BACTERIAL ENDOSYMBIONTS OF ENDOPHYTIC FUNGI: DIVERSITY, PHYLOGENETIC STRUCTURE, AND BIOTIC INTERACTIONS by Michele Therese Hoffman _______________________
A Dissertation Submitted to the Faculty of the DEPARTMENT OF PLANT SCIENCES In Partial Fulfillment of the Requirements For the Degree of DOCTOR OF PHILOSOPHY WITH A MAJOR IN PLANT PATHOLOGY In the Graduate College THE UNIVERSITY OF ARIZONA 2010
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THE UNIVERSITY OF ARIZONA GRADUATE COLLEGE As members of the Dissertation Committee, we certify that we have read the dissertation prepared by Michele T. Hoffman entitled "Bacterial endosymbionts of endophytic fungi: diversity, phylogenetic structure, and biotic interactions.” and recommend that it be accepted as fulfilling the dissertation requirement for the Degree of Doctor of Philosophy. A. Elizabeth Arnold
_______________________________________________________ Date:
4/21/10
Judith L. Bronstein ________________________________________________________ Date: 4/21/10 Marc J. Orbach
___________________________________________________________
Date: 4/21/10
Final approval and acceptance of this dissertation is contingent upon the candidate’s submission of the final copies of the dissertation to the Graduate College. I hereby certify that I have read this dissertation prepared under my direction and recommend that it be accepted as fulfilling the dissertation requirement. A. Elizabeth Arnold ____________________________________ Date: 4/21/10 Dissertation Director
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STATEMENT BY AUTHOR This dissertation has been submitted in partial fulfillment of requirements for an advanced degree at the University of Arizona and is deposited in the University Library to be made available to borrowers under rules of the Library. Brief quotations from this dissertation are allowable without special permission, provided that accurate acknowledgement of source is made. Requests for permission for extended quotation from or reproduction of this manuscript in whole or in part may be granted by the head of the major department or the Dean of the Graduate College when in his or her judgment the proposed use of the material is in the interests of scholarship. In all other instances, however, permission must be obtained from the author.
SIGNED: Michele T. Hoffman
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ACKNOWLEDGMENTS This research would not have been possible without the assistance of many wonderful individuals. I wish to first thank my advisor A. Elizabeth (Betsy) Arnold for her support and guidance these past years. I am forever grateful to have had this opportunity to participate in this outstanding research program. I appreciated that I was given the freedom to explore my own research questions while being rigorously challenged as a scientist. I am also most grateful to the members of my committee, Judie Bronstein for your thoughtful questions and feedback, Marc Orbach for helping me troubleshoot research problems and Sandy Pierson, for all things prokaryotic. I have appreciated all your time, guidance and patience. I want to thank the School of Plant Sciences and the IGERT program for funding and the staff members in Plant Pathology and Microbiology for your kindness and good will. Numerous friends deserve my thanks and gratitude for their support, cheerful spirit, and camaraderie. I wish to especially thank Cara Gibson and members of her phylogenetics lunch club, Jeff Oliver and Chris Schmidt, along with fellow graduate students and faculty: Fabiola Santos Rodriguez, Anne Estes, David Maddison, Randy Ryan, David Bentley, Periasamy (Ravi) Chitrampalam, Scott Kroken, Mary Jane Epps, Jana U’ren, Rhodesia Celoy, Mariana Del Olmo Ruiz, David Jarvis, Gerard White, Baomin Wang, Mary Shimabukuro, Dylan Grippi, Andrea Woodard, Jacob Russell, Eric McDowell, and Bridget Barker. I could not have accomplished this work without the help and guidance of Mali Gunatilaka. You are a special soul Mali and I owe you one million thanks! Finally, I wish to thank my incredible friends and family for their love and support.
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DEDICATION This journey represents the culmination of many years of good intentions, perseverance, and the unwavering belief that dreams can and do eventually come true. I dedicate this work to Edie, the person who encouraged me to dream.
