consists of 1.2Mb of DNA comprised largely of repetitive, AT-rich sequence, with small islands of coding sequence. None of the coding sequence is found in race O [67-70]. Note that this description is startlingly similar to the fraction of the genome of L. maculans which consists of AT-rich, repetitive sequence studded with isolated effectors mentioned earlier [25]. ! 123 ! 4. Molecular Genetics: Genes at Tox1 i. PKS1 The first C. heterostrophus Tox1 gene cloned was PKS1, which was recovered from the Restriction Enzyme Mediated Integration (REMI)-tagged Tox- mutant R.C4.350L [63] using the plasmid rescue procedure. PKS1 maps genetically to Tox1A, on chromosome 12;6 [64,68]. PKS1 was found to be a 7.6kb ORF with four introns. The protein has six enzymatic domains: a β-ketoacyl synthase (KS), acyltransferase (AT), dehydratase (DH), enoyl reductase (ER), βketoacyl reductase (KR), and acyl carrier protein (ACP), plus a degenerate methyl transferase (MeT) domain. PKS1 does not have a chain-terminating thioesterase (TE) domain [68]. It is important to note that PKS1 is not found in race O or any other Cochliobolus species, and that the DNA on both flanks is noncoding and AT rich (~70%) [9]. ii. DEC1 and RED1 The first Tox1B genes cloned were DEC1 and RED1. The genes were recovered from the race T strain C4.PKS.13, a plasmid tagged deletion strain deficient in T-toxin production [67]. DEC1, encoding a decarboxylase, and RED1, encoding a reductase, are adjacent in the C. heterostrophus genome and divergently transcribed. They were confirmed to be unique to race T by DNA gel blot analysis, and like PKS1 are flanked by AT rich (72%), highly repetitive, noncoding DNA [67]. Targeted disruption of DEC1, but not RED1, stopped production of Ttoxin [67]. Deletion of RED1 reduced the amount of T-toxin produced, but toxin is not eliminated unless the red1 deletion is combined with similarly “leaky” red2 or red3 mutants, discovered later [70], as described below. ! 124 ! iii. Was T-toxin the sum of these parts? All three of these genes, PKS1 at Tox1A and DEC1/RED1 at Tox1B, are involved in Ttoxin production (although only pks1 and dec1 mutants are totally tox- [67,68]), and it was hypothesized that perhaps these three genes were sufficient for T-toxin production. To investigate this, PKS1 was introduced into the tox- REMI mutant R.C4.186, which carries a 700kb deletion at Tox1A [64], but transformants remained tox- [71]. Likewise, DEC1 introduced into the tox- mutant C4.PKS.13, which carries a 100kb deletion at Tox1B [64,67], remained Tox[71]. It was thus inferred that more genes at both Tox1A and Tox1B were required for T-toxin biosynthesis (waiting to be discovered in the genomics era). iv. D. zeae-maydis PKS1 Once C. heterostrophus PKS1 was identified, a molecular investigation of D. zeaemaydis and PM-toxin was initiated. The structural similarity of PM-toxin and T-toxin suggested a PCR-based strategy to clone PKS genes from D. zeae-maydis using the KS domain (the most highly conserved domain of a PKS) of C. heterostrophus PKS1 (hereafter ChPKS1) might be productive. One PKS identified in this manner, DzmPKS1, had 82% identity to the KS domain of ChPKS1. Using the KS domain as a starting point, the entire DzmPKS1 gene and its flanking sequence was cloned by a combination of TAIL-PCR [38] and plasmid rescue [68]. The DzmPKS1 protein has 60% identity to ChPKS1 over its entire length, with identity in some functional domain signature motifs as high as 90% [9]. DzmPKS1 and ChPKS1 have identical domain organization, transcript size, (7.6kb after intron splicing), and near identical intron structure, with three out of the four occurring in conserved positions [23]. Attempts to identify more D. zeae-maydis genes required for PM-toxin synthesis resulted in a set of six ToxREMI mutants. As D. zeae-maydis is homothallic, segregation analysis could not be done, ! 125 ! although F1 and F2 progeny were collected to ensure transformants were stable [24]. In contrast to ChPKS1, DzmPKS1 is not flanked by AT rich repeated DNA, but rather by regularly spaced ORFs, including likely transposases. Two of these ORFs, designated DzmRED1 and DzmRED2, encode reductases (nomenclature does not correspond to C. heterostrophus). When any one of DzmRED1, DzmRED2, or DzmPKS1 was deleted, PM-toxin production was lost [23]. All three of these genes, therefore, are required for PM-toxin biosynthesis, although they are not necessarily the only genes involved. 5. Genomics i. PKS2 C. heterostrophus race T strain C4 was first sequenced by the Turgeon/Yoder program at Torrey Mesa Research Institute (TMRI) in 2001. 2x paired-end shotgun sequence coverage was combined with 3x Celera paired end coverage to assemble a 300 scaffold, ~35Mb assembly. In addition, several thousand ESTs were sequenced to aid in annotation. The complexity of Tox1 was confirmed, as all contigs carrying known Tox1 genes (at the time, PKS1, DEC1, and RED1) were located on the smallest (23-46kb) scaffolds in the assembly. Each of these genes was embedded in highly repetitive, AT-rich DNA [9]. Because previous attempts had determined that known Tox1 genes (PKS1, DEC1, RED1) could not be identified in several dozen plasmid, lambda, BAC, and YAC libraries [9], a cDNA subtraction approach was attempted to identify additional Tox1 genes [68,69]. 47 race T unique (expressed in C. heterostrophus race T strain C4, and not its inbred race O sibling C5) cDNAs were sequenced, and 12 were similar to PKSs, 3/12 of which corresponded to PKS1 [68]. The ! 126 ! other nine belonged to a second PKS (PKS2) which mapped to Tox1A and was confirmed race T specific by PCR [69]. Like PKS1, PKS2 is required for T-toxin production [69]. ii. Tox1 associated genome scaffolds The known Tox1 genes, as of the TMRI sequence, included PKS1, PKS2, DEC1, and RED1. These genes mapped to three scaffolds: 4FP (PKS1), 4LU (PKS2), and 3PL (DEC1 and RED1) (Fig. III.2, TMRI designations). These scaffolds were small, highly repetitive, and ATrich [9]. No additional ORFs were found on scaffolds 4FP or 3PL. In the case of 4LU, however, an ORF was found upstream of PKS2. The ORF, designated LAM1, is highly similar to a 3-hydroxyacyl-CoA dehydrogenase, contains two introns, and is 42.47% GC [46]. Scaffold 3PL, containing DEC1, did not originally hold RED1, although the two genes were known to be neighbors from previous sequencing efforts- RED1 was deduced to reside in a gap between two of the contigs composing 3PL. When the gaps were sequenced, two additional reductases, RED2 and RED3, were found and placed on scaffold 3PL. RED2 is 1133 nucleotides with two putative introns, 44.48% G+C. RED3 is 807 nucleotides, with one putative intron, 41.57% G+C [9,70]. Finally, a thorough reinvestigation of the race O/race T cDNA subtraction [69] revealed two additional race T-specific genes: the short-chain dehydrogenase OXI1, and TOX9, a gene with no predicted domains [70]. These genes, on a single 5,129bp contig (OXI1/TOX9), are also flanked by AT-rich DNA, and map to Tox1A [70]. In all, nine genes were identified as belonging to the Tox1 locus prior to the work reported here. Two polyketide synthases, PKS1 and PKS2, the decarboxylase DEC1, the 3hydroxyacyl-CoA dehydrogenase LAM1, the short-chain dehydrogenase OXI1, the three reductases, RED1, RED2, and RED3, and the hypothetical protein TOX9 [70]. These genes ! 127 ! reside on small and AT rich scaffolds, measuring 5,129 to 19,137 bp in length. The Tox1 ORFs themselves are not AT rich, ranging in AT content from 40 to 50%[70]. Of the nine Tox1 genes, PKS1 [68], PKS2 [69], DEC1 [67], LAM1 [46], and TOX9 [70] have all been confirmed as essential for T-toxin production by mutation analysis. red1, red2, red3, and oxi1 mutants are all partially, but not completely, reduced in their ability to produce Ttoxin, and only when combinatorial mutants are generated can T-toxin production be completely halted [70]. iii. PKS1 and PKS2 The domain composition and organization of ChPks1 and ChPks2 is worth narrating. ChPKS1 splices out four introns, resulting in a 2528 amino acid protein. ChPks1 has six complete enzymatic domains (KS, AT, DH, ER, KR, and ACP) and a degenerate MeT [68]. ChPKS1 lacks a chain terminating thioesterase domain at its C-terminus. ChPks2 also possesses KS, AT, DH, ER, KR, and ACP domains, but does not contain a degenerate MeT domain, accounting for the comparatively smaller 2144 amino acid protein. ChPks1 and ChPks2 are 32% identical and 50% similar at the amino acid level in regions of alignment [9]. 6. Phylogenetics of ChPks1 and ChPks2 When a phylogenetic tree of the KS domains of ChPks1 and ChPks2, and those of all known fungal PKS at the time (2005) was constructed, ChPks1 and ChPks2 clustered together in a single large clade [69]. ChPks1 and DzmPks1 clustered together on the outskirts of the clade, near ChPks7. ChPks2, on the other hand, had no closely related ortholog, although it did form a moderately supported (80 bootstrap values) clade with a PKS in Neurospora crassa [69]. Other members of the clade included LovF and MclB, involved in diketide production by Aspergillus ! 128 ! terreus and Penicillium citrinum, producing lovastatin and compactin/citrinin, respectively. No other dothideomycete orthologs appeared in this clade. A later phylogenetic analysis found additional, yet distant orthologs for ChPks1: one in the dung fungus Delitschia winteri, and one in the Penicillium-related saprophyte Talaromyces stipitatus. T. stipitatus is a fungus of great importance for this chapter, yet almost nothing is known about its biology. It can be commonly found in soil, dung, or decaying plant matter, but is generally studied in an industrial setting for its production of hydrolytic enzymes [72,73]. ChPks2 remained mysterious, without close orthologs in any fungal species [70]. Based on these trees, the phylogenetic origins of ChPKS1 and ChPKS2 were far from clear. Horizontal gene transfer, a not unreasonable hypothesis invoked at one time by the Turgeon/Yoder group [68], was thought to be no more likely than rapid loss and duplication, given the tumultuous nature of secondary metabolite genes [74]. This chapter summarizes the progress made utilizing the genomic resources available as of this writing. While our total inventory of Tox1 genes has not greatly increased (only one new T-toxin related gene, TOX10, was discovered), our general understanding of Tox1 has. Instrumental to this has been the discovery of a complete ‘rosetta stone’ Tox1-type locus in L. maculans. This provided us with evidence for the complete coding inventory of Tox1, as well as the location of the race T unique boundary in the genome, which were elucidated as described below. ! 129 ! B. Materials and Methods 1. Strains, media, and culture conditions Minimal medium (MM) contained 10 ml of each of two 100x salts solutions (Solutions A and B), 10ml of Srb’s micronutrient stock solution [75], 10 g glucose, 20g agar (for solid plates), and deionized water to 1 litre [76]. Stock salt solution A contained 10g Ca(NO3)2 !4H2O and deionized water to 100ml. Stock salt solution B contained 2.0g KH2PO4, 2.5g MgSO4!7H2O, 1.5g NaCl and deionized water to 100 ml (pH 5.3). Complete medium (CM) was MM plus 1 g yeast extract, 0.5 g acid-hydrolysed casein and 0.5 g enzymically hydrolysed casein, per liter. Complete medium no salts (CMNS), used for Hygromycin B selection, is CM without salt solution A or B. C. heterostrophus WT race T strain C4 (Tox1+;MAT1-2, American Type Culture Collection [ATCC] number 48331) was used for all initial transformations and strain C2 (Tox1+;MAT1-1;alb1, ATCC 48329) was used as an albino tester for initial crosses. All strains used in this study, including transformants, are listed in Table III.1. Strains were stored at -80oC in liquid CM [76] containing 25% glycerol and were plated onto CM with xylose instead of glucose (CMX) [70] for growth and optimal conidiation. Fungi were grown at 24 °C under fluorescent lights in an alternating 16 hours light 8 hours dark cycle under fluorescent light (Watt-Miser F34 WW/RS/WM, Warm White, General Electric). ! 130 ! Table III.1. Strains used in this chapter Strain name C4 (ATCC 48331) C5 (ATCC 48332) tox10-3 (toxHP, JGI ID# 155299) Species C. het C. het C. het Genotype Tox1;MAT1-2 tox1;MAT1-1 tox10;MAT1-2; hygBR tox10-4 (toxHP, JGI ID# 155299) abc18-1 (JGI ID# 49682) abc18-2 (JGI ID# 49682) abc18-4 (JGI ID# 49682) abc18-1 (JGI C5 ID# 109542) PR1X412 C. het C. het C. het C. het C. het C. het tox10;MAT1-2; hygBR abc18;MAT1-2: hygBR abc18;MAT1-2; hygBR abc18;MAT1-2; hygBR abc18;MAT1-1; hygBR Tox1+;MAT1-1 Hm338 (ATCC 48317) Hm540 20239 A3 20239 A11 Aus 19679-2 PM24 PM3018 J154 IBCN18 Lm893 Lm838 Lm100-3 PI85 C. het Tox1+;MAT1-2 C. het Tox1-;MAT1-1 C. het C. het C. het D. z-m D. z-m L. big L. mac L. mac Tox1-;MAT1-1 Tox1-;MAT1-2 Tox1+; MAT1-2 DzmTox1+;MAT1-1; MAT1-2 DzmTox1+;MAT1-1; MAT1-2 L. mac L. mac L. mac Comment/Reference race T, inbred, [76] race O, inbred, [76] race T, tox- mutant this work, strain C4 background race T, tox- mutant this work, strain C4 background race T, tox- mutant this work, strain C4 background race T, tox- mutant this work, strain C4 background race T, tox- mutant this work, strain C4 background race O this work, strain C5 background race T, a progeny of a cross between PR1C from Poza Rica, Mexico and strain 412, unknown geographical origin, Turgeon lab collection race T, New York, Turgeon lab collection race O, North Carolina, Turgeon lab collection race O, Australia, Turgeon lab collection race O, Australia, Turgeon lab collection race T, Australia, Turgeon lab collection Tox+, Aurora, New York, Turgeon lab collection Tox+, unknown geographical origin, Turgeon lab collection B. Howlett Univ. of Melbourne B. Howlett Univ. of Melbourne Dept. of Plant Pathology & Plant-Microbe Biology (PPP-MB) collection PPP-MB lab collection PPP-MB lab collection PPP-MB lab collection Lm421-3 L. mac PPP-MB lab collection Lm847 L. mac PPP-MB lab collection Lm81 L. mac PPP-MB lab collection Lm846 L. mac PPP-MB lab collection Lm855 L. mac PPP-MB lab collection TS1006 T. stip Louisiana, ARS culture collection (NRRL (ATCC 10500) 1006) JGI ID#s are from strain C. heterostrophus C4v1.0 unless otherwise noted. ! 131 ! 2. DNA manipulations and fungal transformations Fungal genomic DNA was prepared using the Ultraclean Microbial DNA isolation kit (MO BIO). PCR reactions were carried out with Phusion High-Fidelity DNA Polymerase and mastermix (Finnzyme) for generating transformation constructs, or GoTaq DNA Polymerase and mastermix (Promega) for screening mutants, following the manufacturers’ recommendations. Transformations were performed using protoplasting and split marker-based PCR fragments which favor homologous integration [77]. For the latter, 700-1000bp DNA flanking (5’ and 3’ flanks) coding sequence of interest was amplified using primer pairs with extensions complementary to M13Rhyg (TCCTGTGTGAAATTGTTATCCGCT-XXXXXXX on the reverse primer of the 5’ flanking region) and M13Fhyg (GTCGTGACTGGGAAAACCCTGGCG-xxxxxxxx on the forward primer of the 3’ flanking region) (see Fig. S2 in Appendix). 5’ and 3’ flanks were then amplified, along with overlapping segments of the Hygromycin gene (5’ region “HY” amplified with M13RHYG and NLC37, 3’ region “YG” amplified with M13FHYG and NLC38) from the vector pUCATPH, which contains the HygB gene between the Aspergillus nidulans PtrpC promoter and TtrpC terminator. A second round of PCR, using the 5’ flank forward primer with NLC37, and the 3’flank reverse primer with NLC38, results in the 5’ flank being fused to the “HY” hygromycin B construct, and the 3’ flank fused to the “YG” component. All primers are listed in Table III.2. To prepare protoplasts for transformation, 15x100 mm Petri dishes with 10 day old strain C4 cultures on CMX wer scraped using sterile wooden dowels, filtered through sterile cheesecloth, and inoculated into 2x 100mL liquid CM cultures overnight (18h) at 24° C. Two 40ml centrifuge tubes were filled with liquid culture and pelleted in a Sorvall RC-5C centrifuge (SS34 rotor) at 5000 rpm for 5 minutes at 4°. One scoop (10mm diameter glob) each of pelleted ! 132 ! mycelium was transferred to eight 50ml sterile flasks, each containing 10mL enzyme osmoticum [3.27 g NaCl (0.7 M), 1.6 mL Glucanex (we have been using a liquid preparation, kindly provided by C. M. Hjort, Novo Nordisk, Bagsvaerd, Denmark), 0.8 g Driselase (Sigma), H2O to 80 mL, filter sterilized] and shaken gently (40rpm) 30° for 3 hours to release protoplasts. Protoplasts were then filtered through sterile cheesecloth and nylon membrane (SEFAR, 25um pore size) and pelleted by centrifugation as above. The pelleted protoplasts were suspended and re-pelleted four times: the first time in 2x 5ml 0.7M NaCl solution, and then three additional times in 10ml STC [sorbitol, 21.86 g (1.2 M), 1 mL of 1 M Tris–HCl pH 7.5, (10 mM), 0.735 g CaCl2·2H2O (50 mM), H2O to 100 mL] solution. After washing, protoplasts were resuspended in 200uL STC, adjusted to approximately 1*107/ml and kept on ice. Protoplasts were transformed by adding 20ul transformation construct (DNA concentration not determined) to 100uL of protoplasts suspended in STC, and held on ice 5 minutes. 200uL, 200uL, and 800uL aliquots of polyethylene glycol solution [30 g polyethylene glycol, MW 3,350 (60% w/v), 0.5 mL of 1 M Tris–HCl pH 7.5 (10 mM), 0.37 g CaCl2·2H2O (50 mM), H2O to 50 mL] were added, mixing gently and incubating 5 minutes between aliquots. Finally, 1mL STC was added, and protoplasts were dispensed 300ul at a time into 20ml molten recovery medium in Petri dishes and incubated overnight at 30° C. Molten recovery medium was prepared in three flasks, A, B, and C, autoclaved separately, mixed, and held in a water bath prior to use. Flask A contained 1 g yeast extract, 1 g casein hydrolysate (enzymatic), H2O to 50 mL, flask B: 342 g sucrose, H2O to 500 mL and flask C: 16 g agar, H2O to 450 mL. 150ug hygromycin B/mL in 1% agar was overlayed the following day and transformants were allowed one week to emerge at 30° in the dark. Candidate transformants recovered from the hygromycin B overlay were transferred to ! 133 ! selective medium (CMNS containing 50ug Hygromycin B/ml) to confirm resistance. Conidia (asexual spores) were streaked on 5% water agar to separate individual conidia, and individual germinating conidia were transferred to CMX. Only a single nucleus enters a given conidium: single conidiation therefore eleminates heterokaryons. Candidates were then plated onto CMNS HygB (50ug Hygromycin B/ml) and CMX to confirm resistance and for storage in glycerol, respectively. DNA was prepared from candidate transformants and subjected to diagnostic PCR (see Appendix) to verify gene deletion. DNA from candidates with a gene of interest deleted and replaced with the selectable marker carried a band of predictable size when a primer external to the 5’ or 3’ flanking region was combined with NLC37 and NLC38 (internal to HygB), respectively, and if a band was not amplified using primer pairs internal to the gene targeted for knockout (see Fig. S2 in Appendix). All gene deletion and verification primers are listed in Table III.2. ABC18 (JGI C4 ID #49682) deletion constructs were generated using the primer pair 49682upf/49682upr for upstream and 49682dnf/49682dnr for downstream flanking regions. Transformation and gene deletion were performed using both C. heterostrophus race T strain C4 and race T strain C5. Successful deletion was confirmed using the external flanking primer pairs 49682upfext/NLC38, 49682dnrext/NLC37 for integration into the upstream and downstream flanks, respectively, and the internal primer pairs 49682inf/49682inr to verify deletion of ABC18. TOX10 (TOXHP: JGI C4 ID# 155299) was disrupted, rather than completely removed, due to the limited, and AT-rich, nature of its flanking sequence. The disruption construct was synthesized using the primer pair 155299_4upf/155299_4upr for upstream and 155299_4dnf and 155299_4dnr for downstream flanking regions. Correct integration and deletion was confirmed ! 134 ! using the external flanking/internal HygB primer pairs 155299_V2upF/NLC38 (upstream) and 155299_5dnR/NLC37 (downstream). ! 135 ! Table III.2. Primers Primer Sequence (5->3’) Purpose (see Fig. S2) 155299_4upf TGATCTTCAGCGTCACATTT Upstream forward, for disruption of TOX10 (ID: 155299) 155299_4upr TCCTGTGTGAAATTGTTATCCGCTT Upstream reverse, for disruption of AATGCTCTAGGTGGACACG TOX10 (ID: 155299) 155299_4dnf GTCGTGACTGGGAAAACCCTGGCA Downstream forward, for disruption of TCAGTTTCTCACGGCAGTC TOX10 (ID: 155299) 155299_4dnr AGCCTTAGCCCTAAAGAGTCA Downstream reverse, for disruption of TOX10 (ID: 155299) 155299 V2 upf TTTAACGGGCACATCCGTA Upstream external, for verification of TOX10 deletion with NLC38 (ID: 155299) 155299_5dnr GCCCGTGAAATAAAGATGTG Downstream external, for verification of TOX10 deletion with NLC37 (ID: 155299) 49682_upf CCCGATTGGGATAGAAAAC Upstream forward, for knockout for ABC18 (ID:49682) 49682_upr TCCTGTGTGAAATTGTTATCCGCTC Upstream reverse, for deletion for GATGTAGTGCACAGCTCA ABC18 (ID:49682) 49682_dnf GTCGTGACTGGGAAAACCCTGGCT Downstream forward, for deletion for ACGGGAACATGGTTTCAC ABC18 (ID:49682) 49682_dnr CCCTCGACCTTCTTCAACT Downstream reverse, for deletion for ABC18 (ID:49682) 49682_upfext TGGTTCATAGCGTTGTTGTC Upstream external, for verification of ABC18 deletion with NLC38 (ID:49682) 49682_dnrext AATTCTTCCACGTCCCATC Downstream external, for verification of ABC18 deletion with NLC37 (ID:49682) 49682_inf TCGATCAACTCGCTCTTGT Internal to ABC18 (ID:49682) for deletion verification 49682_inr CATAGAGCACGGCTGAAAC Internal to ABC18 (ID:49682) for NLC37 GGATGCCTCCGCTCGAAGTA deletion verification pUCAPTH* SEQ. 1685-1702 NLC38 CGTTGCAAGACCTGCCTGAA pUCATPH SEQ. 2132-2150RC M13RHYG AGCGGATAACAATTTCACACAGGA pUCATPH SEQ. 2865-2888RC M13FHYG CGCCAGGGTTTTCCCAGTCACGAC pUCATPH SEQ. 352-375 LbreakF1 agtggggaacacgaaagat Upstream of left break on C5v3 scaffold 12 (chromosome 12). LbreakR1 ccgcttgcaaaatgaatag Downstream of left break on C5v3 (chromosome 12) LbreakF2 atatgtttcccagcccaat Alternate upstream of left break on C5v3 scaffold 12 (chromosome 12). LbreakR2 gatggtatccgcagaaatg Downstream of left break on C5v3 (chromosome 12) RbreakF1 cgatacatgcaggtccact Upstream of right break on C5v3 scaffold 12 (chromosome 12). RbreakR1 atattgcggacgagacaga Upstream of right break on C5v3 scaffold 12 (chromosome 12). RbreakR2 accgattttcgaggagaag Alternate upstream of right break on C5v3 scaffold 12 (chromosome 12). RbreakF2 gtgcaaatgagcgagtagc Alternate upstream of left break on C5v3 scaffold 12 (chromosome 12). ID numbers cited are the JGI C. heterostrophus C4 protein IDs; * see [63,77] ! 136 ! 3. Genomes utilized Genomes and protein catalogs of C. heterostrophus strains C4 and C5 and L. maculans were acquired from the JGI Mycocosm website [78]. The genome sequence and gene catalog of the additional C. heterostrophus field strains Hm540, Hm338, and PR1X412 [79] were hosted and queried locally. The D. zeae-maydis strain PM3018 (F3018) was sequenced using Illumina technology and assembled with the CLC next generation sequence cell (v3.2) assembler by Dr. Sung Hwan Yun (Soonchunhyung University, Korea) and annotated and queried locally. Annotation of D. zeae-maydis was done de novo using the Hidden Markov Model-based annotater Augustus [80], as reported previously [79]. The T. stipitatus genome was originally sequenced by the Craig Venter Institute, and its sequence and annotations were downloaded from the JGI integrated Microbial Genomes database [81]. The gene catalogs from all additional genomes utilized were downloaded from the JGI Mycocosm [78]. 4. Identification of Tox1 scaffolds and genes The known C. heterostrophus Tox1 proteins (Pks1, Pks2, Dec1, Lam1, Oxi1, Tox9, Red1, Red2, Red3) were retrieved from Genbank and used as blastp queries against the protein catalog of JGI-hosted L. maculans, T. stipitatus, and C. heterostrophus C4 genomes. For unhosted genomes (C. heterostrophus C4 JGI pre-release, C. heterostrophus C4 TMRI, C. heterostrophus PR1X412, C. heterostrophus C4 Hm338, D. zeae-maydis PM3018), Tox1 proteins were queried against the assembled nucleotide sequence using blastx. In parallel, ab initio gene catalogs were created for each genome using Augustus [80], which were in turn queried with the known Tox1 proteins using blastp. Scaffolds matching Tox1 queries were checked for additional Augustus gene calls. ! 137 ! i. In silico subtraction Panseq’s novel region finder [82] was used to find scaffolds in C. heterostrophus strain C4 (query) absent in strain C5 (reference). The MUMmer [83] alignment criteria used were b=200, c=50, d=0.12, g=100, and l=20. The minimum novel region length was set to 500bp. Panseq subtractions using all race T genomes (C4 pre-release, Hm338, PR1X412) as the query used the same alignment criteria, but with a novel minimum region size of 50bp. For manual characterization of candidate race T unique sequence, each C4 scaffold was visually inspected in the JGI comparative synteny browser, aligned to C5 [78]. VISTA alignments were considered for C. heterostrophus C4 to C. heterostrophus C5, L. maculans, and C. sativus. For this analysis, a given scaffold was confirmed as race T unique if it did not have significant alignment to any of these genomes. The whole-genome alignment of C4, C5, Hm540, Hm338, and PR1X412 was inspected in the alignment software MAUVE [84]. This alignment was performed with the pre-release C4 assembly, and each scaffold was assigned to a final release scaffold based on a whole-genome blast alignment. Pre-release scaffold designations are included in Table III.3. A candidate scaffold was considered a Tox1 candidate if it did not have any Locally Colinear Blocks (LCBs) matching to either race O strain (C5, Hm540). Strains with LCBs only in one or more race T strain were still considered candidate Tox1 sequence, as long as they did not match any race O sequence. Each scaffold identified by Panseq as a candidate Tox1 sequence was manually inspected for %AT, repeats, transposons, gene calls, and functional annotations in the C. heterostrophus C4 v1.0 JGI browser [78]. ! 138 ! 5. Tox1 breakpoint identification The MAUVE whole genome alignment was manually scanned to identify regions present in race O strains, but not in race T [84]. Primers spanning the left and right gaps of scaffold 12 (pairs LbreakF1/LbreakR1 and LbreakF2/LbreakR2 for the left, RbreakF1/RbreakR1 and RbreakF2/RbreakR2 for the right) were used to amplify DNA across the proposed gap on C5 scaffold 12 in DNA isolated from race O strains C5 and Hm540, and race T strains C4, Hm338, and PR1X412. The primer pair bk24up1/bk24dn1 was used to amplify DNA by PCR across the gap of scaffold 24. 6. Phylogenetics of Tox1 genes i. Ketosynthase domain sequence extraction All gene catalogs were retrieved from the JGI mycocosm [78], except for those of D. zeae-maydis and T. stipitatus. The draft D. zeae-maydis assembly was annotated de novo by us as noted above. T. stipitatus annotation was retrieved from the JGI integrated microbial genomes viewer [81]. Additional individual protein sequences (bacterial FabF, Delitschia winteri pks1, and the top 20 blast hits for ChPks1 and ChPks2) were retrieved from Genbank. A complete list of species used for tree construction is provided in Table III.4. Ketosynthase (KS) domains were identified in gene models using HMMer3 [85] by searching with our KS HMM model built from the C- and N- terminal domains (PF00109 and PF02801) [79]. Domains were recovered from alignments using Readseq [86] and a custom perl script [79]. ii. Tox1 affiliated protein sequence extraction To search for homologs of Tox1 proteins other than the PKSs, Dec1, Lam1, Oxi1, Tox9, ! 139 ! Tox10, Red1, Red2, and Red3 were used as queries against the NCBI nr protein database to retrieve the top 50 proteins for each query in other species [87]. D. zeae-maydis candidates were identified separately with the standalone blast+ program blastp, and added to the final sequence set [88]. iii. Tree construction All sequences were aligned with MAFFT using default parameters [89] and KS domain alignments were manually annotated to remove columns of poor alignment. Gblocks was used for all other protein alignments with the most relaxed options (allowing for smaller gaps, gap positions in final blocks, and less strict flanking positions) to remove poorly aligned positions [90]. Protein substitution models were evaluated with ProtTest3 [91] and the best model chosen for each alignment (Dec1: LG+I+G, Lam1: LG+I+G, Oxi1: LG+I+G+F, Red1/Red2/Red3: LG+G, Tox9: LG+I+G, Tox10: CpREV+G) and the maximum likelihood tree was inferred with RaxML [92] through the CIPRES phylogenetic gateway [93]. Phylogenetic trees were visualized and processed in Dendroscope [94]. A bootstrap value of 80 was used as a minimum cutoff for “good support” in describing clades. 7. T-Toxin activity assessment T-toxin activity was assayed using E. coli carrying the maize T-URF13 gene on the plasmid pATH13-T [57]. E. coli with pATH13-T, as well as E. coli with pUCATPH lacking the T-URF13 gene (negative control), were plated from glycerol on Luria broth plus ampicillin (100ug/ml) agar (LBA), then a single colony was restreaked on LBA and grown at overnight. pATH13-T colonies were grown at 30°C, and pUCATPH at 37°C. A generous (2mm diameter) sample of this culture was used to inoculate 100ml liquid core medium (M9 salts, 1M MgSO4, ! 