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TABLE OF CONTENTS ABSTRACT.........................................................................................................................7 INTRODUCTION...............................................................................................................9 1.1 Literature review.……………………………….…..................................................9 1.2 Explanation of dissertation format………………………………………………...15 PRESENT STUDY............................................................................................................17 2.1 Geographic locality and host identity shape fungal endophyte communities in cupressaceous trees……………………………………………………………………....17 2.2 Diverse bacterial endosymbionts inhabit living hyphae of phylogenetically diverse fungal endophytes …………………………………………………………...….18 2.3 IAA production by endophytic Pestalotiopsis neglecta is enhanced by an endohyphal bacterium .…………………………………..…………...……………....…18 REFERENCES…………………………………………………………………………..19 APPENDIX A: GEOGRAPHIC LOCALITY AND HOST INDENTITY SHAPE FUNGAL ENDOPHYTE COMMUNITIES IN CUPRESSACEOUS TREES...............24 APPENDIX B: DIVERSE BACTERIAL ENDOSYMBIONTS INHABIT LIVING HYPHAE OF PHYLOGENETICALLY DIVERSE FUNGAL ENDOPHYTES ….….49 APPENDIX C: IAA PRODUCTION BY ENDOPHYTIC PESTALOTIOPSIS NEGLECTA IS ENHANCED BY AN ENDOHYPHAL BACTERIUM..…………….99
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ABSTRACT
This dissertation comprises a series of studies designed to explore the associations between plants and the endophytic fungi they harbor in their above-ground tissues. By viewing endophyte diversity in ecologically and economically important hosts through the lenses of phylogenetic biology, microbiology, and biotechnology, this body of work links plant ecology with newly discovered symbiotic units comprised of endophytic fungi and the bacteria that inhabit them. This work begins with a large-scale survey of endophytic fungi from native and non-native Cupressaceae in Arizona and North Carolina. After isolating over 400 strains of endophytes, I inferred the evolutionary relationships among these fungi using both Bayesian and parsimony analyses. In addition to showing that native and introduced plants contained different endophytes, I found that the endophytes themselves harbor additional microbial symbionts, recovering members of the beta- and gammaproteobacterial orders Burkholderiales, Xanthomonadales, and Enterobacteriales and numerous novel, previously uncultured bacteria. This work finds that phylogenetically diverse bacterial endosymbionts occur within living hyphae of multiple major lineages of ascomycetous endophytes. A focus on 29 fungal/bacterial associations revealed that bacterial and fungal phylogenies are incongruent with each other and did not reflect the phylogenetic relationships of host plants. Instead, both endophyte and bacterial assemblages were strongly structured by geography, consistent with local horizontal transmission.
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Endophytes could be cured of their bacterial endosymbionts using antibiotics, providing a tractable experimental system for comparisons of growth and metabolite production under varying conditions. Studies of seven focal fungal/bacterial pairs showed that bacteria could significantly alter growth of fungi at different nutrient and temperature levels in vitro, and that different members of the same bacterial lineages interact with different fungi in different ways. Focusing on one isolate, I then describe for the first time the production of indole3-acetic acid (IAA) by a non-pathogenic, foliar endophytic fungus (Pestalotiopsis neglecta), suggesting a potential benefit to the host plant harboring this fungus. I show that this fungus is inhabited by an endohyphal bacterium (Luteibacter sp.) and demonstrate that mycelium containing this bacterium produces significantly more IAA in vitro than the fungus alone. I predict that the general biochemical pathway used by the fungal-endohyphal complex is L-tryptophan-dependent and measure effects of IAA production in vivo, focusing on root and shoot growth in tomato seedlings.