140 ! 1M CaCl2, 0.5% casamino acids) with 20mg tryptophan/liter and 100mg ampicillin/liter and shaken overnight. Cultures were diluted 1:10 in liquid core medium without tryptophan, shaken for 1 hour, and activated with 0.5ml 1mg/ml 3-indoleacrylic acid (Sigma). These cultures were shaken for two more hours and stored at 4 ° if not used immediately for up to two weeks. 5ml of induced pATH13-T (or pUCATPH) carrying E. coli culture was inoculated onto 150x15mm LBA dishes and shaken at 30° for one hour. Liquid culture was then poured off and plates allowed to air dry. Agar plugs of fungal cultures were removed from the growing tip of the colony with a cork borer and placed mycelium side down on these plates. For abc18 mutants, plugs were collected with a cork borer at the leading edge of growth, 5mm past the edge, and 5mm behind the leading edge of growth. 8. Comparison of PKS sequence characteristics PKS proteins were blasted against each other either on the NCBI blastp webserver or locally using the standalone blast+ blastp to determine % identity [88]. Predicted transcripts for each gene were downloaded from the JGI Mycocosm, and %AT was calculated using a simple perl script. For D. zeae-maydis, where no transcript catalog was available, %AT was only calculated for PKS1 and PKS2. 9. Maize inoculations and virulence assays W64A-T and W64A-N corn was grown in a growth chamber under 16 h light/ 8 h dark at 24 °C. Seed was planted 3 plants per #6 standard pot in Cornell Mix soil. Plants were inoculated at three weeks, after emergence of the fourth true leaf. At least three replicates (i.e. inoculation ! 141 ! of three independent plants) were set up for each strain, and experiments were repeated at least three times in all cases. All fungal strains were recovered from -80o C glycerol stocks and grown on CMX for ten days prior to inoculation. Conidia were harvested by scraping plates with ~2 mls 0.02% Tween 20 and filtered through cheesecloth to remove hyphal debris. Conidia were counted with a haemocytometer and adjusted with 0.02% Tween 20 to 2x104 conidia/ml. Conidia were spray inoculated onto maize plants using Preval sprayers (Grace Davison Discovery Sciences, Columbia, MD, U.S.A.), at the rate of 2ml/plant. Plants were placed overnight in a mist chamber, then moved to the growth chamber. 4th true leaves were collected after 3 or 5 days and photographed. ! 142 ! C. Results and Discussion 1. Mapping known Tox1 genes to scaffolds in each C. heterostrophus race T assembly and query for previously unknown associated genes The first task with the new JGI race T strain C4 genome assembly and the two additional race T strain (PR1X412 and Hm338) assemblies was to identify where known Tox1 genes mapped and to determine if new Tox1 sequence could be recovered. Before this work, nine Tox1 genes were known, including five genes at Tox1A, i.e., PKS1 and PKS2, encoding polyketide synthases, LAM1 and OXI1, encoding reductases, and TOX9 encoding an unknown predicted protein, and four genes at Tox1B, three of which, RED1, RED2, RED3, encode reductases, and the fourth, DEC1, which encodes a decarboxylase. All of the known Tox1 genes were found in the previous TMRI and current JGI assemblies of strain C4, as well as in the JGI Illumina Velvet assemblies of the race T field isolates (Fig. III.2). There are some differences in scaffold assembly structure among the five sequenced race T strains that likely say less about differences between the Tox1 loci in each strain and more about the rough quality of the Illumina sequencing, and the de novo Velvet assembly of Hm338 and PR1X412. For example, the three reductase/one decarboxylase cluster on scaffolds 82 and 126 in JGI C4 is known to be a single contig from previous individual gene identification and sequencing work [67]. There is no common thread to the four different RED/DEC assemblies, except that DEC1 assembles with at least one other reductase (but not always the same one). This does not mean that the actual locus differs between the three different strains. We hypothesize that the likely source of these discrepancies is assembly issues related to the highly repeated sequences flanking Tox1 genes ([9,70]). This may be unfortunate, ! 143 ! but it is not surprising. The JGI C. heterostrophus race T genomes were sequenced entirely using Illumina technology. Illumina struggles with repeats, and, in each Dothideomycete genome sequenced, those handled entirely with Illumina are markedly lacking in repeat content [95]. Examination of Tox1-related scaffolds and comparison of the four race T genome assemblies with respect to location of known Tox1 genes did not reveal any new candidate genes (but see Section C.11 below for the discovery of TOX10). ! 144 ! C. heterostrophus JGI C4 LAM1 PKS2 83 TOX9 OXI1 TOX10 PKS1 102 74 RED3 RED1 RED2 DEC1 82 126 82 C. heterostrophus LAM1 TMRI C4 PKS2 TOX9 OXI1 M4LU M597 M3L8 PKS1 M4FP RED2 RED3 DEC1 RED1 M3PL M4QW C. heterostrophus LAM1 Hm338 PKS2 673 TOX9 OXI1 TOX10 1027 1215 PKS1 RED1 RED3 RED2 DEC1 721 1172 745 C. heterostrophus LAM1 PR1X412 TOX9 PKS2 OXI1 TOX10 774 135 1431 PKS1 RED3 RED1 DEC1 RED2 813 1539 1126 1440 Figure III.2. Tox1 scaffolds and genes in all sequenced C. heterostrophus race T strains. The inbred race T strain C4 has been sequenced twice, once by TMRI and once by the JGI. Cartoons of both Tox1 assemblies, as well as the JGI race T field strain assemblies for Hm338 and PR1X412, are depicted. Thin bars represent scaffolds (bold numbers below) and tall boxes are ORFs (labeled above). Scaffolds are colored based on JGI C4 organization. Cartoon is not to scale. For JGI C4 scaffold sizes, see Table III.3. 2. In silico subtraction Because it was possible is that the C4, PR1X412, and Hm338 assemblies contain Tox1 sequence that was not assembled with the known Tox1 scaffolds, but map to undiscovered scaffolds, we performed in silico subtraction analysis using Panseq [82]. Initially, the assemblies of C4, Hm338, and PR1X412 (race T) were collectively used as queries against race O strain C5 to identify race T unique sequence conserved in all race T strains, however this approach was too ! 145 ! restrictive. This method identified only 33 novel regions summing to 17kb, only two of which were over 1kb. To relax the constraint, we conducted a Panseq search using only race T strain C4 as the query against inbred race O C5 reference and then found a sum of 182,418bp in the C4 and not the C5 assembly (Table III.3). The filtered C4 unique sequence was manually screened against the JGI VISTA alignment to C. heterostrophus C5 and C. sativus to determine if it was, in fact, unique to C4 (Fig. III.3). Candidate Tox1 sequences were then located in a whole-genome alignment of the three race T strains, C4, Hm338, PR1X412 and two race O strains, C5, and Hm540 using Mauve [84]. Each scaffold was classified as being present in all race T strains, and being absent in race O strains (Table III.3). Sequence that was not found in all race T strains was not necessarily excluded, as there is some earlier evidence that the Tox1 locus may differ between Tox+ strains. For example, the genetic structure of the PR1X412 Tox1 chromosomes differed considerably from other race T strains in karyotype mapping of RFLP probes [65]. ! 146 ! A B %GC# C.#heterostrophus#C5#VISTA# L.#maculans#VISTA# C.#sa2vus#VISTA# Gene#models# Repeats# C Figure III.3. Graphical representation of visual search strategies for race T unique sequence. A: JGI synteny view with C. heterostrophus race O C5 aligned to race T C4. Whole C4 scaffolds are plotted with intensity of shading representing number of C5 scaffolds mapped. Predominantly white scaffolds indicate C4 sequence not found in C5 (e.g., scaffold 82, arrow). B: JGI VISTA browser view of a C4 unique scaffold. There is no VISTA conservation to C. heterostrophus C5, L. maculans, or C. sativus. There are also no gene calls and GC content is low. ! 147 ! C: Mauve whole-genome alignment of C. heterostrophus race O strains C5 and Hm540, and race T strains C4, Hm338, and PR1X412. Tall red vertical lines represent scaffold boundaries. Blocks of synteny, as represented by Locally Colinear Blocks (LCBs), are colored based on representation within each genome. Mauve colored LCBs can be found in every genome; no LCB means that sequence is either unique to that genome, or, more likely, too repetitive to meet the criteria for an LCB. The centered peach LCB (black arrow) is only found in race T strains. Table III.3: Panseq subtracted race T scaffolds C4 JGI scaffold C4 Prerelease scaffolda Race T: Vista support Race T: Mauveb AT- Genes (JGI ID) richc 74 623 82 476 Yes Yes Yes Yes Yes 155299, 45906 Yes 155403, 155406, 45938, 155400 83 440 Yes Yes Yes 67231, 45941 96 ND No ND Yes 0 101 434 Yes Yes Yes repeats 102 459 Yes ND Yes 155492, 155491 107 650 Yes ND No 45971, 182628 111 ND No ND Yes repeats 116 581 Yes No Yes 29135 119 595 Yes ND Yes simple repeats 120 704 No Yes Yes 54905 121 707 No No Yes 155535 122 538 No No Yes 45984 125 751 Yes Yes Yes repeats 126 476 Yes Yes No 155544 132 541 Yes Yes Yes simple repeats 133 507 No No Yes 45993 134 ND Yes ND Yes transposon 136 541 Yes Yes Yes simple repeats 143 ND Yes ND Yes simple repeats 147 776 No No Yes 0 148 ND Yes ND Yes simple repeats 153 589 No No Yes repeats 157 ND Yes ND Yes repeats 158 908 No No Yes 155608 159 ND No ND Yes 0 161 800 Yes No Yes 46016 176 778 Yes Yes Yes 0 177 637 No Yes Yes simple repeats 179 664 Yes Yes Yes 0 190 ND Yes ND Yes simple repeats ! 148 Annotation PKS1, TOX10 DEC1, RED1, RED3 (transposon) LAM1, PKS2 OXI1, TOX9 HP, HP (myb) HP (protease) HP HP trypsin RED2 HP HP HP Scaffold size (bp) 17210 13683 11416 4430 3655 3528 3474 3368 3011 2895 2887 2880 2840 2671 2644 2499 2404 2370 2271 1829 1718 1700 1631 1517 1496 1488 1477 1289 1285 1230 1121 ! 198 566 No Yes Yes 0 1046 200 863 Yes No Yes repeats 1032 aPrerelease scaffold number is included for archival purpose, and because the 5-genome C. heterostrophus alignment was done with the C4 prerelease. Corresponding scaffolds were determined by blast. bSupported by Mauve means there was no LCB, or no LCB present in Hm540 OR C5. Scaffolds without an LCB in one or both of the race T field strains were still considered supported by Mauve if they were absent in race O strains. cDefined as AT-rich if the browser window was below 35% GC for a substantial portion of the scaffold. Once sequence gaps were removed, there were only 109,995bp across 33 scaffolds found in C4 and not in C5 (Table III.3). Removing scaffolds without at least Vista (found in race O strain C5, or C. sativus) or Mauve support (found in race O strain Hm540) left 22 scaffolds summing to 85,231bp (scaffolds 96, 111, 121, 122, 133, 147, 153, 158, and 159 were removed). All known Tox1 associated scaffolds were recovered from this analysis (C4 scaffolds 74, 82, 83, 102, 126) leaving only 36,750bp on 18 scaffolds as novel, candidate Tox1 sequence. Of these, scaffolds 107, 116, and 120 contained gene calls. Scaffolds 101, 119, 125, 132, 134, 136, 143, 148, 157, 177, 190, and 200 are littered with repetitive elements including transposons, or simple sequence repeats. Scaffolds 176, 179, and 198 contain neither gene calls nor repeats. These scaffolds are typically AT-rich, and quite small (1-3kb). This left only four gene calls on the novel candidate Tox1 scaffolds. Proteins 54905 on scaffold 120 and 45971 on scaffold 107 (JGI ID) have predicted secretion tags, but no predicted functional domains, and match to a small set of proteins in other Dothideomycetes (S. turcica, not L. maculans) with homology >60%. Proteins 182628 (scaffold 107) and 29135 (scaffold 116) also have secretion tags and blast hits to hypothetical proteins in a small set of Dothideomycete relatives; but in addition have functional predictions. 182628 has a predicted SANT/Myb DNA-binding domain (IPR001005), and 29135 has a predicted zinc metallopeptidase M domain (IPR006025). While 182628 may sound like a promising transcription factor candidate, the SANT domain belongs to a retroviral subfamily of cellular Myb DNA binding domains. It is not clear what role, if any, a ! 149 ! metallopeptidase could play in production of T-toxin. When these four proteins were blasted against the D. zeae-maydis genome, 45971 (87% max identity) and 182628 (49% max identity) both retrieve the same hit, 82_g1491 the only gene call on scaffold 82. No other proteins were retrieved from this blast. These genes are all candidates for deletion. Previous work on electrophoretic karyotypes of inbred C4 (race T) and C5 (race O) strains revealed race T contains ~ 1.2Mb more DNA than race O [64,96]. As the inbred strains were backcrossed recurrently to the same race T or race O strain and selected each time for ability to produce T-toxin, there are few differences in the inbred lines other than those found at the Tox1 locus (and the mating type locus). The in silico subtraction performed here found only 85kb, including known Tox1 scaffolds, of sequence likely to be unique to race T. What is the sequence of the remaining 1.1Mb of race T DNA, and why wasn’t it retrieved here? One possible explanation is that the vast majority of this DNA is highly repetitive and that the repetitive scaffolds identified above are actually much larger than their assembled size. It is possible that instead of assembling properly, repetitive reads may be mis-assembled into condensed repetitive units of the size seen in Table III.3. The relative proportion of reads that map to these candidate Tox1 scaffolds could confirm this hypothesis if the read density for these scaffolds is extraordinarily high. 3. Identification of candidate reciprocal translocation breakpoints Tox1A and Tox1B loci map genetically to the reciprocal translocation breakpoints of chromosomes 12;6 and 6;12 respectively (Fig. III.1). The precise physical location is as yet undetermined. All known Tox1 sequence is surrounded by highly repetitive, AT rich sequence, and cannot be assembled with any core genome contig. Scaffolds mapping to chromosomes 6 ! 150 ! and 12 of race O align well with the corresponding sequences from race T chromosomes 6;12 and 12;6 (Fig. III.1). Note that the imperfect race T genome assembly means that the presence of a scaffold break along the alignment does not necessarily correspond to a breakpoint. The main tool we possess to pinpoint the breakpoints is our suite of C. heterostrophus genome sequences. As noted, strains C4 and C5 are highly inbred lines, with only 15,894 SNPs in the entire genome alignment [79]. The relevant difference between the two is the ability to produce T-toxin. In addition to these inbred lines, we have three field strain sequences: the race O strain Hm540, and the race T strains Hm338 and PR1X412. Regions of the genome that have continuous synteny in race O, but where synteny is broken or an alignment is missing in race T, may indicate the physical location of the reciprocal translocation breakpoints. To investigate this, whole genome alignments of all strains were made with Mauve. Only two regions could be identified where alignments were continuous in race O scaffolds, but associated with two different race T scaffolds separated by a gap in race T genomes (Fig. III.4). The first candidate breakpoint (Fig. III.4, red bar) is on C5 race O reference genome scaffold S19, and spans 1800bp. This scaffold could not be mapped to the genetic map (see Fig. II.1). The alignment between race O strains C5 and Hm540 (scaffold S10) is undisturbed along this region. but is broken in race T strains. For example, in inbred C4, two scaffolds S41 and S38 align to C5 S19, separated by a gap. The same is true for Hm338 and PR1X412, where Hm338 scaffolds S472 and S453, and PR1X412 scaffolds S431 and S200 map on either side of the C5 S19 breakpoint. Further, three of the four C4 and Hm338 scaffolds have extra sequence on the aligned scaffolds that does not align with C5 (purple bars, Fig. III.4) and is unique to race T. ! 151 ! race O C5 Hm540 S19 S10 S41 C4 race T Hm338 S472 S431 Pr1x412 441800-443600 S38 S453 S200 race O C5 Hm540 C4 S12 Left 726400-726600 S65 S47 S88 Right 732500-735100 S55 race T Hm338 S262 S760 S200 S415 Pr1x412 S1076 S241 Figure III.4. Candidate sequences for the location of race O chromosome 6 and 12 breakpoints. Two regions, identified by whole-genome alignment of race O (C5, Hm540) and race T strains (C4, Hm338, PR1X412), are candidate positions for the location of reciprocal translocation breakpoints on chromosomes 6 and 12. Syntenic sequence in all strains, both race O and race T (green), is disrupted (red) and but then continues in race O strains only. In race T strains, the scaffold ends, often with small overhangs of race T unique sequence (purple). The first candidate breakpoint (C5 scaffold 19, top) is a single 1800bp region unique to race O strains. In race T, aligned scaffolds reach their ends at the breakpoint, or overhang with unique race T sequence (C4, Hm338). The second candidate breakpoint (C5 scaffold 12, bottom) consists of two separate gaps in alignment (200bp and 2600bp), located ~6kb apart. Synteny is continuous in race O strains, but three separate scaffolds align to this region in each race T strain, again with race T unique overhangs. ! 152 ! The second candidate breakpoint also has undisturbed synteny in race O strains (S12 in C5, S65 in Hm540), with gaps where three different scaffolds of race T align (Fig. III.4). Note that C5 scaffold 12 corresponds to linkage group 12 on the genetic map, one of the two race O chromosomes which are proposed to have undergone a reciprocal translocation. This candidate breakpoint region has not one but two separate breaks in synteny. The first is only 200bp long, and the second break is 2,600bp, located 6kb away. Thus three different race T strain C4 scaffolds (S47, S88, S55) align to single scaffold S12 of race O (Fig. III.4). The same pattern is found for strains PR1X412 and Hm338. Most of the race T scaffolds have overhangs unique to race T (purple bars, Fig. III.4). To build a case for these regions corresponding to the breakpoints of the reciprocal translocation between race O chromosomes 6 and 12, we performed PCR with primer pairs designed to amplify DNA across these proposed gaps. An appropriately sized band could be recovered using race O (C5, Hm540), but not race T (C4, Hm338), template DNA (Fig. III.5). This suggests that in race T, these primers anneal to very different locations than in race O, likely on two separate chromosomes if they indeed span the breakpoint. Final confirmation that the race O/T breakpoints occur at these locations is still lacking, and would rely on Southern hybridization using probes on either side of the proposed breakpoint against electrophoretically separated whole chromosomes. ! 153 ! C4# Hm338# Hm540# PR1X412# L" R" C4# C5# Hm338# Hm540# L" R" L" R" L" R" L" R" Figure III.5. A PCR product spanning the proposed Tox1 related breakpoint on chromosome 12 can be amplified from race O, but not race T, DNA. Top: race O (Hm540), but not race T (Hm338, PR1X412, C4), strains are syntenic in these regions when mapped to the C. heterostrophus race O C5 reference strain. Synteny, depicted as brown plots of read depth in the JGI vista viewer, is lost in two distinct regions (L = left and R = right) when DNA from race T strains is aligned. Bottom: two sets of primers each were designed to amplify products spanning the L and R regions. The correct sized product (L = 3kb, R = 1.5kb, small difference between primer pairs) could be amplified from race O (C5, Hm540), but not race T (C4, Hm338), DNA. Smaller bands (<1kb) in lanes for right (R) primers are nonspecific products. Each set of four lanes shows the two left breakpoint (1, 2) and two right breakpoint (3, 4) primer pairs for each DNA template. Marker is at left. ! 154 ! 4. PKS1 and PKS2 and their phylogenetic neighbors Previous phylogenetic work with ChPKS1 and ChPKS2 found only distant orthologs to these genes in other fungi (Section A.6). The most recent published investigation [70] of all known PKS proteins found support for D. zeae-maydis possessing the closest relative to ChPks1, with Delitschia winteri possessing one relative and Talaromyces stipitatus possessing two, and C. heterostrophus’s own Pks7 rounding out the clade. A Eurotiomycete PKS family (from several Aspergillus species, Neosartorya fischeri, and Penicillium chrysogenum) was sister, but clearly separate, from this Pks1 clade [70]. ChPks2, on the other hand, did not form a clade with any other proteins, with a single neighbor present in Phaeosphaeria nodorum. It was assumed, therefore, that, with the exception of D. zeae-maydis, ChPks1 and ChPks2 were unique to C. heterostrophus race T. Given that many more fungal genomes are now available, we repeated our hunt for orthologs. To this end, we constructed phylogenetic trees of the ketosynthase (KS) domains of ChPks1 and ChPks2, populated with the top 20 blast hits for each. In addition, we included the KS domains of all C. heterostrophus, D. zeae-maydis, L. maculans, and T. stipitatus PKSs, as well as bacterial FabF outgroups. ChPks1 formed a single, well supported clade with a D. zeae-maydis PKS (Dzm1066 g6391, MzmPks1 Genbank ID# 40388707) and L. maculans (Lm11520) while D. winteri (Dw Genbank ID# 282160401) and T. stipitatus (Ts645921431) have a single PKS each, sister to this (Fig. III.6). ChPks7 (also found in other Cochliobolus species [79]) groups exterior to this, along with an additional T. stipitatus PKS (Ts645918889). L. maculans and D. zeae-maydis, therefore, both have well-supported ChPks1 orthologs, with weaker candidates in T. stipitatus (potentially 2) and D. winteri. As in previous studies, all other ChPKS1 blast hits form a clade sister to this, consisting largely of PKSs from Eurotiomycete species (Fig III.6, for more distant taxa see full tree in Appendix). ! 155 T. atroviride 358393293 52 85 100 T. virens 358385196 69 ! 55 T. reesei 59482 A. nidulans 350639053 58 A. terreus 115386424 T. stipitatus 645913763 100 0.1 P. chyrsogenum 32119 C. victoriae 29158 100 C. heterostrophus Pks7 55 81 90 98 D. winteri 282160401 T. stipitatus 645921431 C. heterostrophus Pks1 100 72 L. maculans 11520 D. zeae-maydis 1066/_g6391.t1 Pks1% 99 T. stipitatus 645918889 P. digitatum 425773093 99 P. chyrsogenum 80687 A. oryzae 317138801 58 A. fumigatus 238504606 100 A. oryzae 83764427 A. oryzae 391873863 97 A. fumigatus 146323155 31 99A. fumigatus 159128403 100 N. fischeri 002360 A. clavatus 121712301 Figure III.6. Phylogenetic relationship of ChPks1 and relatives. A phylogenetic tree was constructed with the KS domain from a large set of PKSs, including every PKS from C. heterostrophus, L. maculans, D. zeae-maydis, and T. stipitatus, 14 additional species, and the top 20 blast hits for ChPks1 and ChPks2. Each sequence is identified by either the Genbank number, JGI protein ID (C. heterostrophus, T. stipitatus, L. maculans), or Augustus ID (D. zeae-maydis), accompanied by a 3-letter strain abbreviation (Table III.4). Only a portion of the tree showing proteins surrounding ChPks1 is shown here. The full tree is available in the Appendix. Note that ChPks1 (red) groups with L. maculans 11520 and D. zeaemaydis 1066_g6391 with 98% bootstrap support (arrow), and a single protein each from D. winteri and T. stipitatus cluster sister to this. ChPks2 also forms a single, well-supported clade, clustering with PKSs from L. maculans (Lm11513) and D. zeae-maydis (Dzm1284 g8769) (Fig. III.7). These are the first PKSs identified to form a strong phylogenetic relationship with ChPks2. Sister, but with poor support, are a number of sequences from a variety of Sordariomycetes. No other Dothideomycete PKSs cluster with ChPks2, but D. zeae-maydis and L. maculans each have a PKS (Dzm986 g4383 and Lm661, respectively), that clusters, along with T. stipitatus ! 156 84 70 ! 100 89 Lma1999|Lema_T082310.1/9439 Stu185084|estExt_fgenesh1_pg.C_290004/9442 Ptt8903|PTTG_17304.1/9442 Pdi425773714 Ptt9714|PTTG_18319.1/4427 Cvi31369|gm1.9973_g/4427 Ts645913763, in100a dChies1t1i6n870c7t|P,ksb6 ut sister clade (see Appendix for full tree). The finding that D. zeae100 48 Che1229323|Pks6 (2) 2 21maydis possesses6,4 not only aDzm1059/_g6254.t1/4427 ChPKS1 but also a ChPKS2 ortholog, is particularly interesting Stu171983|estExt_fgenesh1_pm.C_3_t10424/4304 because the PM-toxinAcl121704114| family of linear polyketides has a shorter average carbon backbone aim326147|gm1.6147_g/58483 40 compared to the T-toxinFox|10408|FOXG_02741/32457 family. As only a single D. zeae-maydis PKS (MzPKS1) and two Tst645922713/6427 100 reductases (MzRED1, MzRED2) were previously knownPtt8650|PTTG_16988.1/7444 100 to be associated with PM-toxin, it was Lma266|Lema_T002660.1/7419 6 hypothesized that the reduced biosynthetic geneCgr115742|estExt_fgenesh1_pm.C_550003/20435 content (particularly having only one PKS) was Dzm986/_g4383.t1/105524 100 82 Lma661|Lema_T006610.1/94514 connected to the reduced size of PM-toxin [9,70]sm336271734|ref|XP_003350625.1|/88512 Locating an ortholog to ChPKS2 (as well as 69 99 440489879|gb|ELQ69490.1|/169593 many affiliated Tox1 genes, see Section C.8 below)29 440474381|gb|ELQ43129.1|/169593 nullifies this hypothesis. Instead, the 69 100 389625625|ref|XP_003710466.1|/67491 abbreviated structur1e00 of PM-toxin may be dueGgr310792755|gb|EFQ28216.1|/91515 to the slight differences in the reductase and 62 100 Cor477530347|gb|ENH82089.1|/86510 oxidoreductase inventory.Ssc6497|SS1G_01997T0/197404 Bfu472238180|gb|EMR83055.1|/70493 100 Bfu347830656|emb|CCD46353.1|/70493 Bfu40787340|gb|AAR90244.1|/70499 78 0.1 Pch70100|e_gw1.2.3250.1/40459 65 Cgr60275|estExt_Genewise1.C_130160/1417 66 40 G. clavigera 320587585 D. zeae-maydis 1284/_g8769.t1 100 41 C. heterostrophus Pks2 L. maculans 11513 Pks2% 97 V. dahliae 346978574 95 100 C. purpurea 399167504 63 N. haematococca 106474 T. atroviride 358393293 52 85 100 T. virens 358385196 69 55 T. reesei 59482 A. nidulans 350639053 58 A. terreus 115386424 T. stipitatus 645913763 100 P. chyrsogenum 32119 C. victoriae 29158 100 Figure III.7. PhylogCe. hneteerotsitrcophrues Plakst7ionship of ChPks2 and relatives. D. winteri 282160401 See55Fig. 6 f9o0 r methods. Only T. stipitatus 645921431 a portion of the tree showing proteins surrounding ChPks2 is shown here; th98e full treeC. is presentedheterostrophus Pks1 in the Appendix. Note that ChPks2 (in red) groups with L. maculans 11513 a10n0 d D. zeae-maydisL. maculans 11520 1284_g6769 with 100% bootstrap support (arrow). All other PKSs fr8o1 m all o72ther species,D. zeae-maydis including1066/_g6391.t1 the top 20 blast hits, group outside. 99 T. stipitatus 645918889 P. digitatum 425773093 99 P. chyrsogenum 80687 A. oryzae 317138801 ! 58 A. fumigatus 238504606 100 A. oryzae 83764427 157 A. oryzae 391873863 97 A. fumigatus 146323155 31 99A. fumigatus 159128403 ! 5. Polyketide synthases inventories To speculate on the genetic origins and distribution of ChPKS1 and ChPKS2, the distribution of other PKS genes must also be considered. To address this, the complete PKS inventory of C. heterostrophus, D. zeae-maydis, L. maculans, and T. stipitatus, and 14 additional species from within the Pezizomycotina was determined (Table III.4), and the KS domains subjected to phylogenetic analysis (see Fig. S11 in the Appendix for full tree). This utilized 376 PKSs from 7 Dothideomycete, 1 Pezizomycete, 5 Eurotiomycete, 1 Lecanoromycete, 1 Leotiomycete, and 3 Sordariomycete genomes. The PKS inventory for these species varied from 3 in the coprophile Ascobolus immersus to 42 in T. stipitatus, and, in general, fell into classspecific clades, with poor support outside of the class. Table III.4. Species included in PKS KS domain tree Species, strain, versiona Symb Lifestyle (host) Class Cochliobolus heterostrophus C4 v1.0 Cochliobolus victoriae FI3 v1.0 Cochliobolus sativus ND90Pr v1.0 Setosphaeria turcica Et28A v1.0 Didymella zeae-maydis F3018 v1.0 Leptosphaeria maculans ‘brassicae’ JN3 v1.0 Pyrenophora teres f. teres NFNB 15A v1.0 Ascobolus immersus RN42 v1.0 Talaromyces stipitatus ATCC10500 v1.0 Aspergillus fumigatus Af293 v1.0 Aspergillus clavatus NRRL 1 v1.0 Che Cvi Csa Stu Dzm Lm Ptt Aim Ts Afu Acl Neosartorya fischeri NRRL 181 v1.0 Penicillium chyrsogenum v1.0d Nfi Pch phytopathogen (corn) phytopathogen (oats) phytopathogen (cereals) phytopathogen (corn) phytopathogen (corn) phytopathogen (canola) phytopathogen (barley) coprophile saprophyte pathogen (mammalian) opportunistic pathogen (mammalian) saprophyte saprophyte Dothideomycetes Dothideomycetes Dothideomycetes Dothideomycetes Dothideomycetes Dothideomycetes Dothideomycetes Pezizomycetes Eurotiomycetes Eurotiomycetes Eurotiomycetes Eurotiomycetes Eurotiomycetes # KS domains recoveredc 25 24 20 27 14 15 24 3 42 17 23 19 22 ! 158 ! Cladonia grayi Cgr lichen Lecanoromycetes 36 Cgr/DA2myc/ss v1.0 Sclerotinia sclerotiorum Ssc phytopathogen Leotiomycetes 20 1980 ATCC 18683 v1.0 (broad) Trichoderma reesei v2.0 Tri saprophyte Sordariomycetes 14 Nectria haematococca Nha phytopathogen Sordariomycetes 15 MPVI isolate 77-13-4 v2.0 (pea) Fusarium oxysporum f. sp. Fox phytopathogen Sordariomycetes 16 lycopersici 4287 race 2, (tomato) VCG 0030 v1.0 a Except for D. zeae-maydis, all genomes downloaded from JGI web portal, although not all sequenced by them. b Sym: Species abbreviations in tree. c PKS recovered includes some FAS. d P. chrysogenum has no strain designation, as it was a contaminant from the Postia placenta MAD 698R dataset. The top 20 Blast hits for ChPks1 and ChPks2 from the following species are also included: Penicillium digitatum-Pdi, Aspergillus oryzae-Aor, Aspergillus nidulans-Ani, Aspergillus terreus-Ate, Trichoderma virens-Tvi, Trichoderma atroviride-Tat, Claviceps purpurea-Cpu, Verticillium dahliae-Vda, Grosmannia clavigera-Gcl, Colletotrichum orbiculare-Cor, Glomerella graminicola-Ggr, Botryotinia fuckeliana-Bfu, Sordaria macrospora- Sma, Magnaporthe oryzae-Mor. Each PKS from C. heterostrophus, D. zeae-maydis, and L. maculans was classified as to whether or not it had orthologs in C. heterostrophus, D. zeae-maydis, L. maculans, or T. stipitatus, if it was distributed in a Dothideomycete specific clade with strong support, and if it followed the expected species tree or not with its neighbors (Table III.5). Genes that have phylogenetic patterns unlike that expected from a species tree are often speculated to be the product of horizontal gene transfer (HGT). However, the frequent and rapid duplication/loss/recombination of secondary metabolite genes, and the diversity of these genes in individuals within a species (such as in C. heterostrophus [79]) makes garnering strong support for HGT extremely difficult, and other mechanisms may plausibly explain their distribution [74]. Of the 25 C. heterostrophus race T PKSs, there are six clear candidate orthologs in D. zeae-maydis (ChPks1, 2, 3, 6, 21, 25), four in L. maculans (ChPks1, 2, 9, 18), and only one (ChPks1) in T. stipitatus. If the candidate ortholog criteria are relaxed to allow more ambiguous relationships to account for T. stipitatus being a distant relative, orthologs can be postulated for three additional PKSs (ChPks5, 19, and 21)- but in each case there are multiple candidate ! 159 ! orthologs, indicating duplications. Three PKSs (ChPks7, 11, 15) were Cochliobolus-specific, shared only with C. victoriae or C. sativus. Table III.5. PKS inventories and distribution patterns Protein Candidate Description of distribution orthologs ChPks1 ChPks2 ChPks3 ChPks4 ChPks5 ChPks6 ChPks7 ChPks8 ChPks9 ChPks10 ChPks11 ChPks12 ChPks13 ChPks14 ChPks15 ChPks16 ChPks17 ChPks18 ChPks19b ChPks20 ChPks21 ChPks22 ChPks23 ChPks24 ChPks25 DzmPks269 g2961 DzmPks1250 g8623 DzmPks1166 g7942 DzmPks1002 g4912 DzmPks1013 g5164 DzmPks1108 g7085 DzmPks1924 g11384 DzmPks1250 g8622 DzmPks1059 g6254 DzmPks986 g4383 DzmPks1284 g8769 DzmPks1066 g6391 Lm1717 Lm3382 Lm2891 Lm3340 Lm82 Lm6852 ! Dzm1066g6391, Lm11520, Ts645921431a Dzm1284g8769, Lm11513 Dzm1250g8622 Dzm1059g6254 Lm5757 NA Lm1166g7942, Ts Dzm1250g8623 Dzm269g2961 Ch25 Ch21 Ch18 Lm1817 Ch3 Ch6 Lm661 Ch2, Lm11513 Ch1, Lm11520, Ts645921431 Ts645923869 Ch18 * * Dothideomycete clade Discontinuous Dothideomycete clade Dothideomycete clade Cochliobolus specific Dothideomycete clade Dothideomycete clade Dothideomycete clade Cochliobolus specific Dothideomycete clade NA Dothideomycete clade Cochliobolus specific Dothideomycete clade Discontinuous Dothideomycete clade Dothideomycete clade Discontinuous Dothideomycete clade Dothideomycete clade Dothideomycete clade Dothideomycete clade Discontinuous Discontinuous Dothideomycete clade Dothideomycete clade Discontinuous Unique Unique Discontinuous Dothideomycete clade Dothideomycete clade * * * Discontinuous Dothideomycete clade Unique Discontinuous Dothideomycete clade Discontinuous 160 Follows species tree? * * Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes No NA NA No No Yes * * * No Yes NA Yes Yes No ! Lm1817 Dzm1924g11384 Discontinuous No Lm5757 Ch9 Dothideomycete clade Yes Lm3338 Dothideomycete clade Yes Lm1999 Discontinuous Yes Lm266 Ts645922713 Discontinuous Yes Lm661 Dzm986g4383, * * Ts645913767 Lm11513 Ch2, * * Dzm1284g8769 Lm11520 Ch1, * * Dzm1066g6391, Ts645921431 a Talaromyces orthologs included only where there was a close relationship. b ChPks19 has three distant T. stipitatus orthologs with some bootstrap support, omitted as orthologs. * Pks1 and Pks2 related. NA – Not applicable (ChPks13 is a pseudogene [79]). Seventeen C. heterostrophus PKSs formed strong Dothideomycete specific clades, although these were not necessarily unique to the Dothideomycetes. Curiously, ChPks18, responsible for the production of melanin, had good support (>80 bootstrap value) only within the Dothideomycetes. With very few exceptions, phylogenetic support for a given PKS clade is only maintained within the Dothideomycetes and attempts to expand a clade to include species from other fungal classes lack bootstrap support. These exceptions (ChPks4, 17, 20, 25), classified as “Discontinuous,” are not broadly conserved within the Dothideomycetes. Instead, they have only a single Dothideomycete ortholog (if at all), but have candidate orthologs from species in the Eurotiomycete (not T. stipitatus) or Sordariomycete class. This highly unusual distribution pattern is to be expected when comparing secondary metabolism genes from such distantly related species. Even within a particular class, most secondary metabolism genes are not conserved (see Table S20 for PKSs [95]). It is safe to assert, therefore, that the strong support afforded T. stipitatus Ts645921431 clustering near ChPks1 is unlike all other PKSs in this analysis, and it is a robust candidate ortholog. ! 161 ! 6. ChPks1 and ChPks2 comparisons The maximum blast identity of the full C. heterostrophus Pks1 sequence to its closest ortholog in each species is 72% to L. maculans, 61% to D. zeae-maydis, 51% to D. winteri, and 49% to T. stipitatus (and 46% to a second T. stipitatus PKS). All other blast results in Genbank are below 40%. Pks2 is 60% similar (max identity) to its L. maculans ortholog and all other Genbank results are <40% similar. D. zeae-maydis must be blasted separately, as its Pks2 ortholog is not in Genbank. For context, C. heterostrophus PKS9 is 57% similar to its L. maculans counterpart, and PKS18 is 85% similar. Pks1 and Pks2 are therefore not unusually similar to each other in C. heterostrophus and L. maculans compared to other shared PKSs. The %GC of C. heterostrophus, D. zeae-maydis, L. maculans, and T. stipitatus PKS1 and PKS2 transcripts was also considered (Table III.6). The average %GC all PKSs within a species ranged from 47.9-52.5%, with PKS1 and PKS2 transcripts ranging from 49.1-56.4%. L. maculans %GC is slightly higher in all transcripts, and especially for PKS2. This equilibrated %GC does not suggest extraordinary origins for these genes. Table III.6. %GC content in PKS transcripts. All PKSs Species (Average) PKS1 PKS2 C. heterostrophus 51.7 50.8 50.6 L. maculans 52.5 52.4 56.4 T. stipitatus 47.9 49.1 NA D. zeae-maydis* ND 54.7 53.9 PKSs transcript sequences were downloaded from the JGI Mycocosm site, and %GC was calculated using a Perl script. Average %GC is based on transcripts of all PKSs in the genome. *The D. zeae-maydis transcript catalog is not available. Instead, only the PKS1 and PKS2 %GC values were calculated. ! 162 ! 7. T-toxin activity of D. zeae-maydis, L. maculans, and T. stipitatus The discovery of new PKSs highly and moderately similar to ChPks1 and ChPks2 immediately raises questions regarding the corresponding polyketide produced, and the activity of these metabolites. Cultures of L. maculans and T. stipitatus were obtained and tested for Ttoxin activity. C. heterostrophus race T T-toxin and D. zeae-maydis PM-toxin are already known to display biological activity against transgenic E. coli carrying the maize mitochondrial gene and toxin target T-Urf13 (Section A.2), and together with C. heterostrophus race O, serve as positive and negative controls. Startlingly, plugs of T. stipitatus formed a halo around the plug on a lawn of E. coli expressing T-Urf13, but not on control lawns of E. coli carrying a control pUCATPH plasmid without T-Urf13 (Fig. III.8). This makes T. stipitatus the third fungal species, after C. heterostrophus race T and D. zeae-maydis, to show toxicity to T-Urf13 expressing cells. T. stipitatus is not a known pathogen of corn or any other plant, but is rather thought to be a soildwelling fungus. T. stipitatus does not have a clear Pks2 ortholog, although it does have a distant and unsupported Pks2 relative (Tst645913763, see Fig. III.7). The remaining question is: what molecule confers T-toxin-like activity to T. stipitatus? The presence of a Pks1 ortholog coupled with the knowledge that T-toxin and PM-toxin are polyketides overwhelmingly suggests that T. stipitatus produces a polyketide. Both T-toxin and PM-toxin are linear, although their carbon chain lengths vary greatly. The unknown T. stipitatus metabolite, therefore, could vary structurally from T-toxin while still triggering cell death to TUrf13 expressing cells. Absence of a clear PKS2 ortholog (and clear orthologs of all other Tox1 genes, see section C.9 below) makes this a likely scenario, although TsPks1 could work with other unrelated genes (including PKSs) in synthesizing its polyketide. The genomic region ! 163 ! surrounding this T. stipitatus PKS1 gene does not include any genes with predicted domains matching those found in Tox1. Figure III.8. Talaromyces stipitatus displays T-toxin-like activity in an in vitro microbial assay. Plugs of mycelium were plated onto lawns of E. coli carrying the T-URF13 gene (left). C. heterostrophus race T isolates (C4, Aus26239 A3) inhibit E. coli growth surrounding the plug, but race O isolates (C5, Aus19679-2) do not. D. zeae-maydis isolate PM24, producing PMtoxin, also produces a halo, as PM toxin has the same biological activity as T-toxin. T. stipitatus (Ts1006) produced a smaller, but distinct halo, indicating activity against the same T-URF13 target. Plates spread with a control E. coli strain carrying a control plasmid, pUCATPH which lacks the T-URF13 gene (right), are not cleared by mycelial plugs. Like D. zeae-maydis, L. maculans strain JN3 has both a ChPKS1 and ChPKS2 ortholog: it seems likely that it produces a polyketide with T-toxin-like activity. T-toxin assays using Turf13 expressing E. coli carried out in our lab found no evidence of such activity in the L. maculans isolates (Lm893, Lm939, Lm100-3, Lm421-3, Lm847, Lm81, Lm846, and Lm855) that could be found in the Cornell Department of Plant Pathology and Plant-Microbe Biology ! 164 ! isolate collection (Fig. III.9), but with the following caveats. None corresponds to the sequenced L. maculans strain JN3, the geographical origins and genotype of each of these is unknown, and their morphological characteristics are not uniform (data not shown). Furthermore, and perhaps most significant of all, it is likely that many of these isolates would no longer be classified as L. maculans, given the tumultuous history of L. maculans and L. biglobosa (Section A.1.iii). The Turgeon lab is in the process of acquiring the sequenced strain from Australia for testing. Lm893$ Lm838$ Lm100'3$ Pl85% Lm421'3$ Lm847$ Lm81% Lm846% Lm855$ ChC4$ ChC5$ Lm893$ Lm838$ Lm100'3$ Pl85% Lm421'3$ Lm847$ Lm81% Lm846% Lm855$ ChC4$ ChC5$ Figure III.9. L. maculans strains in the PPP-MB collection do not demonstrate T-toxin activity in in vitro assays. Plates prepared as in Fig. III.8. C. heterostrophus race T isolate C4 is a positive control and produces a halo around E. coli carrying the T-URF13 gene, but control race O isolate C5 does not. None of the L. maculans isolates from the Cornell isolate collection (Lm893, Lm838, Lm100-3, Pl85, Lm421-3, Lm847, Lm81, Lm846, Lm855) display T-toxin activity. Clearing is specific to E. coli expressing T-URF13, as E. coli carrying pIGPAPA (right) which lack TURF13 are not affected by any of the mycelial plugs. To overcome these shortcomings, we are collaborating with Dr. Candace Elliott in the laboratory of Dr. Barbara Howlett at the University of Melbourne in Australia (where Lm11513 and Lm11520 were first recognized as having homology to ChPKS1 and ChPKS2) to assay toxin ! 165 ! production in L. maculans JN3. This investigation is still underway, but preliminary results do not support L. maculans producing a molecule with T-toxin activity. There is evidence that the respective ChPKS1 and ChPKS2 orthologs, Lm11520 and ChPKS2 Lm11513 (PKS13 and PKS14 in L. maculans nomenclature) are indeed expressed in 10% V8 under 12h light/ 12h dark conditions [48]; it is not known if they are expressed under the conditions used to assay T-toxin either in our laboratory or in our Australian collaborators’ laboratory. While there is no report of a race highly virulent on a canola host, the genetic structure of the Tox1-like locus in L. maculans strongly suggests that a molecule similar to Ttoxin is produced (Section C.9). Aggressive L. maculans strains, such as L. maculans brassicae JN3 and IBCN18, and L. maculans lepidii IBCN84 possess LmPKS13 and LmPKS14, whereas the less aggressive L. biglobosa strains (IBCN65, J154) do not (C. Elliott, personal communication). This suggests that Lm11520 and Lm11513 could produce a polyketide that results in high virulence on canola, even if that polyketide does not have T-toxin-like activity on T-urf13. 8. Phylogenetic distribution of Tox1 affiliated gene relatives ChPKS1 and ChPKS2 are not the only genes located at Tox1. Phylogenetic and genomic characterization of eight additional Cochliobolus Tox1 genes (DEC1, LAM1, OXI1, TOX9, TOX10, RED1, RED2, RED3) lend support to the suspicion that L. maculans does, in fact, produce a polyketide with T-toxin activity, and that PM-toxin biosynthesis is far more similar to T-toxin biosynthesis than previously thought. On the other hand, the activity displayed by T. stipitatus becomes even more puzzling given the absence of so many genes associated with Ttoxin biosynthesis. ! 166 ! Relevant sections of phylogenetic trees for each C. heterostrophus Tox1 gene and its top 50 blast hits are provided in Figs. III.10-16, and complete trees are available in the Appendix (Figs. S11-S16). To address the convoluted relationship among reductases, a single phylogenetic tree was constructed with ChRed1, ChRed2, and ChRed3 and their orthologs. ChDec1 forms an isolated clade with D. zeae-maydis and L. maculans proteins with 100% bootstrap support (black arrow, Fig. III.10). Within this clade, C. heterostrophus clusters with D. zeae-maydis with 100% bootstrap support, and L. maculans falls outside with no support. The closest blast-derived neighbors reside on a separate, but well-supported branch, and include an additional C. heterostrophus member (Fig. S12), suggesting that these additional proteins are relatives of a different decarboxylase, and that the Tox1 DEC1 is unique to C. heterostrophus, D. zeae-maydis, and L. maculans. There are no orthologs of T. stipitatus nearby. ChLam1 forms a well-supported isolated clade with L. maculans and D. zeae-maydis orthologs (Fig. III.11). The Dothideomycete maize phytopathogen, Macrophomina phaseolina, possesses a Lam1-like protein clustering nearby with poor support. No other proteins (including from T. stipitatus) cluster with any support near ChLam1 (Fig. S13). ! 167 ! 0.1 55 29 55 31 69 100 13 69 64 336260005| 336260030052|404228| 93 44 100 85068394|r 100 100 85068394|r 100 104051852656| 336463110| 71 336463110| 71 485926052| 59 407928516| 336260476| 336260476| 451997090| 42 452845199| 302893743| 452845199| 100 55 31 100 453087398| 477515868| 95 453087398| 475665176| 100 169626440| 169626440| 46127601|r 100 100 Fusarium oxysporum475677038 171695019020| 171695092| Fusarium oxysporum 477522859 100 33627G6r1o9s7m|annia clavigera 336276197| 320594062 100 164429671| Se1to0s0phaeria tu1rc6ic4a4428926870810|30 100 Coc1h0li0obolus heterostrophus 350297343| 25090017040Dec1 350297343| Did1y0m0ella zeae-maydis 1210 100 Dec1% 64 336463440| Leptosphaeria 336463440| maculans 396458983 471573773| 47014777195778|3773| Figure III.10. The decarboxylase Dec1 is unique34t6o322C337.| heteros34t6r32o23p37h| us, D. zeae-maydis, and L. maculans. 100 100 89 89 400602833| 400602833| A phylogenetic tree was constructed using the C. heterostrophus Dec1 protein sequence, and its top 50 blast matches retrieved from Genbank3.428A7569s6|ingle protein342875696| in D. zeae-maydis clusters with the C. heterostrophus D24ec1 sequenc9264e (red) with7896strong boot78strap support, which in turn is on a well supported branch (arrow) with Neighboring taxa are labeled by species nLa.mmeaacnudlaGn140s470e7576ns5625eb1130pa715|an| rkatperofr1t4o0e47075mi76n562511oI3071Dt5| |h.erBdreacnacrhbolexnygltahsses. represent # substitutions/site. See Fig. S12 for 79 full t9r7ee.79 475665102| 97 475665102| 0.1 0.1 322704098| 322704098| Botryotinia fuckeliana 3474B4o1t3r6yo5tinia fuckeliana 347441365 99 99 Glarea lozoyensis 3611305G5l0area lozoyensis 361130550 Macrophomina phaseolina M40a7c9ro1p9h9o3m6 ina phaseolina 407919936 64 64 Leptosphaeria maculans 39L6e4p5to8s9p6h3aeria maculans 396458963 100 75 Lam1%100 Cochliobolus heterostrophuCso2c2h7li9o3b7o6lu3s5hLeatemro1strophus 227937635 Lam1 75 Didymella zeae-maydis 128D4id_ygm87e6ll8a zeae-maydis 1284_g8768 407925638| 407925638| Figure III.11. The 3-hydroxacyl-CoA dehydrogenase Lam1 is unique to C. heterostrophus, D. zeae-maydis, and L. maculans. A phylogenetic tree was constructed using the C. heterostrophus Lam1 protein sequence, and its top 50 blast matches retrieved from Genbank. A single protein in D. zeae-maydis and L. maculans clusters with the C. heterostrophus Lam1 sequence (red) with strong bootstrap support (arrow). A protein from Macrophomina phaseolina (Genbank ID 407919936) clustered nearby with poor support. Branch lengths represent # substitutions/site. See Fig. S13 for full tree. ! 168 17 302882457| 33 ! 0.1 80 100 52 429855427| 95 380493535| 358368173| 80 145251930| 58 350633388| 100 70995420|r 29 238503640| 100 169765079| Macrophomina phaseolina MS6 407917317 100 Neofusicoccum parvum 485922801 Macrophomina phaseolina MS6 407921290 71 Neofusicoccum parvum 485917205 27 Macrophomina phaseolina MS6 407923265 56 53 77 21 Macrophomina phaseolina MS6 407920387 Macrophomina phaseolina MS6 407926197 70 Macrophomina phaseolina MS6 407918562 Marssonina brunnea f. sp. 'multigermtubi' 406865315 21 41 Macrophomina phaseolina MS6 407929697 Cochliobolus heterostrophus 283484401 Oxi1 50 59 Glomerella graminicola 310800434 Leptosphaeria maculans 396458971 Oxi1% 86 Didymella zeae-maydis 234/_g2697 98 Didymella zeae-maydis 1066/_g6389 Neofusicoccum parvum 485923792 115401466| Figure III.12. The short chain reductase Oxi1 is found in many Tox1- taxa, and is duplicated in D. zeae-maydis. A phylogenetic tree was constructed using the C. heterostrophus Oxi1 protein sequence and its top 50 blast matches retrieved from Genbank. A single protein in L. maculans and two in D. zeae-maydis cluster with robust support. The C. heterostrophus Oxi1 sequence (red), along with proteins from several other taxa, cluster with poor bootstrap support. Taxa are labeled by species name and Genbank protein ID. Branch lengths represent # substitutions/site. See Fig. S14 for full tree. ChOxi1 does not group in a well-supported clade, instead it appears in a weakly supported group that includes proteins from D. zeae-maydis, L. maculans, Glomerella graminicola, and M. phaseolina (Fig. III.12). There are two D. zeae-maydis proteins in this clade, both of which are likely associated with PM-toxin production as suggested by their genomic organization (section C.9). Interestingly, there is a greatly expanded group of M. phaseolina proteins sister to the weak Oxi1 clade. Again, there are no T. stipitatus orthologs. ! 169 119184609| 12 115390220| 347830239| 100 ! 97 154323332| 156057957| 303315463| 36 ChTox9 15 4g8 roups with fair support (76%73 238495584| 391872242| bootstrap) in a clade with L. maculans and D. dzm_197/_g 91 72 dzm_1115/_ 169778301| zeae-mayd5i5s orthologs (Fig. III.13). The482812025| Cochliobolus heterostrophus 451992688119191035| closest blast hits from species from a diverse set of 19 Bipolaris sorokiniana 451849551 88 Pyrenophera teres f. teres 330915738922868971| 189203427| 396472589| 119474077| fungal classes (and not including T. stipitatus) group nearby without support.40 82 100 100 Magnaporthe oryzae 389637824 358373241072|072610| 310800832| 89 10698979080|38048139417|7531318| 68 258571639| 472235687| 100 119182464| 99 100 303319197| 347829254| 145241696| 58 350639877| 0.1 Exophiala dermatitidis 37872493077 67541238|r 367049110| 98 Pseudocercospo1ra3 fijiensis 452977842 70 310799229| Talaromyces stipitatus 242760721 6 31 31 Dothistroma sep1to0sporum 452837988 Talaromyces marneffei 212539213 358395046| 45 Coniosporiu4m5 apollinis 4948272889 22 Neofusicoccum parvum 485925466 100 54 20 29 67523149|r 93 67901824|r Talaromyces stipitatus 242821894 119184609| 452845441| Macrophom12ina phaseolina 407929441 115390220| 59 347830239| 97 154323332| Didymella zeae-maydis 92_g1676 156057957| 76 36 15 48 303315463| Cochliobolus heterostrophus Tox9 76 73 238495584| Leptosph3a9e1r8ia72m2a4c2u|lans 396458973 91 169778301| Tox9% 11919D1i0d3y5m|ella zeae-maydis 234_g2969 392868971| 119474077| 89 68 358373217| 106879080| 472235687| 347829254| Figure III.13. The protein of unknown function Tox9 appears unique to C. heterostrophus, D.6 31 Exophiala dermatitidis 378729077 98 Pseudocercospora fijiensis 452977842 Dothistroma septosporum 452837988 zeae-maydis, and L. macula4n5 s. Coniosporium apollinis 494827289 Neofusicoccum parvum 485925466 Macrophomina phaseolina 407929441 and its A phylogenetic tree was cons5t9ructed usi7n6 g the top 50 blast matches retrieved from Genbank. C. heterostrophus Tox9 protein sequence76 Cochliobolus heterostrophus Tox9 Leptosphaeria maculans 396458973 A single protein in L. maculans and two inDidymella zeae-maydis 234_g2969 D. zeae-maydis cluster with the C. heterostrophus Tox9 sequence (red) with moderate bootstrap support (arrow). A second D. zeae-maydis protein is highly divergent and the corresponding branch is truncated (double break lines). See Fig. S15 for full tree. TOX10 is a novel Tox1 gene discovered in the latest assembly of C. heterostrophus C4, adjacent to PKS1 (see Section C.9 below), encoding a small protein with homology to very few known proteins in Genbank and no predicted domains (Fig. III.14). The two closest relatives are in D. zeae-maydis and L. maculans (albeit without good bootstrap support), and are also small proteins without predicted domains. There is no Tox10 ortholog in T. stipitatus. ! 170 ! 0.1 Bacillus cereus 487958209 Cyanothece sp. 281202799 Phaeosphaeria nodorum 169598214 Setosphaeria turcica 482805247 83 62 Macrophomina phaseolina 407925866 92 Cochliobolus sativus 451849807 40 56 68 45 Cochliobolus heterostrophus 477581482 Tox10 26 Leptosphaeria maculans 396458979 40 Didymella zeae-maydis 1284_g8767 Tox10& Thermus oshimai 410731447| Mycoplasma gallisepticum 31544418 Cyanothece sp. 218245179| Figure III.14. The protein of unknown function Tox10 does not form a strong phylogenetic relationship across C. heterostrophus, D. zeae-maydis, and L. maculans. A phylogenetic tree was constructed using the C. heterostrophus Tox10 protein sequence and its top 50 blast matches retrieved from Genbank; highly divergent sequence was manually removed. A single protein each in L. maculans and D. zeae-maydis clusters with the C. heterostrophus Tox10 sequence (red), but with very poor bootstrap support. The dataset formed three other, poorly supported clades: two bacterial, and a third including Macrophomina phaseolina, Cochliobolus sativus, and Setosphaeria turcica. This tree is shown at a larger scale than other Tox1 trees to include all taxa. ChRed1 forms a well-supported clade with D. zeae-maydis 1210g8374 (Fig III.15). There are no L. maculans (or T. stipitatus) orthologs that cluster with ChRed1. Instead, M. phaseolina and N. parvum have single proteins each outside of the ChRed1 grouping. ChRed2 and ChRed3 cluster together with poor support (Fig. III.16). Neighbors include a single D. zeaemaydis reductase (1066g6390) and two L. maculans reductases (11514 and 11518), which cluster ambiguously around ChRed2 and ChRed3. Additional reductases from varied species (but not T. stipitatus) cluster nearby without support. ! 171 6 28 47 55 13 47 226295266| 83 65 327351973| Aspergillus fumigatus 70996344|r 67 31 3 100 261194398| 39 Aspergillus terreus 115397645| ! 31 75 1542827536| 3 N45eosartorya fischeri 119494755| 29450274605| Aspergillus clavatus 121700865| 72 Aspergillus oryzae 169767282| 225561503| Coccidioides posadasii 320036866| C98occidioides 326473399| po8s3adasii 303317974| 21 11 28 55 67 327294940| 27 Coccidioides 9im9mitis 119186175| 2 Cochliobolus heterostrophus _150102| 302655527| 94 0.1 78 10 LeBptaousdp9oh4ianeiariacommapcunCliaaonccsehnl1io39s0bi087so254l4u149s|L99h36e09t2e46r|o8s8t|rophus _150102| Dothistroma61septosporumAs4p5e2rg8i4ll4u0s3nL6ied|pLuteloapsnptoshs6ape7hr5aia1e7rmi2aa3mc7u|arlacnusla1n0s725311|7L|Le 21 22 6 53 Mycosphaerella populorum 453086001| Penicillium chrysogenum 255930451|LLeeppttoosspphhaaeerriaiammaaccuulalannss21301471|L3e|L Zymoseptoria tritici 398412533| 83 Didymella zeae-maydis 403/_g Aspergillus fumigatus 70996344|r Leptosphaeria maculans 10413|L Pseudocercospora fijiensis 452983277| 67 31 Leptosphaeria maculans 6357|Le Didymella zeae-maydis 403/_g 93 Lep5t5oNLspeehpoatfouessMrpiicahaoamccercaor9ucipa3muhlmoapmn3aasc1irnuv8alu6am4np|shL4a1e8s0m5e79o62l7in2|La35410|7L9e26p53to4s7p7h| N34a95eeroiasAamsrptaoecrruyglaailnlfuisssc1th0ee7rrr6ei 71u|1sL91419543795756|45| Leptosphaeria maculans 6357|Le Aspergillus clavatus 121700865| 98 LeptoDspiLdheyampetoreisallapmhzaaeecarueiala-mnmsaay8cd6ui4lsa|Ln1es2m13207/_|Lge8m374 Red1%Aspergillus oryzae 169767282| Cochliobolus heterostrLoepphtoussp4h5a9e3ri8LaeRmpetaodcs1uplahnaser1ia16m4a2c|Lulans 327|Lem Coccidioides posadasii 320036866| Cochliobolus heterostrophus _192271| Leptosphaeria maculans 12272|L C98occidioides posadasii 303317974| 6L7eptosphaeria maculans 12272|L Leptosphaeria maculans 11642|L 98 DidymeClloaczheliaoeb-omluasyhdeiste1r9o5s1tr/o_phus _192271| Coccidioides immitis 119186175| 63 98 Leptosphaeria maculans 525|Lem Figure III.15. The reductase ChRed1 is found only in C. heterostrophus, D. zeae-maydis, and63 Baudoinia compniacensis 449302688| Didymella zeae-maydis 1951/_ 10 Dothistroma septosporum 452844036| Macrophomina phaseolina.52 Methods as in Fig. III.14. A single protein (1210_g374) in D. zeae-maydis clustered with87 Leptosphaeria maculans 1024|Le Leptosphaeria maculans 525|Lem dzm62/_g10 Lep6tospha2e2ri5a3maculanMs y1c0o2s4p|Lheaerella populorum 453086001| gi|1695997 dzm62/_g10 Zymoseptoria tritici 398412533| strong suppgi|4o51r8t58373(ar5r2ow) with C. heterostrophusPgsi|e1u69d5o9c9e7rcospora fijiensis 452983277| Red1 (red), sister to a Macrophomina phaseolina 31protein. A98n additional protein could be foundCochliobolus heterostrophus _N1e3go9i|f94u95s61ic|8o5c3c3um parvum 485922351| in Neofusicoccum parvum sister to Red1, with weak support. See Fig. S16 for full tree.69 31 gi|1891921 55 98 MCaoccrholpiohboomluisnahepthearossetorolipnhau4s0_7193295949767|| 36 75 gi|3309225 69 98 gi|1891921 Didymella zeae-maydis 1210/_g8374 gi|4828078 36 75 gi|3309225 Cochliobolus heterostrophus 45938 Red1 0.1 gi|4828078 Didymella zeae-maydis _1066/_g6390 65 0.1 31 34 42 13 LeDpitdoysmpehlalaerziaeamea-mcualyadniss 1_11501646|/L_g6390 65 Cochliobolus heterostrophus 155544 Red2 Red2,&Red3&Leptosphaeria maculans 11514|L 31 Leptosphaeria mCoachuliaonbsol1u1s5h1e8t|eLrostrophus 155544 Red2 34 42 Cochliobolus heterostrophus 155403 Red3 Leptosphaeria maculans 11518|L 1 29 77 13 Nectria haematococca 302888980| Cochliobolus heterostrophus 155403 Red3 1 Arthrobotrys oligospora 345565698| Nectria haematococca 302888980| Exophiala dermatidis 3787309 Arthrobotrys oligospora 345565698| 29 Coniosporium apollinis 494828354| Exophiala dermatidis 3787309 77 Nectria haematococca 302C8o8n9i5o8s6porium apollinis 494828354| GlareaNleocztoriyaehnaseism3a6t1o1c3o0cc4a 302889586 5 Didymella zeae-maydis 1G91la7r/e_a lozoyensis 3611304 5 84 Figure III.116. The C29hRed2 and Red3 reduc96tase family has members in L. maculansLeptosphaeria maculans 1123D4|idLymella zeae-maydis 1917/_ 84 and D. zeae-maydis. 1 31 29 Cochliobolus het9e6rostrophus _191604| Leptosphaeria maculans 11234|L Cochliobolus heterostrophusC_o1c4h8lio6b9o1l|us heterostrophus _191604| Methods as in Fig.5I8II.14. A single protein in D. zeae-maydis (1066_g6390) and two in10301 Cochliobolus heterostrophus _40416|g Cochliobolus heterostrophus _148691| L. maculans (11514, 11518), clustered with no support with C. heterostrophus Red2 and58 100 Cochliobolus heterostrophus _192351| Cochliobolus heterostrophus _40416|g Red3 (in red). Additional proteins from other fungi clusteredGeomyces destructans 440639301| with them, again withoutCochliobolus heterostrophus _192351| any bootstrap support. See Fig. S16 for84full tree. 69 0 Colletotrichum gloeosporioides 429G8e5o4m16y0ce| s destructans 440639301| 84 Colletotrichum higginsianum 38048C7o4l7le0t|otrichum gloeosporioides 429854160| 69 Ajellomyces capsulatus 154C2o7ll7e0to0t8ri|chum higginsianum 380487470| Clear orthologs with strong bootstrap support can therefore be identified for most Tox10 76Ajellomyces capsulatus 240276986| Ajellomyces capsulatus 154277008| 61 Ajellomyces capsulatus 225554642| 76Ajellomyces capsulatus 240276986| 83 61 genes (PKS1, PKS2, TOX9, DEC1, LAM1) in D. zeae-maydis and L. maculans.Ajellomyces dermatitidis 239614989| 37 83 Ajellomyces capsulatus 225554642| OXI1 and TOX10 A99jellomyces dermatitidis 261187948| Ajellomyces 37 dermatitidis 239614989| 12 Ajellomyces dermatitidis 327357185| A99jellomyces dermatitidis 261187948| Aspergillus clavatus 12117214823 Ajellomyces dermatitidis 327357185| ! 