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INTRODUCTION
1.1 Literature review 1.1.1 Fungal endophytes All lineages of land plants surveyed to date harbor living fungi within their healthy, aboveground tissues (e.g., Carroll, 1988; Petrini, 1996; Davis et al., 2003; Cohen, 2004; Arnold & Lutzoni, 2007; Kauserud et al., 2008; Arnold et al., 2009). These endophytic fungi colonize and persist within living tissues such as leaves and stems without causing overt symptoms of disease (Petrini, 1991; Stone et al., 2000). Fungal endophytes are highly diverse at the species level (Carroll, 1995; Fröhlich & Hyde, 1999; Lu et al., 2004; Rodrigues et al., 2004; Arnold, 2007), represent every major lineage of non-lichenized Pezizomycotina (Higgins et al., 2007; Arnold, 2008), and have arisen multiple times in the evolution of Fungi (Spatafora et al., 2006; Arnold et al., 2009). In life history they range from the systemic, vertically transmitted clavicipitaceous endophytes that have coevolved with their cool-season grass hosts (Class 1 endophytes, Rodriguez et al., 2009) to highly localized, horizontally transmitted endophytes that associate at various levels of specificity with all major lineages of plants (Class 3 endophytes, Rodriguez et al., 2009). Among these, the horizontally transmitted, phylogenetically diverse Class 3 endophytes associated with foliage of long-lived perennial trees are of special interest: individual trees concurrently harbor dozens to hundreds of endophyte species within their asymptomatic photosynthetic tissues (Lodge et al., 1996; Müller & Hallaksela, 1998; Arnold et al., 2003; Göre & Bucak, 2007), raising questions regarding the nature of their
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interactions with hosts. These interactions appear to range from latent pathogenicity to latent saprotrophy, and encompass a variety of defensive mutualisms and physiological costs and benefits (Arnold, 2007). Such interactions change frequently over evolutionary time and may be variable and unpredictable in ecological time frames as well (Arnold, 2008; Arnold et al., 2009). The mechanisms driving variation in such interactions provide the context for my dissertation research. Whether variable outcomes reflect flexible genomic architecture in endophytes or their plant hosts, sensitivity of the interaction or either partner to biotic or abiotic stress, or additional microbial participants in species interactions is not yet clear. In this dissertation, I have focused on the third area, with special attention to the previously unexplored roles of endohyphal bacteria in shaping plant-endophyte interactions.
1.1.2 Endohyphal bacterial symbionts In addition to interacting with environmental and ectosymbiotic bacteria, some plantassociated fungi harbor bacteria within their hyphae (first noted as ‘bacteria-like organisms’ of unknown function by MacDonald and Chandler, 1981). These bacteria, best known from living hyphae of several species of Glomeromycota and Mucoromycotina, can alter the fungal phenotype (see Bianciotto et al., 1996; Bertaux et al., 2003; Bidartondo et al., 2004; de Boer et al., 2005; Waller et al., 2005; Shefferson et al., 2005). For example, Candidatus Glomeribacter gigasporarum colonizes spores and hyphae of the arbuscular mycorrhizal (AM) fungus Gigaspora gigasporarum (Bianciotto
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et al., 2000; Levy et al., 2003; Lumini et al., 2007). When cured of the bacterium, fungal growth and development are suppressed, and the morphology of the cell wall, vacuoles, and lipid bodies changes noticeably. Recovery of a P-transporter operon in Burkholderia sp. from Gigaspora margarita (Ruiz-Lozano & Bonfante, 1999), and the discovery of phosphate-solubilizing bacteria within Glomus mossae spores (Mirabal-Alonso & OrtegaDelgado, 2007), suggest a competitive role in phosphate acquisition and transport by these bacteria within the AM symbiosis. Recently, Partida-Martinez and Hertweck (2005) reported that a plant pathogen, Rhizopus microsporus (Mucoromycotina), harbors an endosymbiotic Burkholderia sp. that produces a phytotoxin (rhizoxin) responsible for the pathogenicity of the fungus. Each of these suggests a pervasive role of these endohyphal bacteria not only on their fungal hosts, but on plant-fungus interactions as well. To date, however, endohyphal bacteria have been studied only in rhizosphere-associated or plantpathogenic fungi. These studies have not encompassed some of the most species-rich fungal lineages, nor the ubiquitous, avirulent endophytic fungi found in above-ground plant tissues.