17274 70982839|r Aspergillus clavatus 121714823 97 39 gi|7098283 74 70982839|r 97 19 10 gi|1194840 39 gi|7098283 19 211025329g6i|191| 94840 12 67903818|r 212532969| ! appear unique to these species, but do not have strong bootstrap support, and furthermore, OXI1, appears to be duplicated in D. zeae-maydis. RED1, RED2, and RED3 are more challenging to classify. RED1 appears to be unique to C. heterostrophus and D. zeae-maydis. RED2 and RED3 can both be found in L. maculans (without support for differentiating the two), but with only one ortholog in D. zeae-maydis. Note that an additional reductase is found adjacent to the L. maculans Tox1 genes (see Section C.9 below), however, it fails to group with any other reductase from any species included in this analysis (data not shown). The final tally of genes, therefore, is constant across all species, with a missing RED2/3 replaced with an extra OXI1 in D. zeae-maydis, and a missing RED1 replaced with an extra unrelated reductase in L. maculans. A challenge to our understanding of T-toxin biosynthesis and the corresponding genetics is the PKS1 ortholog in T. stipitatus. It possesses no clear orthologs for any of the Tox1 associated genes beyond PKS1. How, then, does it produce a molecule with T-toxin-like activity? Blast searches can identify protein models with some similarity to Lam1, Oxi1, Red1, Red2, Red3, Tox9, but there is not generally a reciprocal best-hit relationship between the T. stipitatus and C. heterostrophus proteins. Furthermore, none of these genes reside on the same scaffold as the TsPks1 (Ts645921431). Blast searches with Tox10 and Dec1 return no hits. 9. Genomic organization Once C. heterostrophus Tox1 orthologs were identified in D. zeae-maydis and L. maculans, their relative locations within each genome were mapped (Fig. III.17). ! 173 ! L. maculans LAM1 RED2/3 ABC18 TOX10 PKS2 TOOX9XI1 RED2/3 PKS1 DEC1 RED C. heterostrophus race T LAM1 PKS2 8 TOX9 OXI1 TOX10 PKS1 RED2 DEC1 RED3 RED1 83 102 74 82 126 82 D. zeae-maydis TOX10 LAM1 PKS2 TOX9 OXI171 OXI172 RED2/3 PKS1 RED1 DEC1 Transposon T. stipitatus 1284 234 1066 PKS1 1210 scf_1105507295555 Figure III.17. The full C. heterostrophus Tox1 locus can be found in D. zeae-maydis and L. maculans, but not T. stipitatus. Orthology was assigned based first on phylogeny, and then synteny. Gene orientation is marked with small black arrows. The backbone PKS genes PKS1 and PKS2, as well as the additional Tox1 associated genes DEC1, LAM1, OXI1, TOX9, and TOX10 can all be clearly identified in L. maculans and D. zeae-maydis, although gene orientation is not always conserved across species. L. maculans and C. heterostrophus each have three reductases, and D. zeaemaydis has two. RED2 and RED3 orthologs are colored red, RED1 orthologs orange, and unrelated reductases dark red. An additional OXI1-like gene (OXI1-2) is present in D. zeaemaydis, possibly accounting for this missing reductase. T. stipitatus possesses only a PKS1 ortholog: every other Tox1-associated gene is missing. Cartoon not drawn to scale. The most fascinating aspect of the L. maculans Tox1 locus is not that it possesses all known Tox1 genes, but rather that they are in a tight cluster in the genome. All L. maculans Tox1 orthologs identified above, including the PKSs Lm11520 and Lm11513, are present in a ! 174 ! single genomic cluster on scaffold 8 (Fig. III.17), unlike the more complex organization in C. heterostrophus. Genes that are adjacent in C. heterostrophus Tox1 are for the most part, also adjacent in L. maculans although gene orientation is conserved only for LAM1 and PKS2. LAM1 is adjacent to PKS2, TOX9 is adjacent to OXI1, and TOX10 is adjacent to PKS1. In fact, it was collinearity of TOX10 and PKS1 in L. maculans that lead to the serious consideration of TOX10 as more than a misannotation (see above). The RED2 and RED3 related reductases are also present in this cluster, but not adjacent as in C. heterostrophus. In lieu of RED1, a novel and unrelated reductase is present at the end of the L. maculans cluster. Curiously, an ABC transporter, orthologous to an ABC transporter found in both race O and race T of C. heterostrophus resides in the middle of the L. maculans cluster (ABC18, Fig. III.18). The complete inventory of Tox1 genes can be also be found in D. zeae-maydis, on four scaffolds. There are several structural differences. Scaffold 1066, containing the ChPKS1 ortholog, houses two reductases (one of which is a RED2/RED3 related reductase (Fig. III.16), the other related to OXI1 (Fig. III.12), whereas in C. heterostrophus PKS1 is adjacent to TOX10. Scaffold 1284 has PKS2 and LAM1, like its C. heterostrophus counterpart, but, in addition, contains TOX10. Finally, scaffold 1210 has one reductase and one decarboxylase, unlike C. heterostrophus scaffolds 82 and 126, which have 3 reductases and 1 decarboxylase. Note that the contents of Scaffold 1066 have been reported previously as the known MzmTox1 locus [23]. The genes, labeled MzPKS1 (PKS1), MzRed1 (Red2/3) and MzRed2 (OXI1-2), were identified by Sanger sequencing of a 23kb region that also carried 5 transposons. Comparing L. maculans scaffold 8 to C. heterostrophus race O and race T allows us to clearly define the boundaries of the Leptospharia cluster (Fig. III.18). The L. maculans cluster is flanked on one side by a gene encoding an ABC transporter (ortholog of C. heterostrophus ! 175 ! ABC2) and on the other by a gene encoding an ankyrin type protein, which are present in both C. heterostrophus race O and race T. In fact, these two gene calls are adjacent in both C. heterostrophus races. Because these genes occur in syntenic blocks present in C. heterostrophus race O strain C5 and race T strain C4, and because they map to the distal end of chromosome 13 in C5 [79] it does not seem likely that they are associated with the reciprocal breakpoint housing Tox1, or with T-toxin production. If the syntenic presence of the ABC transporter and ankyrin protein encoding gene truly demarcates the boundary of the Tox1 locus in L. maculans, there are exciting implications for C. heterostrophus. A central dilemma to understanding the Tox1 locus is reconciling the predicted 1.2MB of Tox1 unique sequence and the meager amount of race T unique sequence in the physical assemblies. in silico subtraction of race O from race T strains found little additional sequence, suggesting that the vast majority of this 1.2MB was repetitive. One could never be sure, however, that a gene was not overlooked, or misassembled. With the L. maculans Tox1 locus containing every known Tox1 gene in one place, perhaps every Tox1 gene has in fact been identified. ! 176 ! L. maculans ABC2 LAM1 RED2/3 ABC18 TOX10 PKS2 TOOX9XI1 RED2/3 PKS1 DEC1 RED ANK C. heterostrophus race T LAM1 PKS2 8 TOX9 OXI1 TOX10 PKS1 RED2 DEC1 RED3 RED1 83 102 ABC18 74 82 126 82 ABC2 ANK NPS9 9 21 C. heterostrophus race O ABC18 17 ABC2 ANK NPS9 13 Figure III.18. The L. maculans strain JN3 genome contains all known C. heterostrophus Tox1 genes. The C. heterostrophus Tox1 orthologs, identified by blast and phylogenetic analysis against the L. maculans genome sequence, are located as a single cluster at a single genetic locus on L. maculans scaffold 8 (genomic coordinates 377570-466890). Scaffolds are indicated by thin horizontal bars (scaffold number below line) and colored in relation to the C. heterostrophus C4 assembly. Arrows indicate continuation of the scaffold and blunt ends scaffold boundaries. Genes are labeled based on their ortholog in C. heterostrophus race T and colored according to the corresponding C. heterostrophus Tox1 scaffold. Blue-colored genes signify the boundary of the L. maculans Tox1 locus but are found in both C. heterostrophus race T and race O adjacent to each other, not associated with Tox1 genes. ABC18 (white) is found in both race O and race T, and its ortholog in L. maculans bisects the L. maculans Tox1 locus. Unlike L. maculans, C. heterostrophus Tox1 sequence is spread out across 5 separate smaller scaffolds. ! 177 ! 10. TOX10 and ABC18 Two genes in the LmPKS13 LmPKS14 cluster had orthologs in C. heterostrophus that had not been analyzed for involvement with T-toxin production. The first is an ABC transporter, located between LmOXI1 and LmTOX9. There are no ABC transporters on any known C. heterostrophus Tox1 scaffold, but a comprehensive ABC knockout study was carried out previously, and the ortholog of this L. maculans ABC transporter corresponds to C. heterostrophus ABC18. This transporter is found in both race T and race O, in the middle of a large syntenic region between the two races on scaffold 9 in race T strain C4, and scaffold 17 in race O strain C5 (Fig. III.18). The second gene revealed by comparison with the L. maculans cluster is a hypothetical protein located between LmPKS1 and a reductase. This hypothetical protein, which we have named TOX10, is assembled (connected by a gap) on the same scaffold as PKS1 in the newest C. heterostrophus C4 assembly, and was not found in the TMRI or JGI pre-release assemblies. TOX10 is found in race T, but not race O, C. heterostrophus genomes. It can also be found in D. zeae-maydis, but adjacent to LAM1, not PKS1 (Fig. III.17). Both ABC18 and TOX10 were knocked out separately, for characterization of function related to T-toxin biosynthesis. ABC18 was deleted in both C4 (race T) and C5 (race O) strains of C. heterostrophus. TOX10 was disrupted in C4, but not completely deleted, as flanking regions could not be amplified surrounding the gene, due to the limited amount of sequence to work with, and its highly repetitve nature. Race T abc18 mutants are TOX+ as determined by in vitro and in planta assays (Figs. III.19, III.21). Working under the hypothesis that Abc18 could export the synthesized toxin, strains were grown in liquid culture and filtered. Culture filtrate was still found to be TOX+. Additionally, abc18 mutants were inoculated onto T and N corn, and mutants produced the ! 178 ! elongated chlorotic lesions indicative of T-toxin on T cytoplasm corn, like WT. abc18 race O and race T mutants were unaffected on N cytoplasm corn (Fig. III.21). We can thus conclude that ABC18 does not play a role in T-toxin production or export. Figure III.19. C. heterostrophus abc18 mutants produce and export T-toxin. race%T%WT% abc18&1' abc18&2' abc18&4' race%O%WT% T-toxin assay of abc18 mutants. Each row contains agar plugs taken from 5mm inside the tip of colony growth (left), at the tip of colony growth (middle), and 5mm past the tip of colony growth (right). Plugs from race T abc18 mutants and WT race T (C4), clear pATH13 E. coli, even when taken from the agar 5mm past the tip of fungal growth. Plugs of WT race O (strain C5) have no clearing effect. tox10 mutants, on the other hand, are completely Tox-, as determined by our E. coli assay, and an in planta assay on T-cms corn (Figs. III.20, III.21). tox10 is not reduced in ! 179 ! virulence on N cytoplasm corn (Fig. III.22). While Tox10 is essential for T-toxin production, there are very few hints as to its function. The gene product has no bioinformatically predicted functional domains or secretion signals. Attempts to predict the protein structure by threading using the LOMETS server yielded no results. Based on blast searches, TOX10 appears to be unique to C. heterostrophus race T, L. maculans, and D. zeae-maydis. Even between these three species, the proteins have a maximum identity of only 43% with 92% coverage. Very weak orthologs can be found in S. nodorum and S. turcica (41% max identity, 61% coverage). In the phylogenetic analyses, there was no support for orthology with any proteins, including those of L. maculans and D. zeae-maydis. ! 180 ! race%T% race%T% race%O% tox10&2( tox10&3( tox10&4( Figure III.20. C. heterostrophus tox10 mutants do not produce T-toxin. T-toxin assay of tox10 mutants. The control race T strain C4 (top) and hapX-1 mutant race T strain (top left) form halos, but control race O strain C5 (top right) does not. The three independent tox10 deletion mutants do not produce T-toxin (bottom). ! 181 ! WT#race#T# tox10&3( abc18&1( Figure III.21. C. heterostrophus race T WT and abc18 mutants, but not tox10 mutants, produce lesions typical of T-toxin production on T-cytoplasm corn. C. heterostrophus produces long, streaky, grayish chlorotic lesions that quickly expand past the typical lesion size on T-cytoplasm corn (WT C4, left), rapidly resulting in death of the entire leaf. tox10 mutants produce brown necrotic lesions (tox10-3, middle) typical of those seen on Ncytoplasm corn. abc18 mutants (abc18-1, right) still result in streaky T-toxin symptoms on Tcytoplasm corn. ! 182 ! Average'lesion'length'(mm)' 6" 5" 4" 3" 2" 1" 0" WT# tox10)3# Figure 22. tox10 mutants are not reduced in virulence on N-cytoplasm corn. WT race T (C4) and tox10 deletion mutants were inoculated onto W-64A N cytoplasm corn, which are not affected by T-toxin. When average lesion size was determined, tox10 mutants did not differ from WT. ! 183 ! D. Conclusions Having the genome sequences for multiple C. heterostrophus isolates, and an expanded set of distant and close relative genomes to draw from has propelled our understanding of the Tox1 locus forward. Significant progress has been made regarding the location of the Tox1 breakpoints, the set of coding genes required to produce T-toxin, and the phylogenetic distribution of Tox1-associated genes. We were able to identify two candidate breakpoints for the reciprocal translocation that, along with Tox1, genetically define race T relative to race O. One of these was located on a scaffold already linked to chromosome 12, one of the two reciprocally translocated chromosomes. The other is likely located on chromosome 6, but the scaffold it resides on was not linked by an RFLP to the genetic map. Even with multiple sequenced race T isolates, stitching Tox1 fragments into two complete loci (ie Tox1A and Tox1B) remains an unachievable goal. Fortunately, the unified Tox1 locus in L. maculans likely renders this task less critical. All genes found in the L. maculans Tox1 locus have been identified in both C. heterostrophus and D. zeae-maydis, and it is possible that the complete coding inventory of Tox1 has been identified. But what does the tidy L. maculans Tox1 locus say about the origins of T-toxin? The percent similarity and phylogenetic relationship of the individual genes in the locus does not support a direct transfer from L. maculans to C. heterostrophus: Why, then, are the genes clustered only in L. maculans? And is the AT-rich nature of the Tox1 locus connected to the AT-rich isochore paradigm found in the L. maculans genome? If L. maculans truly does not produce T-toxin, as preliminary T-toxin ! 184 ! activity assays suggest, how could it have served as the source of Tox1 for both C. heterostrophus race T and D. zeae-maydis? The discovery that T. stipitatus possesses T-toxin-like activity is startling, and further compounds this mystery. ChPKS1 was the only PKS with a clear ortholog in T. stipitatus. 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Phytopathology 80: 819–823. ! 197 Chapter IV The role of iron acquisition and homeostasis in Cochliobolus heterostrophus cellular biology and pathobiology 198 Abstract Iron is an essential nutrient, and diligent iron acquisition and management are key traits of a successful pathogen. Fungi can produce nonribosomally synthesized iron chelators (siderophores), or Reductive Iron Assimalation (RIA), to acquire iron in a high affinity manner. Previous studies identified two genes, NPS2 and NPS6, encoding different nonribosomal peptide synthetases (NRPS) responsible for biosynthesis of intra- and extra-cellular siderophores, respectively, in the maize pathogen Cochliobolus heterostrophus. Deletion of NPS6 results in loss of extracellular siderophore biosynthesis, attenuated virulence, hypersensitivity to oxidative and iron-depletion stress, and reduced asexual sporulation, while deletion of NPS2 leads to defective sexual spore development. nps2nps6 double deletion mutants are further impaired in all of these processes. Here it is reported that the alternate high affinity iron acquisition system, RIA, is dispensable for C. heterostrophus. ftr1 and fet3 deletion mutants, lacking the high affinity transporter and ferroxidase components, respectively, required for RIA, have no abnormal phenotypes. When combined with nps6 and nps2nps6 mutants, however, basic morphological defects occur. Both nps6ftr1 and nps2nps6ftr1 mutants display reduced growth and asexual production without supplemental iron, and are progressively reduced in virulence compared to nps6 mutants. The iron-responsive transcription factor HapX also plays an important role in iron homeostasis in C. heterostrophus. hapX mutants are sensitive to iron but not oxidative stress, reduced in virulence, and more impaired in sexual development than nps2 mutants. Thus, intra- and extracellular siderophore acquisition and storage, along with RIA, play redundant but vital roles in C. heterostrophus. 199 A. Introduction 1. Why Iron? Iron is essential for almost every organism on Earth, including humans, animals, plants, and their pathogens. Iron-deficiency anemia is one of the most serious human nutritional problems worldwide [1], and iron is often a limiting nutrient in crop production, particularly in calcareous, alkaline soils [2]. Iron is the fourth most abundant element in the earth’s crust, but bio-available iron is limited due to low solubility [3]. The aerobic environment we find on earth keeps iron oxidized in insoluble oxyhydroxide colloid particles [4,5]. This was not always so, and at the infancy of life, water soluble ferrous iron was available. The advent of oxygen in the atmosphere oxidized iron and reduced its bioavailability, causing a shift in biological metal chemistry, and making the quest for iron a central challenge for life [6]. The battle is not over once iron is acquired: unchelated iron can be damaging to cellular function due to generation of reactive oxygen species (ROS) via the Fenton reaction [7,8]. All organisms must therefore take care to chelate and store iron to prevent unintentional cellular damage, and utilize enzymes that prevent or remove oxygen radicals [9]. ROS, in turn, link iron to many biological processes in fungi as well as plants, such as sexual development, signal transduction, and stress responses [10,11]. Taken together, iron demands a high degree of cellular management for its acquisition, storage, and proper deployment, as do the ROS that iron produces, intentional and accidental.! 200 2. Iron in host interactions i. Mammalian systems Given the vital status for iron in biology, it is no surprise that iron (and ROS) plays a key role in host pathogen interactions. The importance of iron in vertebrate-pathogen interactions is well established. In a human host, no completely free iron is presented to an invading pathogen, as it is bound to a variety of proteins, including haemoglobin, ferritin, transferrin, and lactoferrin [4,12]. Patients with hemochromatosis have iron-saturated transferrin [13], rendering them highly susceptible to bacterial pathogens, such as Vibrio vulnificus, which rarely causes infections in healthy individuals [14]. To successfully invade a healthy host, a fungal/bacterial pathogen must possess some way of siphoning iron. One method is to directly steal host heme from transferrin/lactoferrin [4,13,15]. Alternatively, invaders can secrete siderophores, as discussed in Section 4.ii, which strip host chelators of their iron. Host macrophages, in response to these strategies, actively withhold iron from serum upon infection and sequester microbial siderophores [16]. The most extreme adaptation in response to this battle is the spirochete, Borrelia burgdorferi, which has evolved to forego iron entirely, instead substituting manganese in its metal-requiring enzymes [17]. ii. Plant systems Iron’s role in plant-pathogen interactions is less clear cut. Like vertebrates, plant iron is kept chelated to protect from rogue ROS [18]. Ferritins likely play a large role in storing iron and buffering the plant from oxidative stress, especially in plastids [19]. For intercellular transport of iron, and perhaps therefore more relevant to a fungal invader not privy to plastid stored iron, iron may be chelated by smaller molecules, such as citrate or nicotianamide [20,21]. 201 An impressive study of the biotroph Blumeria graminis f. sp.tritici on wheat, corn, barley, oat, sorghum, and millet demonstrated that Fe3+, but not Fe2+, accumulates at infection sites, as visualized by Prussian blue staining [22]. Furthermore, this iron is host, not pathogen, derived [22]. This result was specific to monocots, and was not observed when Arabidopsis was inoculated with B. graminis f. sp. tritici [23]. Furthermore, no such phenotype was reported in subsequent work by the authors with the necrotroph Fusarium graminearum on wheat [24], so it is unclear whether or not this response is specific to (hemi)biotrophic pathogens of monocots, and therefore not pertinent to necrotrophic pathosystems. Conversely, starving Arabidopsis for iron by washing roots with chelator reduces susceptibility to the necrotrophic fungal pathogen Botrytis cinerea. Increased fungal siderophore production suggested this reduction in disease was related to iron starvation, as opposed to another mechanism such as antimicrobial activity [25]. There is not enough evidence to know if C. heterostrophus experiences an iron starved or replete environment in early infection of maize. 3. Reactive Oxygen Species in plant-microbe interactions Given the redox activity of iron, it is important to always consider iron in the context of ROS. ROS have many compelling direct and indirect implications for plant pathogen interactions, and the so-called ‘oxidative burst’ is a hallmark initial response to pathogen challenge [26,27]. Because ROS are cytotoxic, plant-derived ROS, especially H2O2, could serve a direct, antimicrobial function [26,28]. However, the efficacy of a toxic ROS burst depends on the sensitivity of the pathogen [29], and pathogens, especially necrotrophs [30], may evolve a higher tolerance to ROS. ROS are damaging to the plant as well, so the amount and location of produced ROS needs to be carefully regulated [31]. 202 i. ROS as a host defense signal The ‘oxidative burst’ certainly does more than act as an antimicrobial agent. The apoplastic surge of ROS production is rapid, but also transient [29], mediating the transduction of successful recognition of a pathogen to a defense response [27,32]. The oxidative burst is robust, and is produced by the plant in both effector triggered immunity and PAMP triggered immunity [32]. ROS can mediate activation of defense genes, or activate additional defenses, by redox control of transcription factors, or through other signaling components, such as phosphorylation cascades [33,34]. Finally, in addition to acting as an antimicrobial or a signal transducer, ROS can also be involved in fortifying or modifying existing defenses. ROS can cross-link the host cell wall glycoproteins or lignin and suberin precursors, strengthening the cell wall as a physical barrier to invasion [35,36]. Phytoalexins and plant-produced secondary metabolites can also be released, activated, or stimulated by ROS [37]. ii. ROS as necrotrophic weaponry ROS are not necessarily always working in favor of the plant, especially in the case of necrotrophs. B. cinerea, for example, has been shown to increase in virulence in synchrony with the intensity of ROS, contributed both by the host and the fungus [30]. Likewise, NADPH oxidase mutants defective in ROS generation are greatly reduced in virulence [38]. B. cinerea infection is actually suppressed in an HR-deficient A. thaliana strain [39]. These observations have lead to the longstanding hypothesis that B. cinerea benefits from ROS overproduction and subsequent triggering of the HR, surviving the defense and thriving on the dead tissue [40]. But is Botrytis the archetype, or the exception for necrotrophs? 203 Deleting all catalases (which degrade H2O2) in C. heterostrophus increased sensitivity to oxidative stress, but did not affect virulence [41]. Likewise, deletion of C. heterostrophus ChAP1, encoding a redox-regulated transcription factor mediating oxidative stress response, was first reported as having no effect on virulence [42], then more recently reported to reduce virulence [43]. Superoxide dismutase mutants (which are impaired in detoxifying ROS) were not reduced in virulence (Horwitz, unpublished). C. heterostrophus extracellular and intracellular siderophore mutants are sensitive to oxidative stress as well as to low iron bioavailability, and are reduced in virulence [44,45]. This dual in vitro phenotype, coupled with the intimate relationship between ROS and iron, makes understanding the role ROS plays in C. heterostrophus infection unclear. iii. ROS as a fungal developmental signal ROS are vital components in a great variety of signal pathways in fungi [10,46]. NADPH oxidase (NOX) mutants are impaired in ROS generation, and therefore display a variety of interesting developmental phenotypes. Aspergillus nidulans, Podospora anserina, and N. crassa nox mutants are all variously impaired in sexual development: some are female sterile, others blocked in sexual structure development, and others produce spores which do not germinate [47-49]. Hyphal growth, pigmentation, and appressorium formation can also be affected [48-50]. The mutualistic ryegrass symbiont Epichloe festuceae NOX mutant noxA has a particularly striking phenotype, exhibiting abnormal, unrestricted growth in planta, resulting in an overgrowth of fungal biomass and stunted plant growth [51]. 204 4. Fungal Iron Acquisition systems Regardless of the specific roles iron and ROS play in plant-pathogen interactions, a fungal phytopathogen must still acquire iron from their host. Fungi have two known methods for high affinity iron acquisition: reductive iron assimilation (RIA) and siderophore assisted (Fig. IV.1) [3,52]. Note that some yeast species, such as Saccharomyces cerevisiae, Candida albicans, and Cryptococcus neoformans do not biosynthesize siderophores [53-56], but they can, however, utilize siderophores secreted by other microbes [54]. Additionally, the low affinity ferrous iron transporter Fet4 is considered the standard method of iron acquisition under iron replete conditions [57]. Fet4 transports not only ferrous iron, but other metals, such as copper and zinc [9]. Although it is hypothetically possible that phytopathogens directly utilize host iron chelates such as ferritin, citric acid, or nicotianamine, as vertebrate pathogens utilize lactoferrin for example [15], there is no evidence for such a system. 205 High affinity Extracellular siderophores Nps6 O HN NH O NO ON O O R4 N R3 O Fe O OO N R2 R1 Xenosiderophores Reductive iron assimilation Fe3+ Fre1 Fet3 O O R4 N R3 Ftr1 O HN NH O NO ON O Fe O OO N R2 R1 Low affinity transporter Fet4 Fe2+ Nps2 H HO N O NH O O N H HN H3C N O O Fe CH3 O O HO N NO O O N O N H CH3 Intracellular siderophores Figure IV.1. Fungi possess three modes of high affinity iron acquisition. (1) C. heterostrophus produces the extracellular siderophore coprogen via the NRPS Nps6. Coprogen must be secreted, where it chelates iron and is reacquired. (2) Reductive Iron Assimilation (RIA) is a three step process in which extracellular iron is reduced by the ferric reductase Fre1, and then oxidized and transported across the membrane by the ferroxidase Fet3 and the high affinity iron permease Ftr1. (3) Fungi can also utilize iron from siderophores or chelators produced by other organisms by direct uptake of the ligand. Alternatively, the low affinity ferrous transporter Fet4 can directly transport free ferrous iron. In C. heterostrophus, free iron is chelated by the intracellular siderophore ferricrocin, produced by the NRPS Nps2. i. Reductive iron assimilation RIA is a high affinity acquisition process whereby ferric iron is reduced extracellularly by a ferric reductase Fer1, then oxidized by the iron multicopper oxidase Fet3 while being transported across the plasma membrane by the high affinity iron permease Ftr1; a mechanism originally discovered entirely in S. cereviseae [58,59]. Fet3 was originally linked to iron uptake phenotypically [60], and demonstrated to have a copper dependency by the iron-deficiency 206 phenotype of the copper transporter mutant ctr1 [61]. Fet3 is a cell surface ferroxidase with its active site projected outside the cell [62], providing ferric iron for the iron permease Ftr1 [63]. The two proteins are intimately linked: they co-localize to the plasma membrane [63,64] and kinetic data support a model in which ferric iron flows from Fet3 to Ftr1 without entering the bulk solvent pool of iron [65]. While it was originally discovered and characterized in yeast, the Fet3-Ftr1 high affinity uptake system can be found in almost all available fungal genomes [59], although functional evidence suggests some fungi, such as A. nidulans, do not utilize RIA [66]. The ferric reductase Fre1 works physiologically epistatic to Ftr1-Fet3, supplying ferrous iron to Fet3 [59,67,68]. Interestingly, yeast mutants that produce ferric reductases but lack ferroxidases are sensitive to metal and oxygen-based stress [69], suggesting that ferrireduction of iron mobilizes ferrous iron, but ferroxidation protects the cell from oxidation [59]. ii. Siderophores Siderophores can be found in plants, fungi, and bacteria, although plant siderophores are produced by different means [3,70]. In maize, L-methionine is converted to nicotianamine by way of nicotianamine synthases, and then converted to the phytosiderophore dioxymugineic acid and its derivatives [71]. The phytosiderophore is secreted into the soil from the roots, where it forms Fe3+-chelates, and is then taken up again by the yellowstripe (YSL1) transporter, a member of a subfamily of oligopeptide transporters [72,73]. Plants possess a host of YS genes with specialized functions for movement of iron throughout the plant [74], where it is bound by variety of ligands, including small proteins, citrate, and nicotianamine [75]. It is these ligands, and not phytosiderophores, that a fungal pathogen must compete with. Fungal siderophores fall into one of three chemical groups: aryl caps (catecholates and phenolates), carboxylates, and hydroxamates [70,76]. With the exception of rhizoferrin, all 207 fungal siderophores identified so far are hydroxamates [77]. These fungal hydroxamate siderophores are derived from the nonproteinogenic amino acid ornithine and different acyl groups, which form either rhodotorulic acid, fusarinines, coprogens, or ferrichromes [3]. Fungi often produce and excrete one family of siderophore for iron acquisition, and produce and retain a different, internal siderophore. Aspergillus species, for example, excrete triacetylfusarinin and fusarinine C, and store iron in ferrichrome, ferricrocin, and hydroxyferricrocin. Relevant to this work, C. heterostrophus secretes the extracellular siderophore coprogen, and neocoprogen I and II derivatives [44], while storing iron intracellularly as ferrichrome [45]. Coprogens are linear hydroxamates, while ferrichrome is a cyclic hexapeptide comprised of three N5-acyl-N5hydroxyornithines and three amino acids (glycine, serine, and alanine) [3,78]. iii. Nonribosomal peptide synthetases Once the precursor amino acids are synthesized, they can be linked together to form a siderophore. This is done by nonribosomal peptide synthetases (NRPSs). NRPSs are large, multi-enzymatic and multi-domain proteins that can synthesize a wide array of small peptides, including siderophores [79]. NRPSs can incorporate both D and L forms of the 20 ribosomally used amino acids, as well as non-proteinogenic amino acids (ornithine, imino acids, hydroxy acids) [80]. The final peptide can be linear, cyclic, or branched cyclic, and may be chemically modified further [3]. The result is that NRPSs can produce an extraordinarily large set of peptides, which are often of extreme importance to medicine, industry, and agriculture [81,82]. The corresponding genes are only found in fungi and bacteria and are absent from animals and plants [83]. NRPSs sport a set of functional domains, termed a module, that cooperate in the synthesis of the small peptide. A given NRPS can be monomodular or multimodular. Each module consists of three core catalytic domains: an adenylation (A) domain for substrate 208 specificity and activation via adenylation with ATP, a thiolation (T) domain for substrate attachment, and a condensation (C) domain for linking substrates [84]. Each module is responsible for activating a single substrate, although domains can be used multiple times. NRPSs can also utilize a variety of other enzymatic domains which modify the manufacturing of the peptide in some way, but each module typically has an A, T, and C domain. iv. Transport of siderophores In order for extracellular siderophores to acquire iron in the environment for the fungus, they must be efficiently secreted outside the cell, and taken back up once they are laden with iron. There is some evidence that ABC transporters play a role in this process. For example, the U. maydis gene cluster responsible for production of the siderophore ferrichrome A, as defined by the corresponding NRPS SidFA/Fer3, includes the ABC-transporter Fer6 [85]. The C. heterostrophus ABC transporter ABC6 is located near the coprogen producing NRPS, NPS6. abc6 mutants analyzed by HPLC export fewer coprogens in culture broth than do WT strains, and it is assumed that Abc6 is involved in extracellular coprogen transport (Zhang and Turgeon, unpublished). v. Vacuolar storage Iron is being shuttled not only in and out of the fungal cell, but also between compartments within the cell. Although intracellular siderophores are known to play a role in storing this iron, how it is stored and ultimately utilized is unclear. The best-studied system is S. cerevisiae, which does not produce intracellular siderophores, but can use foreign siderophores. Within the S. cerevisae cell, iron is cycled in and out of the vacuole. Cytoplasmic free iron is likely to be ferrous based on the -250mV potential of the cytoplasm [86], consistent with the vacuolar iron importer Ccc1 being specific for divalent metals [87]. Within the vacuole, iron is 209 likely to be stored as hydrated ferric phosphate species [88,89], limiting its mobilization to the rate of reduction, catalyzed by Fre6 [90]. Ferrous iron is then oxidized by Fet5 to ferric iron, and transported across the vacuolar membrane by Fth1 [91]. Note that this process is a complete reprise of the high affinity RIA system, except that it is occurring at the vacuolar membrane. What about fungi that utilize intracellular siderophores? Unfortunately, this question has only barely been addressed, and only with A. fumigatus [92]. Iron excess results in an increase in ferric bound ferricrocin, as well as expression of CccA, the ortholog of the S. cerevisiae vacuolar iron transporter CCC1. Interestingly, inactivation of CccA, but not ferricrocin biosynthesis, decreases resistance to high iron conditions [92] . 6. Iron acquisition/siderophores in plant-microbe biology and basic metabolism Mutant studies in fungi that are animal and plant pathogens, as well as free living, have revealed much about the role iron acquisition and storage plays in fungal biology. Different fungi produce different combinations of extracellular and intracellular siderophores, and researchers generate mutants that are deficient in either production of siderophore precursors (such as L-ornithine N5-oxygenases deficient sidA/sid1 mutants) which affect all siderophore production, or the NRPS responsible for a specific siderophore [3,93]. This makes cross-system comparisons confusing, although not impossible. RIA is more straightforward, as deletion of either FTR1 or FET3 appears to disrupt RIA, owed to their interdependent membrane localization and chemical function [64]. i. Siderophores in fungal pathogens of animals Both intracellular and extracellular siderophores are required for full virulence of the animal pathogen A. fumigatus [93-96]. Deletion of either intracellular, or extracellular, 210 siderophore production reduces virulence in a murine model [96]. Host macrophages responded differently to A. fumigatus wild-type (WT) and sidA siderophore mutants, indicating that not only are siderophores required for full virulence, but, conversely, that macrophages upregulate iron import and retention, as well as defense responses including tumor necrosis factor expression, in response to siderophores [97]. Extracellular siderophores were also required for A. fumigatus virulence in a fly host (sidA, sidD), although intracellular siderophores (sidC) were not [98]. Histoplasma capsulatum also requires siderophores, as demonstrated by reduced growth and attenuated virulence of sid1 (encoding an L-ornithine monooxygenase) mutants in a murine model [99]. There is evidence that H. capsulatum may possess an RIA system, but no functional confirmation [99,100]. A. nidulans sidA (L-ornithine N5-monooxygenase) and sidC (NRPS producing ferrichrome) mutants had severe growth defects, and oxidative stress sensitivity [66]. SidC (ferrichrome) is important for conidial iron content, oxidative stress tolerance, and homothallic sexual development [101]. ii. Siderophores in plant pathogens A positive role for siderophores in fungal pathogens of plant systems was first described in the Turgeon lab. It was found that deletion of NPS6, encoding an NRPS producing an extracellular siderophore in C. heterostrophus, as well as Fusarium graminearum, Cochliobolus miyabeanus, and Alternaria brassicicola, reduced virulence in all systems to their respective hosts (corn, wheat, rice, and Arabidopsis) [44]. The actual siderophore produced is different in each of these pathogens: coprogen in C. heterostrophus, Nα-dimethyl coprogen in A. brassicicola, and TAFC in F. graminearum [3,44]. Deletion of NPS6 affects more than 211 virulence: it also results in hypersensitivity to reactive oxygen species (ROS), and hypersensitivity to iron depletion [44,102]. Intracellular siderophores also play an important role in C. heterostrophus reproductive biology. Deletion of NPS2, responsible for biosynthesis of the intracellular ferrichrome-type siderophore, ferricrocin, led to loss of sexual spore development in homozygous nps2 X nps2 crosses, although pseudothecia were formed [45]. A similar sexual development phenotype was found for homothallic F. graminearum; deletion of the NPS2 ortholog results in defects in both sexual development and in ferricrocin biosynthesis [45]. Neither C. heterostrophus nor F. graminearum nps2 mutants are reduced in virulence [45], but nps2nps6 double mutants are further reduced in virulence compared to nps6 single mutants [103]. F. graminearum also possesses a third siderophore NRPS, NPS1, whose metabolite is unknown, and nps1nps2nps6 triple deletion strains are even more attenuated in virulence than nps2nps6 strains [103]. These results are consistent with findings by an independent group working with the F. graminearum siderophore biosynthesis mutant sid1 (which is functionally nps1nps2nps6) [24]. Epichloë festucae/Neotyphodium festucae synthesizes two siderophores, an intracellular ferricrocin produced by NPS9/SidC [104,105], and the extracellular ferrichrome type siderophore epichloënin A produced by the NRPS SidN [104]. The mutant sidN no longer produces epichloënin A, and consequently is sensitive to oxidative stress, and grows recklessly in planta, stunting host growth [104], a phenotype reminiscent of E. festuceae nox mutants (Section 3.iii). It is interesting to note an exception to this pattern of extracellular versus intracellular siderophores and virulence: Magnaporthe oryzae ssm1 (intracellular ferricrocin) mutants were reduced in virulence [106], but extracellular coprogen mutants were not [94]. This phenotype is 212 owed to the incredibly complex, appressorium-dependent mechanism by which M. oryzae invades the host. It seems likely that siderophore deficient ROS imbalance is the culprit for this phenotype, rather than the more direct role implicated for extracellular siderophores [3]. iii. Pathogens for which siderophores do not play a role in virulence Siderophores are not required for virulence of all fungal pathogens, however. U. maydis sid1 mutants [107], as well as Microbotryum violaceum rhodotorulic acid deficient mutants (Mutant 45) [108], are both fully virulent on their corn and Silene hosts, respectively. The historical precedent of the U. maydis result discouraged researchers from investigating the role of siderophores in fungal-plant interactions for some time [3,107]. As noted in Section 4, the human pathogens C. albicans and C. neoformans do not biosynthesize siderophores [53-56], and therefore do not require them for virulence. They can, however, utilize siderophores secreted by other microbes [54]. The siderophore permease Sit1/Arn1 is not required for virulence in C. neoformans and C. albicans in a mouse model [109,110], but in C. albicans, sit1 mutants are required for invasion of epithelial cells, where iron cannot be acquired from blood [110]. iv. Pathogens in which reductive iron assimilation is essential for virulence RIA was first described as important for virulence in the animal pathogen C. albicans [55]. C. albicans has two FTR1 genes, CaFTR1 and CaFTR2, but only CaFTR1 is required for growth in iron-limiting conditions and successful systemic invasion of mice. CaFTR2, curiously, displayed an expression pattern inverse to CaFTR1, being induced under iron replete conditions and repressed under iron-limiting conditions [55]. C. albicans appears to rely on RIA to acquire iron bound to host transferrin in serum [111], but it also utilizes the siderophore transporter system to acquire iron from epithelial cells where serum is not available [110]. C. 213 neoformans also possesses two FTR1 orthologs, one of which is required for full virulence in mice and iron acquisition [53]. Deletion of the FET3 ferroxidase ortholog CFO1 also reduced virulence on mice [112]. Partial disruption of FTR1 in the opportunistic human pathogen Rhizopus oryzae reduces virulence in diabetic ketoacidosis mice (which have elevated iron serum levels, facilitating infection) [113]. R. oryzae does, in fact, produce the extracellular siderophore rhizoferrin, but it is insufficient to acquire iron from serum [114]. The first plant pathogen demonstrated to require RIA for virulence was U. maydis [85]. Fer2, encoding a high affinity permease orthologous to Ftr1, and Fer1, encoding a multicopper oxidase orthologous to Fet3, are clustered and iron regulated, and mutant strains lacking either are strongly affected in virulence [85]. Colletotrichum graminicola also requires RIA for full virulence on maize [115]. C. graminicola possesses two FET3 orthologs, Fet3-1 and Fet3-2; corresponding single mutants are reduced in virulence. This is due in large part to impaired appressorium formation, as single mutants are restored when plants are wounded prior to infection. Double fet3-1/fet3-2 mutants, however, are reduced in virulence on wounded leaves, implicating a direct role in virulence [115]. v. Pathogens that do not require reductive iron assimilation for virulence F. graminearum fgtr1 fgtr2 double mutants were like WT for both growth phenotype and virulence, although either of the corresponding genes could complement growth of yeast ftr1 mutants [116]. fgtr1fgtr2 RIA mutant strains had increased SIDA expression comparable to that seen in iron-depleted WT strains, however, suggesting RIA is utilized to acquire iron in F. graminearum [116]. Similarly, deletion of FtrA in A. fumigatus had no effect on virulence in mice [95]. sidAftrA double deletion strains were unable to grow on most media, including blood 214 agar plates, without ferricrocin supplementation,, demonstrating that A. fumigatus has no machinery for direct utilization of host iron chelates [95]. With the exception of Aspergillus nidulans [66,117], RIA genes are present in all fungal genomes examined [59]. All pathogens, therefore, favor one system, siderophore assisted or RIA, over the other and there is no fungal pathosystem that does not require either RIA or siderophore production for virulence. What determines this choice is unknown, although it is intriguing that, thus far, all necrotrophs rely on siderophores, while biotrophs use RIA. In the only case where double mutants have been made [sidAftrA in A. fumigatus [95]], growth was severely limited. Note sidA eliminates intracellular as well as extracellular siderophore biosynthesis. sidAftrA mutants can therefore be thought of as triple mutants, with the difference between extracellular/intracellular siderophore and ftrA double mutants unexplored. No RIA/siderophore acquisition mutant has been tested without also disrupting intracellular siderophore production. 7. Iron regulation It is clear that proper regulation of iron genes is of the utmost importance for all organisms, and even more so for pathogens. The regulatory machinery for transcription of iron related genes has been well studied in several model organisms, particularly S. cerevisiae and Aspergillus species [9,118]. i. Iron regulation in S. cerevisiae In S. cerevisiae, transcriptional responses to iron starvation depend on Aft1 [9,119]. Aft1 is the master low-iron sensing transcription factor, acting directly and in coordination with other transcription factors to fine tune gene expression in response to a variety of inputs [9]. Deleting 215 AFT1 prevents expression of iron-related genes, a dominant aft1 mutant leads to constitutive overexpression of iron genes, and Aft1 binds directly to promoter regions of iron regulated genes [119-121]. There is an AFT1 paralog, AFT2, of unclear function: aft2 mutants on their own are like WT, but aft1;aft2 double mutants are extra sensitive to iron-limited growth [122]. Together, the Aft1/Aft2 regulon consists of at least 26 genes, all partially characterized and pertaining to iron biology [9]. This includes the RIA encoding genes FET3 and FTR1, their vacuolar equivalents FET5 and FTH1, and the siderophore transporters ARN1-ARN4 [9]. There are exceptions to the dictatorship of Aft1: Yap5, for example, activates expression of the vacuolar iron transporter CCC1 [123]. ii. Regulation beyond S. cerevisiae While understanding the AFT1 expression system is important, it does not inform our understanding of filamentous fungi, as AFT1 is not found outside the Saccharomycotina [3]. The next best-studied fungi are A. fumigatus and A. nidulans, which do share many orthologs with C. heterostrophus. In A. fumigatus, and most filamentous fungi, iron regulation revolves around the feed-back loop of two transcription factors: the GATA-factor SreA and the bZip-factor HapX [117,124,125]. SreA and HapX, in turn, are not found in S. cerevisiae [3,9]. SreA was first shown to regulate siderophore biosynthesis and iron uptake in A. nidulans [126,127], and later to function analogously in A. fumigatus [124]. HapX and SreA are not the final word in iron regulation: other regulatory factors influence expression based on the status of related factors, such as oxidative stress via Yap1 and hypoxia via SrbA [128,129]. During iron sufficiency, SreA is upregulated and HapX is downregulated. SreA then represses expression of RIA and siderophore-mediated iron uptake genes by binding to the consensus sequence ATCWGATAA [12,124,130]. During iron starvation, HapX is upregulated 216 and SreA is downregulated, which activates expression of RIA and siderophore-mediated iron uptake genes. Additionally, HapX rations iron utilization by repressing iron-consuming pathways, such as heme biosynthesis, respiration, and ribosome synthesis [124,125]. HapX and SreA, then, directly oppose one another in a manner which ensures iron acquisition and storage genes are expressed at a level appropriate to the availability of iron [12]. Deletion of either SreA or HapX impacts cellular iron homeostasis, and results in a reddish hyphal pigmentation in Aspergillus [124,125]. Furthermore, attempts to generate hapX sreA double mutants in both A. nidulans and A. fumigatus have failed, as the double mutant is synthetically lethal [124,125,131]. While both regulators are clearly essential for iron regulation, only deletion of HapX, and not SreA, reduces virulence of A. fumigatus in mice [124,125]. hapX mutants are also reduced in virulence in the mammalian pathogens C. albicans and C. neoformans [130,132], as well as the plant pathogen F. oxysporum [133]. Likewise, the SreA ortholog Sfu1 is not required for systemic virulence of C. albicans [134]. C. heterostrophus possesses both HapX and Sre1 orthologs, although only Sre1 has thus far been studied [43]. Our investigation of C. heterostrophus iron metabolism, therefore, benefits from our knowledge of iron acquisition and homeostasis in many other systems. The aim of this chapter is to investigate how unexplored components of C. heterostrophus iron biology (particularly RIA and HapX) impact the fungus and its virulence. 217 B. Materials and Methods 1. Fungal strains and culture conditions Minimal medium (MM), used for stress plates, contained 10 ml of each of two 100x salts solutions (Solutions A and B), 10ml of Srb’s micronutrient stock solution [135] 10 g glucose, 20g agar (for solid plates), and deionized water to 1 litre [136]. Stock salt solution A contained 10g Ca(NO3)2!4H2O and deionized water to 100ml. Stock salt solution B contained 2.0g KH2PO4, 2.5g MgSO4!7H2O, 1.5g NaCl and deionized water to 100 ml (pH 5.3). Complete medium (CM) was MM plus 1g yeast extract, 0.5g acid-hydrolysed casein and 0.5g enzymically hydrolysed casein, per liter. Complete medium no salts (CMNS), used for Hygromycin B selection, was CM without salt solution A or B. Complete Medium Xylose (CMX) was CM with with xylose instead of dextrose [137]. CMX+Fe and MM+Fe were CMX or MM with 100uM ferric citrate (Fischer) added prior to autoclaving, unless stated otherwise. Sach’s medium was used for crosses, and contained 1g Ca(NO3)2!4H2O, 0.19g K2HPO4, 0.25g MgSO4, 0.85g CaCO3, in 1 liter water, vacuum filtered through Whatman #1 paper, and mixed with 20g agar [136]. C. heterostrophus WT race T strain C4 (Tox1+;MAT1-2, American Type Culture Collection [ATCC] number 48331) was used for all initial transformations and strain C2 (Tox1+;MAT1-1;alb1, ATCC 48329) was used as an albino tester for initial crosses. All strains used in this study, including transformants, are listed in Table IV.1. Strains were stored at -80oC in liquid CM [136] containing 25% glycerol and were plated onto CMX for growth and optimal conidiation. Fungi were grown at 24 °C under fluorescent lights in an alternating 16 hours light 8 hours dark cycle under fluorescent light (Watt-Miser F34 WW/RS/WM, Warm White, General Electric). 218 2. Iron Iron source supplementation was tested on solid MM with ferric citrate (Sigma), ferrous sulphate 7·H2O (Mallinckrodt), and ferric chloride 6·H2O (Fluka). Iron was added to media after autoclaving from aliquots of 1mM sterilized stock solutions. Cultures were grown for six days in the dark and photographed. For liquid culture observation, spore suspensions were prepared by scraping culture plates into liquid medium and filtering through sterile cheesecloth. 105 conidia were added to 10ml liquid CM, with or without 100uM ferric citrate. Cultures were grown for 18 hours at room temperature shaken at 150rpm, and observed under a Leica DM5500 Epifluorescence microscope using Differential Interference Contrast (DIC) at the Boyce Thompson Institute Plant Cell Imaging Center, and images were taken using Qcapture Pro 6.0. Table IV.1. C. heterostrophus strains used in this study. Straina Genotype Comments and/or References C4 (ATCC 48331) Tox1;MAT1-2 Race T, inbred laboratory strain, WT [136] C2 Tox1;MAT1-1 Race T, inbred laboratory strain, WT [136] C9 Tox1;MAT1-1 Race T, inbred laboratory strain, WT [136] CB7 Tox1;MAT1-1;alb1 Race T, inbred laboratory strain, WT [136] ChDsred RFP;hygB;MAT1-2 Dsred [138] expressed under the citrate lyase promoter (pIGREDPAPA) Turgeon unpublished Chnps2-1 nps2;hygB;MAT1-2 [45] Chnps2-2 nps2;hygB;MAT1-2 [45] Ch1449-T1-1 nps2;hygB;MAT1-2 [45] Ch1449-T1-5 nps2;hygB;MAT1-1;alb1 [45] Chnps6-1 nps6;hygB;MAT1-2 [44] Chnps6-1-R2 nps6;hygB;MAT1-1 [44] Chftr1-1 ftr1;hygB;MAT1-2 This study, iron permease FTR1 deletion Chftr1-2 ftr1;hygB;MAT1-2 as above Ch1721-R1 ftr1;hygB;MAT1-1;alb1 This study, progeny of cross Chftr1-1 X C2 Ch1721-R3 ftr1;hygB;MAT1-2;alb1 as above Chfet3-1 fet3; hygB; MAT1-2 This study, FET3 ferroxidase deletion Chfer3-2 fet3; hygB; MAT1-2 as above Chfet3ftr1-1 fet3;ftr1;hygB;MAT1-2 This study, deletion of both FET3 and FTR1 Chfet3ftr1-2 fet3;ftr1;hygB;MAT1-2 as above Ch1495-T1-1 nps2;nps6;hygB;MAT1-2 [103] Ch1495-T1-6 nps2;nps6;hygB;MAT1-1 [103] 219 Ch1495-T1-8 nps2;nps6;hygB;MAT1-2 [103] 1496-T1-1 nps2;nps6;hygB;MAT1-1 as above 1496-T1-8 nps2;nps6;hygB;MAT1-1 as above 1722-R1 nps6;ftr1;hygB;MAT1-1 This study, progeny of cross Chnps6-R2 X Chftr1-1 1722-R3 nps6;ftr1;hygB;MAT1-2 as above 1723-T2-2 nps2;ftr1;hygB;alb1 This study, progeny of cross Chftr1-1 and 1449-T1-5 1723-T2-4 nps2;ftr1;hygB as above 1731-T6-3 nps2;nps6;ftr1;hygB; MAT1-1 This study, progeny of 1722-R-1 X Chnps2-1 1743-T6-1 nps2;nps6;ftr1;hygB; MAT1-2 This study, progeny of 1721-R-3 X 1495-T1-6 1744-T6-6 nps2;nps6;ftr1;hygB, alb1 Repeat of above cross 1754-R6 nps6;fer1;hygB This study, progeny of Chnps6-R1 X Chfer1 1754-R7 nps6;fer1;hygB as above 1755-R11 nps6;fer1;ftr1;hygB This study, progeny of Chnps6-R1 X Chfer1ftr1 1755-R18 nps6;fer1;ftr1;hygB as above 1761-R7 RFP;hygB;MAT1-1 This study, progeny of Dsred X C2 1761-R9 RFP;hygB;MAT1-1 as above 1762-T1-1 RFP;nps6;hygB This study, progeny of Chnps6 -1 X 1761-R7 1763-T3-1 RFP;nps6;hygB This study, progeny of Chnps6-1 X 1761-R9 1764-T1-1 RFP;nps6;ftr1;hygB This study, progeny of 1722-R3 X 1761-R7 1765-T2-3 RFP;nps2;nps6;hygB This study, progeny of 1495-T1-1 X 1761-R7 1765-T3-5 RFP;nps2;nps6;hygB as above 1766-R2 RFP;nps2;nps6;ftr1;hygB This study, progeny of 1743-T6-1 X 1761-R7 Chabc6-1 abc6;hygB Zhang and Turgeon, unpublished Chabc6-1nps6-1 abc6;nps6;hygB Zhang and Turgeon, unpublished ChhapX-1 hapx;MAT1-2 This study ChhapX-3 hapx;MAT1-2 This study 1749-R17 sre1;MAT1-1 sre1 mutant, [43] aNomenclature: Chnps2-1 is a strain in which NPS2 is deleted, ‘-1’ indicates transformant #1; 1449-T1-1 is cross #1449; T1-1 is tetrad #1, ascospore 1. 1766-R2 is the second random progeny recovered (rather than part of a tetrad) from cross #1766. Strains are pigmented unless denoted alb1 in genotype, for albino strains. If mating type is not included, it was not tested and is unknown. 3. Plant cultivars and growth conditions Corn cultivar W64A with N cytoplasm was used in this study. Corn was grown in a growth chamber under 16 h light/ 8 h dark at 24 °C. Seed was planted 3 plants per #6 standard pot, in Cornell Mix soil. Plants were inoculated as described below at three weeks, after emergence of the fourth true leaf. 4. DNA manipulations and fungal transformations Fungal genomic DNA was prepared using the Ultraclean Microbial DNA isolation kit (MO BIO). PCR reactions were carried out with Phusion (Finnzyme) DNA polymerase and mix 220 for generating transformation constructs, or GoTaq (Promega) for screening mutants, following the manufacturer’s recommendations. Transformations were performed using split marker-based homologous integration [139,140]. 700-1000bp DNA flanking (5’ and 3’ flanks) the coding sequence was amplified using primer pairs with extensions complementary to M13Rhyg (TCCTGTGTGAAATTGTTATCCGCT-XXXXXXX on the reverse primer of the 5’ flanking region) and M13Fhyg (GTCGTGACTGGGAAAACCCTGGCG-xxxxxxxx on the forward primer of the 3’ flanking region). 5’ and 3’ flanks were then amplified, along with overlapping segments of the gene for resitance to Hygromycin B (5’ region “HY” amplified with M13RHYG and NLC37, 3’ region “YG” amplified with M13FHYG and NLC38) from the vector pUCATPH, which contains the HygB gene between the A. nidulans PtrpC promoter and TtrpC terminator. A second round of PCR, using the 5’ flank forward primer with NLC37, and the 3’flank reverse primer with NLC38, results in the 5’ flank being fused to the “HY” hygromycin B construct, and the 3’ flank fused to the “YG” component. To prepare protoplasts for transformation, 15x100 mm Petri dishes with 10 day old strain C4 culture on CMX were scraped using sterile wooden dowels, filtered through sterile cheesecloth, and inoculated into 2x 100mL liquid CM cultures overnight (18h) at 24° C. Two 40ml centrifuge tubes were filled with liquid culture and pelleted in a Sorvall RC-5C centrifuge at 5000 rpm for 5 minutes at 4°. One scoop (10mm diameter glob) each of pelleted mycelium was transferred to 8 50ml sterile flasks, each containing 10mL enzyme osmoticum [3.27 g NaCl (0.7 M), 1.6 mL Glucanex (Novo Nordisk), 0.8 g Driselase (Sigma), H2O to 80 mL, filter sterilized] and shaken gently (40rpm) 30° for 3 hours to release protoplasts. Protoplasts were then filtered through sterile cheesecloth and nylon membrane (SEFAR) (25um pore size) and 221 pelleted by centrifugation at 5000 rpm for 5 minutes at 4°C. The pelleted protoplasts were suspended and re-pelleted four times: the first time in 2x 5ml 0.7M NaCl solution, and then three additional times in 10ml STC [sorbitol, 21.86 g (1.2 M), 1 mL of 1 M Tris–HCl pH 7.5, (10 mM), 0.735 g CaCl2·2H2O (50 mM), H2O to 100 mL] solution. After washing, protoplasts were resuspended in 200uL STC, adjusted to approximately 1*107/ml and kept on ice. Protoplasts were transformed by adding 20ul transformation construct (DNA concentration of PCR product not determined) to 100uL protoplast suspended in STC, and held on ice 5 minutes. 200uL, 200uL, and 800uL polyethylene glycol solution [30 g polyethylene glycol, MW 3,350 (60% w/v), 0.5 mL of 1 M Tris–HCl pH 7.5 (10 mM), 0.37 g CaCl2·2H2O (50 mM), H2O to 50 mL] was added, mixing gently and incubating 5 minutes between aliquots. Finally, 1mL STC was added, and protoplasts were dispensed 300ul at a time into 20ml molten recovery medium [consisting of three flasks, A, B, and C, autoclaved separately, mixed, and held in a water bath prior to use. Flask A: 1 g yeast extract, 1 g casein hydrolysate (enzymatic), H2O to 50 mL. Flask B: 342 g sucrose, H2O to 500 mL. Flask C: 16 g agar, H2O to 450 mL.]. Protoplasts were mixed into the molten recovery medium and allowed to solidify overnight at 30° C. Then 150ug Hygromycin B /mL in 1% agar was overlayed, and transformants were allowed one week to emerge at 30° in the dark. Candidate transformants recovered from the hygromycin B overlay were transferred to selective medium (CMNS containing 150ug Hygromycin B/ml) to confirm resistance. Conidia were streaked on 5% water agar to separate individual conidia, and individual germinating conidia were transferred to CMX. Only a single nucleus enters a given conidium: single conidiation therefore removes heterokaryons. Candidates were then patched onto CMNS HygB (150ug Hygromycin B/ml) and CMX to confirm resistance and for storage in glycerol, 222 respectively. DNA was prepared from candidate transformants and subjected to diagnostic PCR (Fig S2, Table IV.2) to verify gene deletion. DNA from candidates with a gene of interest deleted and replaced with the selectable marker carried a band of predictable size when a primer external to the 5’ or 3’ flanking region was combined with NLC37 and NLC38 (internal to HygB), respectively, and if a band was not amplified using primer pairs internal to the gene targeted for knockout (see Fig. S2 in Appendix for PCR transformation construct generation and verification schematic). All gene deletion and verification primers are listed in Table IV.2. Table IV.2. Primers used in this study. Primer Sequence (5’->3’) Chftrupfa cagataaatggcaccagact Chftrupr Chftrdwnf Chftrdwnr TCCTGTGTGAAATTGTTATCCGCTgtg catcccgtctctgta GTCGTGACTGGGAAAACCCTGGCGat gtattgggaggcctaga cctgtgttccatccttttac Chftrinf ggcgtgataggaacctttt Chftrinr tttgctcaagactccagaca fet3upf caaacacgagttgctgtacg fet3upr fet3dnf fet3dnr TCCTGTGTGAAATTGTTATCCGCTtcc taggtgcaagtgaatga GTCGTGACTGGGAAAACCCTGGCGa gctgtactggatggtcctg atcccgtctctgtaccatga fet3inf cgttgaagaaggcactttgt fet3inr ttgttgaagtcgatggtgtg fet3dwn2f fet3dwn2r GTCGTGACTGGGAAAACCCTGGCGct agaagacacagccccatc gacaccatgtatggggaaag fet3extupf gcatgaagccatcacaaagt fet3extdnr fet3extfardnr ggcgcagatgattagacaga aggggagtggttatgattgc Purpose (See Fig. S2) Upstream forward, for deletion of FTR1 (ID: 104817) Upstream reverse, for deletion of FTR1 (ID: 104817) Downstream forward, for deletion of FTR1 (ID: 104817) Downstream reverse, for deletion of FTR1 (ID: 104817) Internal to FTR1 (ID: 104817) for deletion verification Internal to FTR1 (ID: 104817) for deletion verification Upstream forward, for deletion of FET3 (ID: 104814) Upstream reverse, for deletion of FET3 (ID: 104814) Downstream forward, for deletion of FET3 (ID: 104814) Downstream reverse, for deletion of FET3 (ID: 104814) Internal to FET3 (ID: 104814) for deletion verification Internal to FET3 (ID: 104814) for deletion verification Downstream forward, for deletion of FTR1 and FET3 (IDs:104817, 104814) Downstream reverse, for deletion of FTR1 and FET3 (IDs:104817, 104814) Upstream external, for verification of FET3 and FTR1/FET3 (IDs:104817, 104814) deletion with NLC37 Downstream external, for verification of FET3 (ID: 104814) deletion with NLC38 Downstream external, for verification of FTR1/FET3 (IDs:104817, 104814) deletion with NLC38 223 Fet4UpF caactagctctggcgtttg Upstream forward, for deletion of FET4 (ID: 33695) Fet4UpR TCCTGTGTGAAATTGTTATCCGCTgat Upstream reverse, for deletion of FET4 gctactggagcgtctg (ID: 33695) Fet4DnF GTCGTGACTGGGAAAACCCTGGCGc Downstream forward, for deletion of gatcaagaatggcgtaag FET4 (ID: 33695) Fet4DnR ccaattcgatagggcataa Downstream reverse, for deletion of FET4 (ID: 33695) Fet4Upext ggtgcaaaagaaaagcaca Upstream external, for verification of FET4 (ID: 33695) deletion with NLC37 Fet4Dnext cggtacaagaacagaacgtg Downstream external, for verification of FET4 (ID: 33695) deletion with NLC38 Fet4inF atcttcacgcctcttttcc Internal to FET4 (ID: 33695) for deletion verification Fet4inR gtggctcctcattggtaca Internal to FET4 (ID: 33695) for deletion verification hapxupf acctacctcgcataccaagc Upstream forward, for deletion of HAPX (ID: 1021239) hapxupr TCCTGTGTGAAATTGTTATCCGCTagc Upstream reverse, for deletion of HAPX attgtggtggatggtta (ID: 1021239) hapxdnf GTCGTGACTGGGAAAACCCTGGCctat Downstream forward, for deletion of agggtggcgttcgtct HAPX (ID: 1021239) hapxdnr aggacccaaggcaaacatac Downstream reverse, for deletion of HAPX (ID: 1021239) hapxupext ccccaactgatcactccaa Upstream external, for verification of HAPX (ID: 1021239) deletion with NLC37 hapxdnext ttagcccctctgatgaaggt Downstream external, for verification of HAPX (ID: 1021239) deletion with NLC38 NPS6F1 GCCTGAGCTTCGCTTCTGTAT Internal primer for NPS6 (ID: 128080) NPS6R1 GCAACTTTGCCAGTTTGTCAG Internal primer for NPS6 (ID: 128080) NPS2F1 CAGTTTTCAACGTCCGATTCA Internal primer for NPS2 (ID: 128084) NPS2R1 GCCTCCTATTGCAAGTTCACC Internal primer for NPS2 (ID: 128084) NLC37 GGATGCCTCCGCTCGAAGTA pUCAPTH SEQ. 1685-1702 NLC38 CGTTGCAAGACCTGCCTGAA pUCATPH SEQ. 2132-2150RC M13RHYG AGCGGATAACAATTTCACACAGGA pUCATPH SEQ. 2865-2888RC M13FHYG CGCCAGGGTTTTCCCAGTCACGAC pUCATPH SEQ. 352-375 aNomenclature for knockout and verification primers: Chftrupf: Ch = C. heterostrophus, ftr = gene (FTR1) up = upstream flank, f = forward primer. See Fig. S2 in the Appendix for knockout and verification schematic. 5. Construction of polymutant strains A list of crosses set up for recovery of mutants with double and triple gene deletions is available in Table IV.3. Transformants of strain C4 (MAT1-2) were initially crossed to strain C2 (MAT1-1) to generate albino and pigmented strains of both mating type. All WT genes were replaced with the hygB gene, conferring resistance to hygromycin B. Tracking hygromycin B resistance, therefore, also tracks gene deletion (assuming Hygromycin B has correctly replaced 224 the targeted gene, as confirmed by diagnostic PCR of candidate transformants, see Fig. S2). Single and polymutants were recovered from crosses by collecting random hygromycin B resistant progeny, or by tetrad analysis. For tetrads, the full set of progeny (tetrad) from a given ascus was collected, in isolation from other asci to ensure each ascospore was from the same ascus. Meiosis occurs inside each ascus, and so the complete meiotic set (four pairs of twins) of ascospores will be found in each ascus. In crosses between hygromycin resistant parents, if hygromycin B sensitive progeny are observed in a tetrad, the tetrad must contain contain WT progeny, and by extension the polymutant containing each mutation carried by both parents. Diagnostic PCR reactions with the same primer pairs used to confirm successful gene deletions were used to verify the genotype of polymutants collected by both random and tetrad isolated ascospores (see Fig. S2). Table IV.3. Crosses and selfs set up in this study. Cross Parent strains WT X WT C4 X C2 ftr1 X WT Chftr1-1 X C2 ftr1 X ftr1 Chftr1-1 X 1721-R2 ftr1 X nps6 Chtr1-1 X Chnps6-1-R2 ftr1 X nps2 Chftr1-1 X 1449-T1-5 fet3 X WT Chfet3-1 X C2 fet3ftr1 X WT Chfet3ftr1-1 X C2 ftr1nps6 X nps2 1722-R-1 X Chnps2-1 ftr1 X nps2nps6 1721-R-3 X 1495-T1-6 fet3 X nps6 Chfet3-1 X Chnps6-R2 fet3ftr1 X nps6 Chfet3ftr1-1 X Chnps6-R2 fet4 X nps2nps6ftr1 Chfet4-1 X 1731-T6-3 Dsred X WT ChDsred X C2 Dsred X nps6 Chnps6 -1 X 1761-R7 Dsred X nps2nps6 1495-T1-1 X 1761-R7 Dsred X nps6ftr1 1722-R3 X 1761-R7 Dsred X nps2nps6ftr1 1743-T6-1 X 1761-R7 hapX X sre1 hapX-1 X 1749-R17 hapX X sre1 hapX-3 X 1749-R17 hapX X WT hapX-1 X C2 hapX X WT hapX-1 X C9 hapX X WT hapX-1 X CB7 a For strain designations, see Table IV.1 225 6. Asexual development Asexual sporulation of WT and nps6, ftr1, nps6ftr1, nps2ftr1, nps2nps6, nps2nps6ftr1, and nps2nps6ftr1fet4 mutant strains was quantified. Strains were grown on CMX with and without exogenous application of iron (100 uM ferric citrate), for seven days at 24 °C under fluorescent lights with 16 hours light 8 hours dark alternating fluorescent light and photographed. Plates were then completely scraped with 0.2% Tween 20 in H2O to collect conidia, filtered through sterile cheesecloth, and suspended in 10 ml 0.2% Tween 20. The concentration was determined by counting spore suspensions under a stereomicroscope with a haemacytometer, and the number of spores per plate estimated for each plate. 7. Feeding plates nps2nps6ftr1 was grown for 5 days on CMX+Fe. Small culture scrapings (as small as possible, 1mm X 1mm) were transferred to CMNSHygB plates. Care was taken to collect mycelium and conidia only, without any iron-rich medium. For supplementation with a WT culture, the nps2nps6ftr1 triple mutant was set up in a grid pattern, and a patch of WT mycelium was added to a corner outside the grid. When supplementing with any other culture, culture filtrate was used instead of a solid piece of mycelium to prevent the supplementing culture from overgrowing the nps2nps6ftr1 mutant. For this, 50ml cultures were inoculated with 5x5mm scraped conidia and shaken overnight. Cultures were centrifuged for 5 minutes at 10,000RPM at room temperature in a tabletop centrifuge (HERMLE) to pellet mycelium. 20ul supernatant was transferred to wells carved into the agar with a cork borer, set up in corners outside the grid of nps2nps6ftr1 mutants. Plates were grown for two (nps2nps6), three (abc6nps6) or five days (hapX) at 24 °C under 16 hours light 8 hours dark alternating fluorescent light and photographed. 226 8. Stress sensitivity assays Sensitivity to H2O2, the superoxide-generator KO2, the membrane-permeable iron chelator, 2-2’-dipyridyl (2DP), and the membrane-impermable iron chelator bathophenanthroline disulfonate (BPS) was assessed. A fresh stock solution of each stress agent was prepared for each experiment (1M KO2, 10 mM 2DP, and 100 mM BPS solutions in water) and the stress agents were added to MM after autoclaving and cooling to approximately 48° C. Fresh MM plates with the stress agents were prepared for each experiment. All experiments were carried out in the dark at room temperature, and allowed to grow for six days before being photographed. Colony diameters were measured from leading tip to leading tip of hyphal growth; the extent of growth within that diameter was not considered. 9. Virulence Virulence assays of C. heterostrophus on maize were carried out as described previously [44]. At least three replicates (i.e. inoculation to three independent plants) were set up for each strain, and experiments were repeated at least three times in all cases. All fungal strains were recovered from -80o C glycerol stocks on CMX with 100uM supplemental ferric citrate (if experiment included nps6ftr1 or nps2nps6ftr1, otherwise, plain CMX was used) for inoculation. After strains were grown for 10-14 days, conidia were harvested by scraping plates in 0.02% Tween 20, and filtered through cheesecloth to remove hyphal debris. Conidia were counted with a haemocytometer and adjusted with 0.02% Tween 20 to 2x104 conidia/ml. Conidia were spray inoculated onto maize plants using Preval sprayers, 2ml/plant. Infection took place overnight in a mist chamber, after which plants were moved to 227 the growth chamber. 3rd or 4th true leaves were collected after 5 days and photographed. At least 50 independent lesions were measured for each strain for each experiment. For iron supplementation, maize leaves were rubbed with 1 mM ferric citrate solution containing 1g/L glucopon 215 CS UP (Fluka), one hour prior to inoculation. The iron-treated plants were incubated in a growth chamber under continuous light until inoculation. Fungal inoculation was carried out in the same way as the standard inoculation assay, except that spore suspensions with/without 0.5 mM ferric citrate were prepared. Control plants inoculated with 0.5 mM ferric citrate without fungal spores were set up as a control. A fresh stock solution of ferric citrate (10 mM in water) was prepared for each experiment. 10. DAB staining and confocal microscopy Inoculation and growth conditions for confocal microscopy and DAB staining were the same as above, except fungal spores were as concentrated as possible- one entire 100x15 petri dish per inoculation was scraped into 50ml 0.02% Tween 20 in H2O, and the entire 50ml was used in the inoculation. For DAB staining, leaf squares were submerged in DAB solution (1mg/mL, pH 3.7) and shaken for 4 hours at 50rpm, then transferred to 3:1 ethanol:acetic acid, and shaken overnight at 50rpm to clear leaves. Prior to confocal microscopy, fungal mutants in this experiment were first crossed to a red fluorescent Dsred strain [138], previously transformed using the pIGREDPAPA vector (Liu, Oide, and Turgeon, unpublished) for visualization. Tetrads were collected for nps6, nps2nps6, and nps6ftr1 X Dsred crosses; random progeny were collected for WT X Dsred and nps2nps6ftr1 X Dsred crosses. Progeny were assayed for fluorescence under under a Leica DM5500 Epifluorescence microscope, and screened for HygromycinB resistance on CMNS HygB 228 (50ug/ml) plates. Complete tetrads that contained HygromycinB sensitive, nonfluorescent progeny (and therefore, the intended fluorescent mutant in the same ascus) were screened by PCR to verify genotype (see Fig. S2 in Appendix). Fluorescent nps2nps6ftr1 progeny were collected by phenotype (inability to grow on MM and CMX) and fluorescence. Inoculation was carried out as above for DAB staining. Fourth leaves were collected one day (24 hours) and five days after inoculation, and trimmed with a scalpel to ~1mmx1mm squares along either side of the major vein. For microscopy, leaves were examined in a Leica TCS SP5 Laser Scanning Confocal Microscope at the Boyce Thompson Institute Plant Cell Imaging Center. Images were collected by exciting samples at 488nm with an Argon laser and collecting emissions at 566nm (Dsred) and 675nm (chloroplasts). Images were processed in ImageJ [141]. 11. Evaluation of fertility C. heterostrophus crosses (Table IV.3) were set up as described previously [45]. Small mycelium scrapings of parental strains were placed in triplicate on either side of a 5mmx5mm autoclaved corn leaf, on Sach’s medium. Crosses between the WT strains C4 (MAT1-2) and C2 or CB7 (MAT1-1) were set up as a control for all experiments. Fertility was evaluated based on the average number of pseudothecia per cm2 senescent corn leaf and the average number of asci per pseudothecium. For hapX crosses, Sach’s medium was complemented with 1mM ascorbic acid, 100uM ferric citrate to alter ROS and iron status (alongside plain Sach’s). Pseudothecia were counted in two separate experiments on three separate cross plates each. At least five pseudothecia per cross were collected, squashed, mounted, and observed under a Nikon Eclipse E600 stereo 229 microscope to count ascospores. All asci in the mount were characterized as empty, containing one, two, three, or more than three ascospores, and represenatitve asci were photographed using Spot (Diagnostic Instruments) imaging software. The entire experiment was carried out twice. 230 C. Results and Discussion 1. Construction and characterization of RIA mutants. RIA mutants were generated by deleting C. heterostrophus FTR1 and FET3 genes. Orthologs of the S. cerevisiae iron permease FTR1 and multicopper oxidase FET3 in the RIA pathway (Table IV.4) were identified initially by blast search [142] with the yeast genes against the C. heterostrophus C5 genome [143]. FET3 and FTR1 were adjacent in the genome, allowing them to be deleted separately, as well as with a single construct. The yeast low affinity ferrous iron transporter FET4 was also used as a blast query, and again a single ortholog was identified and deleted. All candidate deletion strains were single conidiated to eliminate heterokaryons and confirmed by PCR using primer pairs internal to the deleted gene, and a set of primer pairs in which one primer is internal to the selectable marker, and the other is external to the the 5’ or 3’ flank used to dete the gene (see Materials and Methods, and Fig. S2 in Appendix). Table IV.4. Genes studied in this chapter. C. heterostrophus S. cerevisiae A. fumigatus function FTR1 FTR1 FTRA High affinity iron permease (RIA component) FET3 FET3 FETC multicopper oxidase/ferroxidase (RIA component) FRE1 (Not FRE1 FREB Ferric reductase (RIA component) deleted) SRE1 NA SRE1 Transcription factor, repressor under iron replete conditions HAPX NA HAPX Transcription factor, activator under iron depleted conditions FET4 FET4 FET4 Low affinity ferrous iron transporter NPS2 NA SIDC NRPS biosynthesizing intracellular NPS6 siderophore NA aSIDD NRPS biosynthesizing extracellular siderophore ABC6 NA NA ABC transporter adjacent to NPS6. aSIDD is the NRPS responsible for production of extracellular siderophores in A. fumigatus, but the product, fusarinine, is a different siderophore than C. heterostrophus’s NPS6 (coprogen) 231 None of these mutants (fet3, ftr1, fet3ftr1, fet4) was different from WT with respect to growth, sexual and asexual development, oxidative and iron stress, or virulence. Because the extracellular siderophore mutant nps6 is different from WT in all of these phenotypes [44], we can conclude that C. heterostrophus prefers high affinity siderophore iron acquisition over RIA. C. heterostrophus, like all other fungal species where both extracellular siderophore production and RIA have been disrupted, favors one method or the other. 2. Construction of high affinity iron acquisition polymutants. While siderophores may be the preferred method of iron acquisition for C. heterostrophus, it is possible that RIA plays a secondary or backup role. For example, nps2 intracellular siderophore mutants have no phenotype except sterility when crossed to another nps2 strain [45], however all nps6 and nps2- associated phenotypes are exacerbated in nps2nps6 double mutant strains. 232 Figure IV.2. Plating tetrads on different media allows rapid identification of desired polymutants. Eight ascospores from a single ascus in a cross between parents nps6ftr1 (1722-R-1) X nps2 (Chnps2-1) were patched on to CMX, MM, CMX+iron, and CMNS containing hygromycin B media. Position of each progeny is the same across plates, WT progeny are indicated with red arrows, and triple nps2nps6ftr1 mutant progeny with blue arrows. All eight progeny grow on CMX+iron. nps2nps6ftr1 progeny do not grow on CMX and MM, indicating that they require iron. WT progeny do not grow on selective medium containing hygromycin B. Pigmentation was used as an internal control for 1:1 segregation. Pigmentation is controlled by a single gene, ALB1, which is unlinked to the other genes in this study. Therefore, we next generated combinatorial mutants lacking ability to produce intracellular siderophores (nps2), extracellular siderophores (nps6), and/or RIA (ftr1, fet3, 233 ftr1fet3) components. To accomplish this, sexual crosses were performed (Table IV.3) and either random progeny or complete tetrads (seven or eight progeny from a single ascus) were collected. Tetrad collection allows for simplified selection of desired combinatorial mutants, because products of meiotic division remain in the same ascus. An ascus from a cross between two mutant parents that contains WT progeny must also contain progeny that have both parental mutant genotypes (Fig. IV.2). nps6ftr1, nps2ftr1, nps6fet3, nps6fet3ftr1, and nps2nps6ftr1 combinatorial mutants were collected in this way and confirmed by PCR using the same primer schematic to determine correct knockout construct integration (see Fig. S2). nps2nps6 double mutants were generated previously [103]. 3. Growth and asexual development of combinatorial iron mutants i. RIA and siderophore double mutant growth While RIA mutants display WT morphology, mutants lacking both RIA and extracellular siderophores (nps6ftr1, nps6fet3, nps6fet3ftr1) grow at a reduced rate on CMX, and with less pigmentation, compared to WT (Fig. IV.3). Note that mutants lacking extracellular siderophores (nps6), intracellular siderophores (nps2), or both (nps2nps6) still grow at WT rates on CMX [103]. Growth could be restored to WT levels by providing 100uM iron supplements (Fig. IV.3). This demonstrates that while RIA may not be required for growth in the presence of NPS6, without it, the fungus is so taxed for iron it cannot grow properly. Note that nps6fet3, nps6fet3ftr1, and nps6ftr1 are phenotypically the same. This suggests that deleting FTR1, FET3, or FTR1 and FET3 is sufficient to disrupt RIA. This is not surprising, as studies in yeast demonstrate that Fet3 and Ftr1 corequire each other for RIA to operate, as the two proteins products escort one another to the membrane [64]. Further experiments described in this thesis 234 focus on ftr1 mutants for RIA as they are phenotypicaly the same, with respect to iron, as fer1 and fer1ftr1. On MM, growth of nps6ftr1 was almost totally abolished, and addition of supplemental iron could not fully restore growth to WT levels. (Fig. IV.4) All other strains, except nps2nps6ftr1 and nps2nps6ftr1fet4 mutants (see below) are able to grow on MM without supplemental iron. nps2nps6 mutants were reported to display abnormal pigmentation on MM [103]. That phenotype was replicated here, although there was some variation in pigmentation within and between strains. The nps6 mutant was also reduced in pigmentation, while it was previously reported to be like WT on MM, suggesting there may be variation between these experiments. 235 WT# CMX# fet3% fet3&r1% &r1% CMX# +Fe# nps6&r1% nps6fet3% nps6fet3&r1% nps2nps6&r1% CMX# CMX# +Fe# Figure IV.3. Impaired growth of double and triple mutants on CMX. RIA mutants fet3, fet3ftr1, and ftr1 grow like WT on CMX, and do not benefit from supplemental iron (top row). RIA and extracellular siderophore mutants nps6ftr1, nps6fet3, and nps6fet3ftr1, however, are delinquent in growth on CMX ,with reduced pigmentation and colony diameter (bottom row). Adding supplemental iron (100uM ferric citrate) can restore colony growth rate, although colony morphology remains altered, with less aerial hyphal growth. nps2nps6ftr1 mutants, missing RIA, extracellular siderophores, and intracellular siderophores, are extremely reduced in growth on CMX, extending only several millimeters past the inoculum plug. Supplemental iron restores growth to resemble nps6ftr1. ftr1 and nps2nps6ftr1 photos, separated by the white line, are from a separate experiment under identical conditions. 236 ii. nps2nps6ftr1 nps2nps6ftr1 triple iron mutants, lacking not only RIA and siderophore iron acquisition mechanisms, but also intracellular siderophores and thus the ability to store iron internally as well, are almost completely unable to grow on CMX, and do not sporulate (Fig. IV.3). Even with supplemental iron, colony morphology is significantly altered compared to nps6ftr1. nps2nps6ftr1 is absolutely unable to grow on MM, and while iron supplementation slightly restores growth, it is not to the extent of nps6ftr1 (Fig. IV.4). Examining liquid CM cultures of nps6ftr1 and nps2nps6ftr1 revealed that hyphal morphology of germinated conidia is abnormal without supplemental iron both mutants (Fig IV.5). Without iron, nps6ftr1 growth is thin and abnormal, with hyphae frequently changing direction of growth. nps2nps6ftr1 hyphae do not grow at all in CM, but instead become melanized, suggestive of senescence (Fig. IV.5). Supplemental iron alleviates colony morphology defects for nps6ftr1, but not growth rate; nps2nps6ftr1 grows with supplemental iron, but morphology is still defective. As reported previously, nps2, nps6, and nps2nps6 strains show WT growth and morphology in vitro (Fig. IV.4) [103]. The single RIA mutants (ftr1, fet3,fet3ftr1) generated in this study, as well as the nps2ftr1 double mutant, also grew like WT (Figs. IV.3, IV.4) 237 MM# MM+Fe# MM# MM+Fe# WT# nps6% 'r1% nps2'r1% nps2% nps2nps6% nps6'r1% nps2nps6'r1% nps2nps6'r1fet4% Figure IV.4 Impaired growth of double, triple, and quadruple mutants on minimal medium. WT, all single mutant strains, and nps2ftr1 and nps2nps6 double mutants are able to grow on minimal medium (MM) with and without supplemental iron. nps6ftr1 double mutant growth is highly impaired on MM, and iron supplementation (100uM ferric citrate) partially, but not completely, restores growth. nps2nps6ftr1, and nps2nps6ftr1fet4 mutants are completely unable to grow on MM without supplemental iron, on which they grow like nps6ftr1 mutants. 238 CM# WT#(C4)# nps6%r1( CM+Fe# nps2( nps6%r1( Figure IV.5. Observation of nps6ftr1 and nps2nps6ftr1 conidia in liquid culture shows abnormal colony growth. Liquid cultures of WT, nps6ftr1, and nps2nps6ftr1 mutants were grown overnight in CM and CM supplemented with 100um ferric citrate. WT growth is thick in both CM and CM + Fe. Without iron, nps6ftr1 growth is thin and abnormal, with hyphae frequently changing direction of growth. Supplemental iron partially restores this abnormal morphology, but growth is still not as dense as WT. nps2nps6ftr1 hyphae do not grow at all in CM, but instead become melanized, suggestive of senescence. Supplemental iron allows nps2nps6ftr1 mutants to grow, but morphology is still abnormal. 239 iii. nps2nps6ftr1fet4 It should be noted that CMX (and CM) is iron-replete without the addition of supplemental iron. The yeast extract in CMX alone (50ug/g iron) provides ~ 0.9uM iron, and nanomolar contaminant iron is present in most lab reagents. This casts doubt on the functionality of a low affinity iron uptake system in C. heterostrophus (ie, Fet4). In Aspergillus, for example, high affinity systems are thought to be active only under iron depleted conditions [117]. If RIA/siderophore double mutants are unable to grow normally with the 0.9uM iron found in CMX, it seems unlikely they are able to utilize the abundant iron through low affinity means. If C. heterostrophus utilizes a low affinity ferrous iron transporter, nps2nps6ftr1 mutants should show a preference for ferrous salt. When testing different concentrations of chelated and unchelated ferric and ferrous iron, no such preference was found (Fig. IV.6). Instead, a threshold concentration between 2.5uM and 25uM of ferric or ferrous salt, as well as chelated ferric iron, is able to supplement growth. To further test the role of fet4 in C. heterostrophus iron acquisition, we generated an nps2nps6ftr1fet4 mutant, lacking RIA, internal and external siderophores, and the low affinity ferrous iron transporter Fet4. nps2nps6ftr1fet4 mutants are morphologically identical to nps2nps6ftr1 mutants, and support the suspicion that Fet4 is not involved in iron acquisition in C. heterostrophus (Fig. IV.4). It is possible that C. heterostrophus relies on either RIA or siderophores for iron acquisition under iron replete conditions. Alternatively, C. heterostrophus may utlize another unidentified (yet insufficient) iron acquisition mechanism. 240 Ferrous' sulphate' 0.25uM' 2.5uM' 25uM' Ferric' chloride' Ferric' citrate' Figure IV.6. Ferrous salt, ferric salt, and ferric citrate supplement nps2nps6ftr1 mutant growth. nps2nps6ftr1 mutant strains are unable to grow on CMX (see Fig. IV.3). To remedy this, supplemental iron was added to CMX in the form of ferrous sulphate (Fe2+ salt), ferric chloride (Fe3+ salt) and ferric citrate (Fe3+ chelated) at 0.25, 2.5, and 25 uM. A threshold concentration of 25uM for any form of iron was able to partially restore growth and asexual development. No excess beyond this amount could further complement the nps2nps6ftr1 phenotype. Plates were grown for 5 days in the dark, resulting in the lack of pigmentation seen here. v. Asexual spore production Asexual conidia production was assessed on complete medium with and without supplemental iron for all strains (Fig. IV.7). WT, nps2, nps6, ftr1, nps2ftr1, and nps2nps6 strains all produce 5-10x107 conidia/plate, with or without 150uM supplemental ferric citrate. nps6ftr1, however, produces 100 fold fewer spores than this on CM; supplemention with ferric citrate restores spore production to that seen in nps2nps6. nps2nps6ftr1 strains completely fail to 241 produce conidia on CMX, but spore production can be restored, albeit to 10 fold fewer spores (1x106) than WT per plate. Conidia/plate, 100000000" 10000000" 1000000" 100000" 10000" 1000" 100" 10" 1" CMX" CMX"Fe" Figure IV.7. nps6ftr1 and nps2nps6ftr1 are impaired in asexual spore production. WT (C4) strains, as well as nps2, nps6, ftr1, nps2ftr1, and nps2nps6 single and double mutants produce 1x107 conidia/plate. nps6ftr1 mutants produce 100-fold fewer, and nps2nps6ftr1 mutants do not produce conidia on CMX. With 50uM supplemental iron, nps6ftr1 conidia production is largely restored. nps2nps6ftr1 mutants are also able to produce conidia (but 10-fold less than WT) with supplemental iron. The nps2nps6ftr1fet4 tetramutant, lacking RIA, siderophores, and the low affinity ferrous iron transporter, is phenotypically identical to nps2nps6ftr1. The severity of the growth defect observed in nps6ftr1 strains suggests that RIA plays an essential backup role in iron acquisition for C. heterostrophus. Single deletion of ftr1 does not alter any obvious biological processes, however. This is consistent with the long line of evidence that fungi favor one high affinity iron acquisition system over the other, and for C. heterostrophus, siderophores are the favored tool. Conventional understanding, however, suggests that low affinity iron uptake systems are sufficient for growth and development under iron replete conditions. Our results, however, suggest that in the absence of siderophores, C. heterostrophus relies on RIA, not low affinity transporters, for iron acquisition. Removal of both 242 RIA and siderophores impairs growth and asexual development, and only by supplying high quantities of iron (much higher than would be found naturally) can growth be restored. Unlike NPS6, FTR1 expression was not repressed by the iron-responsive transcription factor Sre1 [43], supporting the idea that RIA is used under “standard” iron conditions, and not reserved for iron starvation. Additional removal of NPS2 results in extremely limited growth without supplementation, likely due to an already iron limited fungus being extra-sensitive to disruption to iron and oxidative stress homeostasis. 4. Can iron acquisition mutants be fed iron by other strains? nps2nps6ftr1 is incapable of growing on CMX when plated alone. When plated alongside other strains, for example, when whole tetrads are plated together (see Fig. IV.2), some growth was observed, often in abnormal patterns. We developed an assay to test whether this was because the nps2nps6ftr1 strain was utilizing factors produced by other strains. nps2nps6ftr1 mycelia and conidia was laid out in a grid pattern on CM-hygromycin B (CMHygB) plates (Fig. IV.8). WT mycelia and conidia, which cannot grow on medium containing hygromycin B, were placed in one corner of the plate. After 2 days, nps2nps6ftr1 mutants that were adjacent to the WTstrain were able to grow, but not those farther away. Furthermore, colony growth habit was not radial: mutants grew towards the WT strain, but not away from it. nps2nps6ftr1 is therefore able to utilize, and sense, some factor produced by WT C. heterostrophus. 243 nps6% WT# WT# nps2nps6% nps2% Figure IV.8. nps2nps6ftr1 growth is restored by proximity to WT and nps2, but not nps6 or nps2nps6 cultures. nps2nps6ftr1 is unable to grow in isolation on CMNS hygB plates, but grows (poorly) when plated alongside other strains (see Fig. IV.2). A small amount of conidial scrapings (without agar medium) was transplanted in a grid pattern onto CMNS HygB plates. (Left) WT (C4) C. heterostrophus was patched onto the upper right corner of the grid. While WT cultures are sensitive to Hygromycin B and do not grow, nps2nps6ftr1 proximal to the WT fungus is able to grow, and after 3 days has grown substantially towards the WT patch. nps2nps6ftr1 transplants distant to WT are unable to grow. (Right) When nps2np6ftr1 is grown adjacent to culture filtrate from liquid cultures of WT, nps6, nps2, and nps2nps6, only WT and nps2 culture filtrate can restore growth. Given that nps2nps6ftr1 is unable to produce siderophores, it is likely that one of these molecules is the source of the restored growth. Feeding plates were therefore set up with culture filtrates of WT, nps6, nps2, and nps2nps6 strains in four wells situated at the corners of the plate (Fig IV.8). As before, nps2nps6ftr1 located adjacent to WT extract were well restored in growth, as were those next to nps2, although slightly less so. Patches adjacent to nps6 and nps2nps6 filtrates were unable to grow. We hypothesize that coprogen siderophores produced by Nps6 by WT and nps2 mutants mobilize in the medium and feed nps2nps6ftr1. nps2 mutants may 244 produce slightly less Nps6, or export less coprogen, as they are slightly reduced in feeding nps2nps6ftr1. Both nps6 and nps2nps6 lack coprogens, and therefore cannot feed nps2nps6ftr1. Coprogens produced by Nps6 must be secreted into the environment, chelate iron, and find their way back to the growing fungus, where the iron is acquired, and the siderophore recycled. The ATP-driven efflux ABC transporters have been characterized as siderophore exporters, whereas siderophore uptake is facilitated by MFS/SIT1 [110]. The C. heterostrophus ABC transporter ABC6 maps directly adjacent to NPS6, and is therefore a likely candidate for siderophore transport. Previously, abc6 and abc6nps6 double mutants were generated and subjected to HPLC analysis for siderophores in broth and mycelium (Zhang, Gibson and Turgeon, unpublished). abc6 mutants had significantly less coprogen in broth samples than WT, while nps6 and abc6nps6 strains lacked coprogens entirely. When culture extracts are used in feeding experiments, however, the results are quite different (Fig. IV.9). abc6 mutants clearly outperform WT strains in feeding nps2nps6ftr1, but only in the presence NPS6, as abc6nps6 mutants are completely unable to feed nps2nps6ftr1. It is not clear what underlies this apparent contradiction between the HPLC and feeding plate data. One hypothesis is that the previous HPLC data (although these are the same strains) are erroneous, and abc6 mutants actually have more, not less, coprogen in their broth. It is possible that Abc6 is involved in the uptake, not export, of coprogen (although this is not how ABC transporters typically function). Repeating the HPLC experiments is therefore necessary. Another explanation is that while abc6 exports less siderophore into liquid culture, the resulting broth is somehow more capable of restoring growth to nps2nps6ftr1 than WT broth. Because nps6abc6 is unable to restore nps2nps6ftr1 growth, coprogens are still required for this restoration, and it therefore seems unlikely to be the case. 245 nps6% WT# abc6% abc6nps6% Figure IV.9. abc6 mutants overfeed nps2nps6ftr1 triple mutants in an NPS6-dependent manner. nps2nps6ftr1 feeding plates were set up using culture filtrates as in Fig. IV.7, and grown for five days. The ABC transporter mutant abc6 overfeeds, rather than underfeeds, nps2nps6ftr1 mutants, as adjacent triple mutants grow faster and lusher than WT-fed strains. abc6nps6 double mutants do not complement nps2nps6ftr1 growth. 5. Stress sensitivity Deletion of NPS6 leads to hypersensitivity to oxidative stress and iron depletion in C. heterostrophus, and double deletion of NPS2 and NPS6 results in further sensitivity to these stressors [44,103]. Single deletion of of NPS2 does not affect sensitivity to these stresses [45]. These mutants, and the additional combinatorial mutants generated in this study, were assayed for sensitivity to two categories of stress. The oxidative stressors used were H2O2, which can be converted via the Fenton reaction to hydroxy radicals, and KO2, which challenges the fungus with superoxide. For low-iron stress the membrane-permeable iron chelator 2,2’-dipyridyl (2DP) and the membrane-impermeable iron chelator, bathophenanthroline disulfonate (BPS) were used (Figs. IV.10, IV.11). Not surprisingly, ftr1 and nps2ftr1 grow like WT at the concentrations of each stressor tested. nps6 growth is further inhibited than WT for each stress tested. nps6ftr1 growth is delayed on MM without stress, and compared to this baseline, growth 246 is comparable to nps6 for each stress. nps2nps6 growth is more severely disturbed than nps6, when stress concentrations allowed a distinction. nps2nps6ftr1 was not included, as it is not capable of growing on MM. Figure IV.10. Iron mutants are sensitive to oxidative and iron stress. All iron polymutants in this study were plated onto minimal medium (MM) containing increasing concentrations of the oxidative stressors H2O2 and KO2, the membrane permeable iron chelator 2DP, and membrane impermeable iron chelator BPS. Average colony diameter was measured from leading edge to leading edge of hyphal growth, and is displayed in Fig. IV.11. nps2nps6ftr1 is unable to grow on MM without stressors, and so is not included here. 247 Colony&Diameter&(mm)& 70" 60" 50" 40" 30" 20" 10" 0" 70" 60" 50" 40" 30" 20" 10" 0" H2O2& 2DP& MM& H2O2&4mM& H2O2&8mM& Colony&diameter&(mm)& 70" 60" 50" 40" 30" 20" 10" 0" MM& 2DP&75&uM& 2DP&150uM& Colony&diameter&(mm)& 70" 60" 50" 40" 30" 20" 10" 0" KO2& MM& KO2&6&mM& KO2&12&mM& BPS& MM& BPS&100uM& BPS&300uM& Colony&diameter&(mm)& Figure IV.11. Iron mutants are progressively sensitive to oxidative and iron stress. Average colony diameter of stress plates shown in Fig. IV.10 was determined for each strain on each stress. ftr1 and nps2ftr1 mutants are like WT for all stressors. As reported previously, nps6 is more sensitive than WT, and nps2nps6 is, in turn, more sensitive than nps6 on all stressors (not shown at the presented iron stress concentrations) [103]. nps6ftr1 mutants growth is comparable to nps6, although its reduced growth on MM is a complicating factor. These findings confirm that, although deletion of NPS2 alone has no effect on sensitivity to ROS or to low iron, deletion of NPS2 has a clear effect when deleted in combination with NPS6 [103]. nps6ftr1 mutants, on the other hand, were not further reduced than nps6 on these stress plates. ftr1, nps2, and nps2ftr1 mutants, however, are not sensitive to any of the stressors tested here compared to WT. This confirms that Nps6 is sufficient for dealing with both oxidative and iron stress. That nps2nps6 is further sensitive than nps6, while nps6ftr1 is not, suggests that Nps2, but not RIA, can play a secondary role in dealing with these stressors. Yet, 248 np6ftr1 is reduced in growth, while nps2nps6 is not, consistent with RIA, and not intracellular siderophores, being required for high affinity iron acquisition. nps2nps6ftr1 mutants do not grow on MM without supplemental iron, and could therefore not be tested for sensitivity to these stressors. 6. Virulence of iron mutants is reduced It was determined previously that deletion of C. heterostrophus NPS6 leads to reduction in virulence to maize, while deletion of NPS2 does not affect virulence [44,102]. As with oxidative stress and iron chelation stress, double deletion of NPS2 and NPS6 leads to attenuation of virulence to maize beyond that observed for nps6 strains [103]. The full set of iron acquisition and storage mutants generated here, as well as those described previously, were inoculated onto maize plants to evaluate virulence (Figs. IV.12, IV.13). ftr1 and nps2ftr1 strains were not reduced in virulence compared to WT. As in previous experiments, nps6 was reduced compared to WT, and nps2nps6 was further reduced beyond symptoms caused by nps6. nps6ftr1 mutants were similar to nps2nps6. nps2nps6ftr1 mutants were extremely reduced in virulence. Rather than forming lesions, they formed white flecks only. It is possible that these white flecks are not lesions at all, but the accumulation of callose indicative of an incompatible reaction where C. heterostrophus is arrested at the surface. Both nps6ftr1 and nps2nps6 are further reduced in virulence compared to nps6, but only nps2nps6 is more sensitive than nps6 to iron and oxidative stress. The reduced growth of nps6ftr1 in vitro could explain this; alternatively, a factor other than iron or oxidative stress may be involved in planta. 249 WT#(C4)# nps2%r1( nps6( nps6%r1( nps2nps6( nps2nps6%r1( Figure IV.12. Virulence of iron mutants is progressively reduced compared to WT. Representative fourth true leaves from N-cytoplasm W64-A corn inoculated with C. heterostrophus WT and iron mutants are shown. Leaves were photographed after five days. Lesions are dark, long, and necrotic on WT and nps2ftr1 inoculated leaves. nps6 mutants cause smaller brown lesions, while nps6ftr1 and nps2nps6 cause much smaller, lighter lesions with small dark centers. nps2nps6ftr1 mutants do not cause lesions at all, but rather infrequent, small, white flecks. 250 Average'lesion'length'(mm)' 4.5" 4" 3.5" 3" 2.5" 2" 1.5" 1" 0.5" 0" WT#(C4)# nps2,r1# nps6# nps2nps6# nps6,r1# nps2nps6,r1# Plain" Iron" Figure IV.13. Iron mutants are progressively reduced in virulence with or without supplemental iron. Average lesion size of fourth true leaves from N-cytoplasm W64-A maize inoculated with WT and iron mutants was determined. nps2ftr1 (and ftr1, not shown) mutant lesion sizes are like WT. As shown previously, nps6 mutants are reduced in virulence, and nps2nps6 mutants are further reduced than nps6. nps6ftr1 is comparable to nps2nps6, while nps2nps6ftr1 mutants are extremely reduced in virulence compared to WT. Iron was added to the inoculum, and painted onto the leaves of iron treated plants prior to inoculation. This supplementation slightly increased lesion size for not only the iron mutant inoculated plants, but also WT inoculated plants. i. Iron complementation Attempts to restore virulence of any of these strains by adding exogenous iron (Fig. IV.13) and/or the antioxidant ascorbic acid (data not shown) were unsuccessful. Introducing any solution into maize leaves, which are extremely hydrophobic, is difficult. More invasive measures, such as vacuum infiltration or removing leaves and soaking them in solution, were too destructive to the leaf, when combined with the pathogen. Selective restoration of combinatorial mutant phenotypes with iron or ROS scavengers could be very informative for separating the iron chelation sensitivity and oxidative stress sensitivity in planta, but the experimental 251 methodology for supplementation has not been adequately developed for the Cochliobolus/maize pathosystem. ii. Diaminobenzidine A key goal of this study was to differentiate between the role iron and oxidative stress play in iron acquisition mutant phenotypes. One approach is subcellular staining of reactive oxygen species (ROS). Diaminobenzidine (DAB), which stains ROS indirectly by has been used in a variety of plant-fungal systems [144-146] with varying success. Because host cell death is mediated by ROS, and C. heterostrophus is a necrotroph, for DAB to be of use in indicating subcellular localization of ROS or iron, a time point must be identified where ROS are formed, but host tissue has not been extensively killed. C. heterostrophus forms appressoria and penetrates the leaf within 4 hours after inoculation. By 8 hours, DAB staining in WT strains is ubquitious, completely obfuscating the site of infection. This indicates that within 8 hours the cells surrounding the site of penetration have been killed (data not shown). With the caveat that DAB cannot differentiate host cell death from other types ofROS produced, infected leaves were stained with DAB at 24 hours and 5 days post infection (Fig. IV.14). At 24 hours, strong DAB accumulation could be seen in WT, nps6, and nps6ftr1 infected leaves, presumably immediately surrounding points of infiltration. nps2nps6 and nps2nps6ftr1 challenged leaves look more like uninoculated controls, without DAB accumulation. Note that at 24 hours, lesions are already forming, and can be seen by the naked eye if held up to the light. 252 WT# nps6% nps6'r1% nps2nps6% nps2nps6'r1% no#inocula+on# Figure IV.14. DAB staining is reduced in maize leaves inoculated with iron mutants. Inoculated leaves were stained with the reactive oxygen species and cell death dye Diaminobenzidine (DAB) one day (left in each pair) and five days (right in each pair) after inoculation. In WT and nps6 challenged leaves, DAB precipitates form at discrete locations after 24 hours, and ubiquitously throughout the leaf by five days. nps6ftr1 and nps2nps6 form fewer localized DAB precipitates at 24 hours, and much smaller, contained DAB stained areas at five days. nps2nps6ftr1 does not form visible DAB precipitate at any stage of infection. 253 At 5 days, when lesions are plainly visible on the leaf, DAB staining is rampant in WT and nps6 challenged leaves. nps6fr1 and nps2nps6 have discrete lesion shaped DAB stains, often with stain accumulating at the edge, without staining in the middle. nps2nps6ftr1 and uninoculated leaves do not show DAB accumulation. iii. Confocal imaging To further investigate how virulence is impaired in these mutants, Dsred (red fluorescent protein) expressing strains were constructed for WT and nps6, nps2nps6, nps6ftr1, and nps2nps6ftr1 mutants by crossing the respective mutants to the Dsred expressing strain 1761-R7 (Table IV.3). These strains were inoculated onto maize and observed in planta at 24 hours and 5 days after inoculation (Fig. IV.15). Confocal observation at 24 hours found no clear difference in spore germination or appressorium formation among the WT, nps6, nps6ftr1, and nps2nps6 strains. in planta colonization by both nps6ftr1 and nps2nps6 mutants, however, was delayed compared to WT or nps6 at 24 hours (Fig. IV.15). For all strains, sites of fungal entry were often accompanied by autoflourescence originating from the challenged epidermal cell (Fig. IV.16). After 5 days, fungal growth had halted after spreading 100-500µm, and was surrounded by autofluorescence, whereas for WT and nps6, regions colonized by fungi were overrun at this point. When combined with the lesion measurement and DAB data, this suggests that both nps2nps6 and nps6ftr1 are able to enter the leaf and begin the infection process. However, they fail to colonize the leaf extensively, and are arrested, resulting in patches of autofluorescence, perhaps corresponding to a host defense response. 254 one#day# WT#(C4)# five#days# nps6%r1# nps2nps6%r1# Figure IV.15. Iron mutants are progressively impaired in colonizing maize leaves. Dsred expressing strains of WT, nps6, nps6ftr1, nps2nps6, and nps2nps6ftr1 mutants were constructed and inoculated onto maize plants. Leaves were visualized by confocal microscopy after one and five days. The scalebar (lower right of each image) is set to 50um. WT (and nps6 mutants, not shown) were able to colonize the length of several epidermal cells after 24 hours, and by five days had extensively colonized the leaf. nps6ftr1 (and nps2nps6, not shown) was reduced in both initial and late stage colonization, and at five days was restricted to an autoflorescent patch of plant tissue. nps2nps6ftr1 was able to germinate and form appressoria, but did not typically penetrate or grow inside the leaf. When it did, it did not progress beyond the initial cell colonized. 255 nps2nps6ftr1, on the other hand, had fewer conidia to observe/sample at both 24 hours and 5 days, although similar numbers to e.g., WT, were applied initially, suggesting that some conidia fail to attach to the leaf. Those that could be observed were able to germinate and form appressoria. No in planta fungal growth could be observed at 24 hours. At 5 days, fungal growth was not observed beyond the initial site of penetration in any case. nps2nps6& nps6'r1& Figure IV.16. Host autoflorescence accompanies fungal entry. Dsred expressing strains of nps2nps6 and nps6ftr1 have breached the leaf 24 hours after inoculation. Conidia are still attached (open white arrows), and the precise site of penetration can be identified ( closed yellow arrows). Autofluorescence of the penetrated epidermal cell and surrounding cells occurs in an emission spectrum indistinguishable from Dsred. Additionally, strong autofluorescence surrounded sites where nps2nps6ftr1 had entered the leaf. Combined with the white flecks observed on whole leaves, and the lack of DAB staining, this suggests that nps2nps6ftr1 does not advance as a pathogen beyond a non-host interaction. The strong, localized autofluorescence, combined with white flecking, suggests callose deposition, a hallmark of penetration resistance. 256 7. The iron-responsive transcriptional activator HapX The diversity and severity of phenotypes observed in iron acquisition and storage mutants demonstrates the pivotal role iron plays in fungal biology. Regulation of genes that contend with iron acquisition, storage, metabolism, and ROS response is therefore of top priority for the cell. The regulatory players in C. heterostrophus are like those in the Aspergilli, as its genome contains both a Sre1 repressor, primarily active under iron replete conditions, and a HapX activator, primarily active under iron depleted conditions. previously, the SRE1 gene was deleted in C. heterostrophus [43] and expression assays confirmed that NPS6 but not FTR1 arerepressed under Fe replete conditions in an sre1 dependent manner. sre1 mutant growth is greatly slowed in vitro, and virulence is correspondingly reduced. Here, hapX mutants were constructed to further understand iron regulation. Targeted gene knockout constructs were generated by PCR, and confirmed using internal and external primer pairs (Table IV.2), as described in Section IV.B.4 (see Fig. S2 for deletion and confirmation schematic). i. Growth Unlike sre1 mutants, hapX mutants are like WT in terms of growth in vitro (data not shown). It is possible that sre1 mutants have an abnormal morphology specifically because of the metabolic costs of a lack of genetic repression, including costly secondary metabolites, not simply because of perturbed iron-related expression. hapX mutants, on the other hand, are missing activity of an activator, and not a repressor. They would therefore not experience this tax on their metabolism. ii. Stress sensitivity Stress assays with iron chelators, showed that hapX mutants are sensitive to low iron stress. The minimum inhibitory concentration of the membrane permeable iron chelator 2DP 257 was lower for hapX mutants than for WT; hapX (and nps6) did not grow at 200mM 2DP, unlike WT which did (Fig. IV.17). Likewise, hapX mutants are also more sensitive to the membrane impermable iron chelator BPS than WT. hapX growth was perturbed at 100uM BPS, while WT growth was the same at 100, 200, and 300 uM BPS (Fig. IV.17). nps6 mutants grew slightly less than hapX mutants on iron stress plates. hapX mutants were not sensitive to the oxidative stressor hydrogen peroxide, and strains grew like WT at up to 8mM H2O2, known to inhibit growth of control nps6 mutants. Average'colony'diameter'(mm)' Average'colony'diameter'(mm)' Average'colony'diameter'(mm)' 60" 50" 40" 30" 20" 10" 0" WT" 2DP' nps6" hapX1" hapX3" MM" 75"uM"2DP" 150"uM"2DP" 225uM""2DP" 60" 50" 40" 30" 20" 10" 0" WT" BPS' nps6" hapX1" hapX3" MM" 100uM"BPS" 200uM"BPS" 300uM"BPS" 60" 50" 40" 30" 20" 10" 0" WT" H2O2' nps6" hapX1" hapX3" MM" 2mM"H2O2" 4mM"H2O2" 8mM"H2O2" Figure IV.17. hapX mutants are sensitive to iron, but not oxidative, stress. WT, and hapX (two independent transformants, hapX-1 and hapX-3), and nps6 mutants were grown on MM containing increasing concentrations of iron chelative (2DP and BPS) and oxidative (H2O2) stress for five days in the dark. nps6 was unable to grow on 8mM H2O2, whereas hapX mutants grew like WT at this concentration. On iron stress plates, however, hapX mutant growth was reduced compared to WT. hapX mutants were unable to grow at 150uM 2DP, and were severely reduced in growth at all concentrations of BPS. nps6 was slightly more sensitive than hapX to both 2DP and BPS. 258 HapX is therefore required for responding to iron chelation stress, but not oxidative stress. It is likely that in 2DP and BPS media, the fungus responds to a reduced supply of available iron by activating NPS6 in a HapX dependent manner. Loss of HapX does not affect responses to oxidative stress, although nps6, nps2nps6, and nps6ftr1 mutants are sensitive to such stressors. This suggests that NPS6 and NPS2 are required for dealing with oxidative stress, but not in a HapX dependent manner. That nps6ftr1 mutants, like nps2nps6 mutants, are more sensitive to oxidative stress than nps6 mutants suggests that iron acquisition itself is an important component of dealing with oxidative stress. hapX mutants may therefore be compensated in some ways by activity of other regulators, such as Chap1 [42], which activates genes in response to oxidative stress. The nps2nps6ftr1 feeding assay (Figs. IV.8, IV.9) is one way to examine how ironrelated gene expression is altered in hapX mutants. If hapX mutants cannot activate expression of NPS6, for example, the amount of coprogen siderophore produced should be reduced, and hapX should be less effective at feeding nps2nps6ftr1. This is not the case (Fig IV.18), as a hapX mutant is able to supplement nps2nps6ftr1 mutants like WT does. This suggests that hapX mutants are not impaired in siderophore production, either because NPS6 is not regulated by HapX, or because NPS6 is sufficiently expressed by other factors in the absence of HapX. 259 hapx% WT#(C4)# nps6% abc6% Figure IV.18. hapX mutants feed nps2nps6ftr1 siderophores as WT. Grids of nps2nps6ftr1 mycelium scraping serve as an indicator for siderophore production. Culture filtrate from WT (C4), hapX, nps6, and abc6 colonies was collected and applied into wells in the corners of the nps2nps6ftr1 grid. Plates were photographed after five days. WT and hapX culture filtrate complemented nps2nps6ftr1 mutants to the same degree, suggesting hapX is supplying similar siderophore levels as WT. nps6 culture filtrate is unable to complement nps2nps6ftr1 mutants, whereas abc6 complements nps2nps6ftr1 mutants better than WT (as in previous experiments, Fig. IV.9). iii. Sexual development HapX upregulation of genes in response to iron depletion is complimentary to Sre1 suppression of genes in response to iron replete conditions. In A. fumigatus and A. nidulans, loss of both HapX and SreA results in synthetic lethality [117]. To determine if this was the case for C. heterostrophus, hapx mutants were crossed to sre1 mutants [43]. sre1 X hapX crosses, and control hapX X WT crosses, all produced pseudothecia but failed to produce asci with a full set of ascospores (Fig. IV.19). Note that sre1 X WT (C9), as well as WT (C4 X C9) control crosses, all produced tetrads with ascospore numbers typical of WT crosses (7-8 spores). In 260 contrast, when hapX mutants are involved in a cross, pseudothecia contained predominantly 90% empty asci (Fig. IV.20), and the rest of the asci contained a one - three ascospores. Figure IV.19. hapX mutants are impaired in ascospore development. WT crosses (C4 X CB7) produce full asci with (typically) eight coiled ascospores per ascus (left). Crosses between hapX mutants and the CB7 WT tester strain (as well as hapX X sre1, data not shown) do not produce healthy asci. The majority of asci are empty, and those asci that do form ascospores produce only a few (1-3) ascospores/ascus. Example ascospores are labeled with blue open arrows, asci with red closed arrows. Scalebar is set to 50µm. 261 Percent'asci'containing'ascospores' 100%# 90%# 80%# 70%# 60%# 50%# 40%# 30%# 20%# 10%# 0%# hapX11# hapX11#ascorbic#acid# WT# 4+#ascospores# <4#ascospores# WT#ascorbic#acid# Figure IV.20. hapX mutant crosses do not produce complete tetrads. WT crosses (C4 X CB7) produce full asci with up to eight ascospores per ascus. 80% of asci contain four or more ascospores, 10% contain less than four, and 10% are empty. Crosses between hapX mutants and the CB7 WT tester strain (as well as hapX X sre1 mutants, data not shown) do not produce healthy asci. The majority of asci are empty (90%), and those asci that do form ascospores produce fewer than four ascospores/ascus. Application of ascorbic acid (or iron, data not shown) to the cross plates could not complement this phenotype, although it did slightly reduce the % healthy asci produced in WT crosses. Interestingly, in CB7 (albino WT) X hapX (pigmented) mutant crosses, the ratio of albino to pigmented pseudothecia was no different from control crosses, and albino pseudothecia (hapX was acting as the male) were still unable to produce regular asci. This phenotype is therefore not a case of female sterility (as is the case for ssk1, [147]), or a recessive trait only seen in self crosses (such as nps2 X nps2 producing empty asci [45]), but a dominant trait. Furthermore, all ascospores recovered from hapX (hygR) X CB7, C2, C9 (hygS) crosses are 100% hygromycin sensitive (Fig. IV.21). This suggests that ascospores are viable when theyhave a functional copy of HAPX. This phenomenon occurs in pseudothecia in which hapX strains act as either the male or the female. 262 CMX$ CMNS$Hygromycin$B$ C9$ hapX%1' C9$ hapX%1' Figure IV.21. All hapX X WT progeny are WT (Hygromycin B sensitive). When viable progeny are collected from hapX X C9 (WT pigmented) crosses, none of those that survive retain the hygR phenotype, suggesting they are contain WT copies of HAPX. Ascospores that do not grow after collection may be the missing hapX progeny. nps2 X nps2 crosses also fail to produce ascospores, although nps2 X WT crosses produce healthy progeny [45]. Because hapX is an iron regulatory gene, it seems reasonable that the sexual phenotype observed with hapX mutants is connected to this similar but distinct phenotype seen in nps2 homozygous crosses. nps2 X nps2 crosses could be partially rehabilitated to produce ascospores by supplementing cross plates with 250uM excess iron, raising the question of whether this defect can be complemented by adding supplements [45]. To address this, cross plates were set up with Sach’s (standard crossing medium), with 100uM ferric 263 citrate, and Sach’s with 1mM ascorbic acid (Fig. IV.22). Ascorbic acid is an antioxidant, and given the role ROS plays in developmental cues for fungi it seemed possible that this could alter the hapX mutant, or even the WT, phenotype. C4'X'CB7' Sach’s' Sach’s+'' ascorbic'acid' Sach’s+'' ferric'citrate' hapX%1'X'CB7' Figure IV.22. hapX mutants are male and female fertile in sexual crosses. Crosses between pigmented WT (C4) and hapX strains (hapX-1 and hapX-3, not shown) to the albino WT tester strain CB7 produced black and white pseudothecia for both WT and hapX mutants, demonstrating that hapX is like WT in its ability to perform as a male and female partner. Addition of 1mM ascorbic acid or 100uM ferric citrate did not disrupt or alter the production of pseudothecia for either WT or mutant crosses. Ascorbic acid plates formed a halo of precipitate around the autoclaved corn leaf substrate. Iron (data not shown) and ascorbic acid did not alter the number of pseudothecia or asci produced in WT crosses (Fig. IV.20). Crosses involving hapX mutants remained severely impaired, with no change in the number of ascospores per ascus. Interestingly, the Sach’s with 264 ascorbic acid plates were cloudier than regular Sach’s, and a halo of clearing could be observed around the crosses (Fig. IV.22). This could be indicative of a large amount of ROS produced. iv. Virulence hapX mutants were significantly reduced in virulence (Fig. IV.23). The average lesion length was 2mm for hapX, and 5mm for WT (Fig IV.23). This level of reduction is equivalent to that seen in nps6, but not the further reduced nps2nps6 and nps6ftr1 mutants. hapX mutants still produce T-toxin, and therefore still produce T-toxin induced lesions on T-cytoplasm corn (Fig. IV.23) equivalent to WT. This reduction in virulence is particularly illuminating when combined with the observation that hapX is sensitive to iron-chelation, but not oxidative, stress. A central mystery to the role of Nps6 in virulence was differentiating between iron starvation and oxidative stress sensitivity as a result of iron starvation. hapX mutants are able to cope with oxidative stress in vitro: we can therefore propose that oxidative stress is not the primary stress C. heterostrophus faces in the infection process. If it were, hapX mutants would be able to deal with it, and would not be reduced in virulence. Instead, it is reduced to levels similar to those seen in nps6. This suggests that the primary challenge C. heterostrophus encounters in planta is iron acquisition, and that nps6 mutants are reduced in virulence first due to their inability to acquire iron in planta, not their sensitivity to host generated ROS. This reasoning assumes that the reduction in virulence seen in hapX mutants is primarily due to the mutant nps6 gene phenotype. This is likely not true, as we expect HapX to regulate a larger set of iron-related genes. Even so, it does inform our understanding of the iron/ROS debate with regard to infection. 265 A WT#(C4)# hapX%3' B 7" 6" 5" Average'lesion'length'(mm)' 4" N"corn" 3" T"corn" 2" 1" 0" WT# hapX(1# hapX(3# Figure IV.23. hapX is reduced in virulence on maize. (A) hapX mutants were reduced in virulence when inoculated onto N cytoplasm corn, forming smaller lesions than WT. (B) Average lesion length was determined for WT and hapX mutants on T and N cytoplasm corn. hapX mutants formed smaller lesions (1.5-2mm compared to 5mm for WT) on N cytoplasm corn. On T cytoplasm corn, hapX mutants were not distinguishable from WT, indicating hapX mutants are not impaired in their ability to produce and deliver T-toxin. 266 D. Conclusions C. heterostrophus follows the general pattern for high affinity iron acquisition seen by other organisms. In requiring siderophore-mediated iron acquisition for virulence and oxidative/iron stress management, it is unaffected by loss of RIA alone. RIA and siderophore double mutants, however, are extremely perturbed, and defective in basic growth and development. Only one other study has generated RIA/siderophore double mutants, in A. fumigatus and these mutants, too, suffered grave morphological defects. The A. fumigatus sidA/ftrA double mutants lack both extracellular and intracellular siderophore production, as SIDA is required for biosynthesis of both extracellular and intracellular siderophores [95]. Our study with C. heterostrophus is therefore unique in distinguishing between extracellular and intracellular siderophore mutants, in combination with RIA mutants. Interestingly, loss of intracellular siderophores and RIA (nps2ftr1) does not affect stress sensitivity, growth or virulence. As long as NPS6 is present, the fungus remains like WT. The phenotypes observed for hapX mutants reveal several things about C. heterostrophus iron biology. hapX is required for dealing with iron, but not oxidative, stress. hapX mutants are also reduced in virulence. nps6 mutants are sensitive to oxidative and iron stress, and also are reduced in virulence. This distinction could suggest that nps6 mutants are reduced in virulence because of their sensitivity to low iron stress and that sensitivity to oxidative stress is coincidental. Alternatively, it could suggest that C. heterostrophus responds to the iron and oxidative challenges of the host plant in a hapX dependent manner, in such a way that genes responsive to oxidative stress (for example, SRE1) cannot compensate. 267 The defect seen in ascospore development of hapX mutants is intriguing. hapX mutants are both male and female fertile, as demonstrated by pigmented and albino pseudothecia produced in crosses to the albino strain CB7. In C. heterostrophus, mutants impaired in sexual development are often less able to act as the female partner (as seen in ssk1 and skn7 histidine kinase mutants, [147]). Alternatively, the defect in nps2 sexual development is only seen when nps2 is crossed to nps2; nps2 X WT crosses develop normally [45]. 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Each linkage group (1-15, and the dispensable chromosome B1) is represented by three lines: the left solid line for the race T parent, the dotted right line for the race O parent, and linkage between markers (middle). Breaks in this middle line indicate where genetic linkage could not be determined, but Southern hybridization to chromosome blots confirmed their presence on the chromosome. Two race O chromosomes 6 and 12, corresponding to two race T chromosomes 6;12 and 12;6, form a cruciform linkage group. The Tox1 locus maps to the intersection of the four-armed linkage group. 288 First round PCR UpF UpR DnF DnR 5' Flank (Up) YFG Second round PCR and integration UpF 3' Flank (Dn) NLC37 5' Flank (Up) HY NLC38 YG NLC38 M13FHyg M13RHyg NLC37 HygB DnR 3' Flank (Dn) 5' Flank (Up) YFG 3' Flank (Dn) Verification of transformation Upext InF NLC37 InR NLC38 Dnext 5' Flank (Up) HygB 3' Flank (Dn) Figure S2. Generalized schematic for gene deletion and confirmation of selectable marker integration. Split marker PCR [2] is a multi-step PCR process for targeted deletion of a gene and integration of a selectable marker. First, 700-800bp DNA immediately upstream (5’ Flank up) and downstream (3’ Flank Dn) of Your Favorite Gene (YFG) are amplified, using primers UpF and UpR) and DnF and DnR, respectively. The upstream reverse (UpR) and downstream forward (DnF) primers (drawn as black and red overhangs) include adapters matching the HygB gene. HygB, carried on the pUCATPH plasmid is also amplified in two overlapping segments, using primer pair M13RHyg/NLC37 for the 5’ end (HY) and M13FHyg/NLC38 for the 3’ end (YG). Next, the purified PCR products are combined in two reactions: the 5’ upstream and HY fragments are joined with primer pair UpF/NLC37, and the 3’ downstream and YG fragments are joined with the primer pairs DnR/NLC38. Flanking and HygB fragments are joined due to the adapters on UpR and DnF. These two PCR products are used together as a transformation construct. Successful targeted replacement of YFG with HygB is confirmed using three PCR reactions. First, a band will not be amplified from gene deleted strains using a YFG internal primer pair (InF, InR), but will from WT and ectopic integrants. Second and third, a band will 289 appear only in successfully gene deleted transformants using a primer external to the flanking region construct (5’ Upext and 3’Dnext) combined with a primer internal to HYG. Ectopic transformants will not produce correctly sized bands for these PCR reactions, nor will WT. Figure S3. Genetic distance correlates with physical distance on the C. heterostrophus map. RFLP markers located on the same scaffolSduwpepreleumseedntotaprlyotFgigenuerteic2distance against physical distance. Genetic distances between RFLPs determined by Tzeng et al. [1]. Available online at doi:10.1371/journal.pgen.1003233.s001. Figure S4. A C. heterostrophus dispensable chromosome is present in some but not all C. heterostrophus strains. Mauve alignment of the genome of strains Hm540 and C5. Colored blocks [Locally Collinear Blocks (LCB)] indicate matches between genomes. Note there are only a few matches (colored blocks) in Hm540 to scaffold S16/chromosome B1 in C5. The C5 S16 scaffold (~750 kb) is shown in its entirety below the whole genome alignment. Available online at doi:10.1371/journal.pgen.1003233.s002. 290 Figure S5. C. sativus SSR sequences anchor sequenced scaffolds to the genetic map. Genetic map of C. sativus based on 68 SSR markers, 102 amplified fragment length polymorphism (AFLP) markers, 34 RFLP markers, two polymerase chain reaction–amplified markers, the mating type locus (CsMAT), and the barley cultivar-specific virulence locus (VHv1). Of the 37 linkage groups, 30 were assigned to 16 of the 157 scaffolds based on alignment of mapped SSR markers to the sequence assembly of ND90Pr. Linkage groups are on the left (open bars, numbered at the top) and assembled scaffolds (solid black bars, numbered at the top) are on the right. The start point of each scaffold is at the top. Dotted lines connect the SSR loci on the genetic map and physical map. AFLP markers flanking the VHv1 locus and the locus itself are highlighted in red (Fig. I.7). The scale bar corresponds to 500 kb. Available online at doi:10.1371/journal.pgen.1003233.s003. 291 Figure S5 292 Figure S5 (continued) LG5 (147) G98EV-a 13.3 (262) CsSSR_95 14.0 (2) E-AG/M-CG-293 18.3 (18) E-AT/M-CG-318 17.0 (187) CsSSR_95_2 19 32.5 (160) CsSSR_95_3 30.4 4 (216) CsSSR_20 9.8 (218) CsSSR_22 7.8 (217) CsSSR_21 21.0 (219) CsSSR_23 31.0 (193) CsSSR_23_3 LG4 0.0 (126) B149EV-b (58) E-AC/M-CC-188 25.2 (123) C70EV 10.1 (221) CsSSR_27 30.1 3.5 19.0 0.0 0.0 (172) CsSSR_27_2 (146) B107P (93) E-AT/M-AC-638 (94) E-AT/M-AC-637 (175) CsSSR_27_3 LG15 (223) CsSSR_29 35.3 (224) CsSSR_30 20.1 (183) CsSSR_30_3 7.7 (81) CsMAT 20.2 0.0 (52) E-AT/M-CA-317 (41) E-AT/M-CT-353 LG31 2.0 (60) E-AC/M-CG-343 (168) CSSR_30_4 LG17 (198) CsSSR_31_3 29.9 0.0 17.7 16.6 3.1 10.6 1.0 0.0 1.0 1.9 24.0 0.0 0.0 0.0 0.0 0.0 0.0 4.9 (119) E-GT/M-AG-109 (106) E-GC/M-AG-115 (63) E-AT/M-CC-611 (226) CsSSR_32 (227) CsSSR_33 (228) CsSSR_34 (99) E-AT/M-AC-191 (26) E-AA/M-CT-208 (151) B114EV (120) E-GT/M-AG-105 (75) E-AG/M-CT-703 (108) E-GC/M-AG-82 (107) E-GC/M-AG-104 (82) VHv1 (4) E-AG/M-CG-121 (59) E-AC/M-CG-910 (71) E-AG/M-CA-207 (7) E-AA/M-CC-614 5 40 LG32 4.0 (229) CsSSR_35 3.0 (230) CsSSR_36 (231) CsSSR_37 LG33 (139) B88EV-f 9.0 (232) CsSSR_38 31.3 (233) CsSSR_39 6 LG34 0.0 3.1 12.0 (5) E-AG/M-CG-110 (127) B149EV-c (197) CsSSR_39_3 (234) CsSSR_40 22.2 (182) CsSSR_40_2 7 Figure S4, cont’d 293 Figure S5 (continued) 294 Figure S6. Analysis of the mating type region in Cochliobolus spp. and S. turcica. S. turcica 28A, C. sativusND90Pr, C. carbonum 26-R-13, C. heterostrophus Hm540, and C. heterostrophus PR1x412, and the reference C. heterostrophus C5 strains are MAT1-1, while the others are MAT1-2. 10 kb regions flanking the MAT idiomorphs were aligned for each mating type. In all cases, the order of genes immediately surrounding the MAT locus (~20 kb) was conserved. Genes flanking the MAT locus, differ from those flanking MAT in other ascomycetes. JGI ID numbers are shown for strain C5 (MAT1-1) and C4 (MAT1-2). S. turcica had the most variation compared to the Cochliobolus MAT region, although the MAT genes were well conserved. Approximately 500 bp of the 5′ region and ~3 kb of the 3′ region around the MAT gene were more variable than other regions when all genomes were compared. Available online at doi:10.1371/journal.pgen.1003233.s004. 295 Figure S7. Maximum likelihood tree of NRPS AMP-binding (AMP) domains identified using Augustus. RAxML using the RTREVF model with a gamma distribution was used to infer the maximum likelihood tree and bootstrap support was determined using the fast-bootstrap method with 1000 bootstrap replicates. See Materials and Methods. AMP domains are color-coded by species. Plain branches at the top of the tree are AMP domains from related adenylating enzymes (e.g., acyl CoA ligases, etc). AMP domains corresponding to the C. heterostrophus reference set (Fig. II.3, Table II.4, Table S6,) are indicated on the right of each group. Bootstrap values above the branches. Certain additional AMP domains correspond to well-known metabolites from fungi outside the Cochliobolus and Setosphaeria strains.Gibberella fujikuroi FusS produces fusarin C, Fusarium heterosporum EqiS produces Equisetin,Aspergillus fumigatus GliP produces Gliotoxin, Leptosphaeria maculans SirP produces sirodesmin, Magnaporthe oryzae Ace1 produces an unknown metabolite involved in pathogenicity, Fusarium spp. Esyn1 produces enniatin, AAR, involved in lysine biosynthesis,Tolypocladium inflatum TiSimA produces cyclosporin, Penicillium spp., produces penicillin, δ-(L-α-aminoadipyl)-L-cysteine-D-valine (ACV), C. carbonum HTS1 produces HC-toxin, Hypocrea virens Tex1 produces peptaibols, Alternaria alternata AMT produces AMT toxin, Claviceps purpurea PS1(LPS1) is an NRPS that along with lysergic acid, produces an ergot alkaloid,Metarhizium anisopliae PesA unknown product, Epichloë festucae perA produces Peramine. Genewise trees are available online at doi:10.1371/journal.pgen.1003233.s005. 296 Figure S7 297 Figure S7 (continued) 298 Figure S7 (continued) 299 Figure S7 (continued) 300 Figure S7 (continued) 301 Figure S7 (continued) 302 Figure S8. Maximum likelihood tree of PKS ketosynthase (KS) domains identified using Augustus. RAxML using the WAGF model with a gamma distribution was used to infer the maximum likelihood tree and bootstrap support was determined using the fast-bootstrap method with 1000 bootstrap replicates. See Chapter II Materials and Methods for tree construction methodology. Plain branches at the top of the tree are KS domains from related enzymes. KS domains are color-coded by species as in Fig. S6. Bootstrap values are above the branches. PKS11 is a C. heterostrophus ortholog of the Fusarium verticillioides PKS for fumonisin. Genewise trees are available online at: doi:10.1371/journal.pgen.1003233.s006. 303 Figure S8 304 Figure S8 (continued) 305 Figure S8 (continued) 306 Figure S8 (continued) 307 Figure S8 (continued) 308 Disease level 9 8 7 6 5 4 3 2 1 0 WT ∆115356 Figure S8 Figure S9. Quantification of spot blotch disease induced by the C. sativus wild type and mutant (Δ115356) on barley cv. Bowman. Disease rating was taken at 7 days after inoculation and is based on a 1 to 9 scale. Four replicates were used. Error bar indicates the standard deviation. Available online at doi:10.1371/journal.pgen.1003233.s007. 309 Figure S10. Quantitative real-time PCR analysis of S. turcica PKS gene (protein ID 161586) during infection of maize cultivar W64A-N. Gene expression was normalized based on the expression of the β-actin gene. Values are relative expression levels compared to that in mycelia grown on LCA medium. Samples were collected at 3, 5, 6, 7, and 8 days after inoculation. Error bars indicate the minimum and maximum relative expression values of the gene. Available online at doi:10.1371/journal.pgen.1003233.s008. 310 Figure S11. Complete ChPks1 and ChPks2 phylogenetic tree. A phylogenetic tree was constructed with the KS domain from a large set of PKSs, including every PKS from C. heterostrophus, L. maculans, D. zeae-maydis, and T. stipitatus, 14 additional species, and the top 20 blast hits for ChPks1 and ChPks2. Each sequence is identified by either the Genbank number, JGI protein ID (C. heterostrophus, T. stipitatus, L. maculans), or Augustus ID (D. zeae-maydis), accompanied by a 3-letter strain abbreviation (Table III.4). More detailed explanation in Figs. III.6, III.7. 311 Figure S11 PKS25 PKS22 PKS23 PKS21 312 Figure S11 (continued) PKS18 PKS20 PKS19 313 Figure S11 (continued) PKS11 PKS12 PKS14 PKS15 314 Figure S11 (continued) PKS17 PKS16 315 Figure S11 (continued) PKS4 PKS3 PKS5 PKS9 316 Figure S11 (continued) PKS8 PKS10 PKS6 317 Figure S11 (continued) PKS2 PKS7 PKS1 318 Figure S12. Complete decarboxylase Dec1 phylogenetic tree. A phylogenetic tree was constructed using the C. heterostrophus Dec1 protein sequence, and its top 50 blast matches retrieved from Genbank. See Fig. III.10 for annotation. 319 Figure S12 320 Figure S12 (continued) DEC1 321 Figure S13. Complete Lam1 phylogenetic tree. A phylogenetic tree was constructed using the C. heterostrophus Lam1 protein sequence, and its top 50 blast matches retrieved from Genbank. See Fig. III.11 for annotation. 322 Figure S13 323 Figure S13 (continued) 324 Figure S13 (continued) 325 Figure S13 (continued) LAM1 326 Figure S14. Complete oxidoreductase Oxi1 phylogenetic tree. Full tree from Fig. III.12. 327 OXI1 Figure S15. Complete Tox9 phylogenetic tree. Full tree from Fig. III.13. 328 Figure S15 329 Figure S15 (continued) TOX9 330 Figure S16. Complete reductase phylogenetic tree. Full tree from Figs. III.15, III.16. 331 Figure S16 RED2 RED3 332 Figure S16 (continued) 333 Figure S16 (continued) 334 Figure S16 (Continued) 335 Figure S16 (continued) 336 Figure S16 (continued) RED1 337 B. Supplemental Methods 1. Mapping scaffolds onto the C. heterostrophus genetic map. i. Physical and genetic distance An RFLP map with 125 markers was constructed previously using C. heterostrophus race O field strain Hm540 and race T C-strain B30.A3.R.45 as parents (Fig. S1) [1]. When RFLP marker sequences were used as blast queries against the strain C5 genome assembly, 62 markers could be placed unambiguously. Placed markers were used to unite scaffolds and genetic linkage groups (Fig. II.1, Tables S1, S2). Twenty-nine scaffolds (denoted by internal JGI IDs, Table S2) were combined to create 16 genetically grounded scaffolds (30.6 Mb total), 14 of which could be oriented by the presence of multiple markers; 4.94 Mb of sequence on 28 scaffolds was left unplaced (Fig. II.1). In addition, 24 unplaced fosmids that could not be placed in the assembly, but which contained valid sequence, were added as individual scaffolds (864 kb). For markers that mapped to the same scaffold, genetic distance could be directly compared to physical distance. Twenty nine such pairwise comparisons were made, showing an average ratio of 13 kb/cM (ranging from 4.2-30 kb/cM). For a number of the linkage groups, the sum physical distance is very near previous estimates for chromosome size: assemblies for linkage groups 4, 7, 12, 13 and 14 are all within 300 kb of predictions based on Clamped Homogeneous Electric Field (CHEF) gels [1,3] (Table S1). Other assembled linkage groups have much more physical sequence than previously estimated. Chromosome 10, for example, estimated by CHEF gels to be 1.9 Mb, appears to be 3.4 Mb when assembled, with the caveat 338 that each scaffold is linked by a single marker only (the MAT locus and RFLP B285) (Figs. II.1, S1, Table S1). Chromosome 1, estimated to be 3.1 Mb but assembling to 4.2 Mb, is a more robust example, as each scaffold is linked by at least two RFLPs (Figs. II.1, S1, Table S1). There is a gross difference in physical size and genetic distance between markers B277 and B91 on this chromosome (Figures II.1, S1), however, suggesting that recombination is very rare between these two markers, or that a structural rearrangement has occurred on this chromosome in the sequenced C5 strain. Chromosomes 2 and 3, on the other hand, assemble to only 2.5 Mb, while CHEF methods estimate 3.7 and 3.6 Mb, respectively. For total C5 genome size, however, previous estimates (34.6 Mb, [3]) are very near that of the final assembly (36.46 Mb, Table S1, Figs. II.1, S1). Note that the total assembled genome size of isogenic C. heterostrophus strain C4 is 32.93 Mb. This is less than predicted since we know that a unique region (not in strain C5) encoding the Tox1 locus is ~1.2 Mb. Therefore, based on the C5 genome size, we estimate the C4 genome size should be ~ 36.46 + 1.2 = 37.66 Mb. Kodama et al [3] predict ~ 35.8 Mb. This discrepancy is likely due to the fact that C4 was sequenced using Illumina, an approach that results in underestimations of repeat content coupled with the fact that the 1.2 Mb Tox1 region contains an abundance of difficult to assemble repeats [4,5]. ii. Breakpoint linkage group Strains C5 and C4 are progeny of a backcross series selected on the basis of whether or not they carried the Tox1 locus responsible for T-toxin production [6]. As noted above, estimates from chromosome separation gels suggest that there is at least 1.2 Mb of DNA in race T strain C4 that is not present in race O strain C5 [1,3,4]. It has been shown previously that the Tox1 locus maps to the intersection of a four armed linkage group consisting of two race O chromosomes (chromosomes 6 and 12) and two race T chromosomes (chromosomes 6;12 and 339 12;6) that have undergone a reciprocal translocation with respect to the race O pair (Fig. S1) [1,3]. Mapping of scaffolds to the genetic map identified scaffolds 11027.2 and 11057 mapping to race O linkage group 6 (chromosome 6) and scaffolds 11053 and 11035 mapping to C5 linkage group 12 (chromosome 12), which were combined to form scaffolds 6 and 12 in the final assembly (Figs. II.1, S1). Chromosome 6 (1.3 Mb) of C5 is well covered by isogenic race T C4 scaffolds 13 (900 kb) and 33 (270 kb). Although both of these scaffolds aligned to a single chromosome (6) of C5, they are likely split onto the two reciprocally translocated chromosomes, 6:12 and 12:6, of C4 (Fig. S1). C5 chromosome 12 (1.8 Mb) is primarily covered by C4 scaffold 20 (56 kb), scaffold 29, (161 kb), scaffold 47 (146 kb), scaffold 55 (80 kb), and scaffold 11 (1 Mb). On both chromosome 6 and 12, there is a region between aligned C4 scaffolds where there is poor coverage (Fig. II.3). To date, no scaffold has been identified that clearly spans the breakpoints of the reciprocal translocations, likely due to the repetitive nature of Tox1-associated DNA. This leaves the exact physical positions of the 1.2 Mb of Tox1 DNA unresolved. iii. Dispensable chromosome Many markers mapped to linkage group B1 (scaffold 16, JGI ID 11041) which corresponds to the dispensable chromosome on the original map (Fig. S1). This chromosome is present in the parent race T strain B30.A3.R.45 [1] used to build the RFLP map and is also in reference race O strain C5. It was not, however, present in the second parent (Hm540) of the original genetic map. Because of this, there are no frequencies of crossing over to calculate genetic distance for this chromosome. When C. heterostrophus Hm540 was aligned to the C5 reference genome, only small, non-co-linear islands of the B1 chromosome could be mapped with Hm540 sequences, although 340 Hm540 sequences aligned well to other C5 chromosomes/scaffolds (Fig. S3). This gap was specific to C. heterostrophus Hm540, as strain C4, Hm338 and PR1x412 sequence reads could be mapped to this region. This supports the argument that chromosome B1 is a dispensable chromosome and that it is missing from strain Hm540 (Fig. S3). This chromosome did not meet dispensable chromosome criteria set in Ohm et al. [7], as the gene density (392.9 genes/Mb) is too high and repeat content (4.89%) too low. No known virulence factors map to the C. heterostrophus B chromosome. Thus, while the genetic evidence for chromosome B1 being dispensable is strong, it does not fit the pattern of several other dispensable chromosomes. There were no RFLP markers linking this scaffold to our map, and the JGI synteny browser allowed us to assess the presence and co-linearity of Scaffold 16 (chromosome B1) in other strains and species. Scaffold 16 (chromosome B1) is present, in its entirety, in isogenic strain C4 (scaffold 18), along with small regions (<1 kb) distributed on other scaffolds. When aligned to C. sativus, however, there is almost no co-linearity, although the majority of the scaffold has alignments, generally < 20 kb long (not shown). This pattern of reduced co-linearity of this chromosome/scaffold, compared to conservation for the rest of the genome is also evident when comparisons are made to other species examined here. iv. Telomeres Twelve candidate telomeric sequences were identified in the JGI C5 assembly out of the complete inventory of 32 when scaffolds were searched for repeats with telomeric sequence CCCTAA. Six were placed on mapped scaffolds, with only a single linkage group (14, scaffolds 11055 and 11038) having both telomeres. Six additional telomere calls were on unplaced scaffolds (S18, 23, 27, 50, 52, 57), however, there are no data linking these scaffolds to any particular chromosome. 341 2. Mapping scaffolds to the C. sativus genetic map To add more markers to the previously constructed C. sativus genetic map [8] and associate scaffolds with linkage groups, 121 polymorphic simple sequence repeat (SSR) markers were identified in the assembly sequences of the ND90Pr and ND93-1 parents. Of these, 106 segregated in a 1:1 ratio, while 15 exhibited distorted segregation, ten at the 5% and five at the 1%, significance level. A final genetic linkage map (Fig. S4) was constructed with 68 SSR markers and 140 previously mapped markers, including 102 amplified fragment length polymorphism (AFLP), 34 RFLP and two PCR markers, plus the mating type (MAT) and the VHv1 locus associated with virulence of pathotype 2 isolate ND90Pr on barley cv Bowman [8]. The markers (208 in total) were associated with 37 linkage groups which had at least two markers, when a minimum logarithm of odds (LOD) value of 4.0 and a maximum theta of 0.3 were used in the MAPMAKER program [9]. Since 30 of these linkage groups contained SSR markers, they could be associated with 16 scaffolds, summing to 29.32 Mb. Seven linkage groups were unassigned (Fig. S4). When DNA sequences of the two AFLP markers (E-AG/MCA-207 and E-AG/M-CG-121, Fig. S4), co-segregating with the virulence locus VHv1, were used as blast queries against the ND90Pr genome assembly, E-AG/M-CG-121 mapped to coordinates 2,132,630-2,132,734 of scaffold 5, while E-AG/M-CA-207 mapped to a thioesterase gene (protein ID 42084), unique to ND90Pr on scaffold 40, on a small contig (6,399 bp) that is likely linked to scaffold 5 because it carries the second AFLP marker that co-segregates with the VHv1 locus (Fig. S4). Comparison of the physical and genetic distances between 32 pairs of SSR markers that mapped to the same scaffold indicated that the ratios varied and ranged from 0.67 to 13.28 kb/cM with an average of 5.07 kb/cM). 342 3. Mating type region comparisons Cochliobolus, like other heterothallic Dothideomycetes, has two mating type idiomorphs [10,11], MAT1-1 and MAT1-2, and these were identified in all genomes. The C. heterostrophus reference C5, Hm540, and PR1x412, C. carbonum 26-R-13 and S. turcica strains were MAT1-1, while the others were MAT1-2. 10 kb regions flanking the MAT idiomorphs were aligned for each mating type. In all cases, the order of genes immediately surrounding the MAT locus was conserved (Fig. S5). Thus the larger collection of isolates confirms earlier data regarding which genes are encoded on the MAT flanks and that these are indeed different from genes flanking MAT loci in other ascomycetes [11]. S. turcica, not surprisingly, had the most variation in these regions compared to the Cochliobolus regions, however the same genes were present (Fig. S5). Although the MAT and flanking genes themselves were well conserved compared to those in the other species, 500 bp of the 5’ region and ~3 kb of the 3’ region (Fig. S5) were highly variable compared to these regions in Cochliobolus. There were other, smaller, gaps elsewhere in the 20 kb region analyzed. The number of SNPs called when aligning MAT regions (excluding the MAT genes themselves when comparing MAT1-1 to MAT1-2) was lowest when comparing within C. heterostrophus species (Table S11). When C. heterostrophus strains of the same mating type were compared, there were only 0 [C5 (MAT1-1) and PR1x412 (MAT1-1)], 15, [C5 (MAT1-1) and Hm540 (MAT1-1)] or 12 [C4 (MAT1-2) and Hm338 (MAT1-2)] SNPs called. Within the species, (i.e., comparing C. heterostrophus MAT1-1 flanks to MAT1-2 flanks), however, there were 100-200 SNP calls in each alignment. C. heterostrophus C4 and C5 are the most similar strains on the genome scale (Table II.3), and the average number of bps/SNP would predict only a single SNP across the 20 kb flanking regions outside the MAT genes. Instead, there were 91 343 between C4 and C5, on par with the other MAT1-1 flank to MAT1-2 flank comparisons across mating type within the species. For all other C. heterostrophus strains and Cochliobolus species aligned to C5, the number of SNPs called was slightly higher than the number predicted by the whole-genome average bps/SNP. Aligning the 20 kb of MAT flanking DNA across species produced 1500-1823 SNPs, regardless of whether or not the comparison was for regions carrying the same or different MAT genes, and over twice as many SNPs when aligned to S. turcica, when either C4 (MAT1-2) or C5 (MAT1-1) was used. C. carbonum (MAT1-1) and C. victoriae (MAT12) had only 121 SNPs when aligned to each other, fewer than comparing C. heterostrophus C5 (MAT1-1) to C. heterostrophus Hm338 (MAT1-2). The observation that the similarity between MAT regions of C. carbonum and C. victoriae is comparable to that of MAT loci within C. heterostrophus strains is consistent with their close phylogenetic relationship, and, ability to cross to one another. 344 C. References 1. 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