1.1.3 Endohyphal bacteria: hidden drivers of plant-endophyte interactions? In this dissertation, I demonstrate that foliar endophytes harbor diverse, apparently horizontally acquired, and facultative bacterial endosymbionts, and provide the first evidence for their effects on foliar endophytes and plant-endophyte interactions. I have focused on host plants representing the Cupressaceae (cedars and cypresses), an economically and ecologically important family of conifers. The family comprises 130-
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140 species of trees and shrubs in approximately 27-30 genera and encompasses the widest geographic range of any conifer family, with species ranging from 70°N to 55°S. Well-known species include coastal redwood (Sequoia sempervirens), giant sequoia (Sequoiadendron giganteum), and bald cypress (Taxodium spp.). In the mid-elevation woodlands and forests of Arizona and much of the intermountain west, Cupressaceae form the dominant component of tree communities. Species of Juniperus and Cupressus are especially common both naturally and as ornamentals, with species such as Platycladus orientalis (Oriental arbor-vitae) commonly planted as well. Endophytic fungi have been examined in a small number of cupressaceous hosts previously (Carroll 1978, 1995; Camacho et al., 1997; Deckert, 2000; Hata, 1996; Arnold et al., 2007), but never in an explicit geographic context, nor with the geographic breadth and depth of the present work. I first address the degree to which endophytic fungal associates of trees in the Cupressaceae differ across the geographic ranges of hosts. In this work I examine the role of native vs. non-native status in shaping endophyte communities as well. My goal is to understand the intrinsic variation in fungal communities encountered and acquired by plants in different geographic areas. Using both genotype-level and phylogenetic analyses, I show that (1) conspecific trees in widely separated geographic areas (southwestern vs. southeastern US) harbor different endophyte communities, and (2) that native and introduced trees in the same geographic areas differ in their endophyte assemblages. That study yielded a tremendous diversity of previously unknown endophytic fungi and a new understanding of their phylogenetic relationships.
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Next, I examined the potential for such endophytic fungi to harbor endohyphal bacteria. For this work, I concentrated on fungi associated with healthy foliage of Cupressaceae at a smaller geographic scale, with a particular focus on exploring fungal communities associated with plants on multiple mountain ranges within Arizona. After extracting total genomic DNA from several axenic endophyte cultures, I discovered the presence of measurable bacterial DNA within those DNA samples. After ruling out contamination (Hoffman and Arnold, 2010), I sequenced the 16S ribosomal DNA region of representative strains and recovered members of the beta- and gamma-proteobacterial orders Burkholderiales, Xanthomonadales, and Enterobacteriales, including putative Ralstonia pickettii, Variovorax paradoxus, Luteibacter rhizovicina, Pantoea spp., and numerous novel, previously uncultured bacteria. I localized these bacteria within hyphae and confirmed the viability of both the bacteria and the hyphae harboring them using Molecular Probes Live-Dead stain. Phylogenetic analyses of fungi and their associated bacteria then provided strong evidence that bacteria can occur within endophytic members of four classes of Ascomycota. My work suggests that these are facultative associates of their fungal hosts, and highlights the lack of congruence among bacterial, fungal, and host plant phylogenies. After developing methods to cure fungi of their endohyphal bacteria, I examined the effects of bacterial endosymbionts on endophytic growth rates in vitro. Specifically, by cultivating fungi on growth media containing the antibiotic ciprofloxacin, I produced fungal cultivars that were confirmed by microscopy and PCR to lack endohyphal bacteria (Hoffman and Arnold, 2010). By comparing the growth of replicate clones with and
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without bacteria on various media, at different temperatures and at different pH levels, I found that these bacteria had variable and sometimes unpredictable effects. One strain, 9143, proved especially interesting because (1) I could identify the both the fungus and bacterium with reasonable certainty using morphological and phylogenetic methods, (2) I could separate the fungal and bacterial partners in vitro, and (3) this particular genus (Pestalotiopsis) is well known for its production of numerous medicinally important secondary metabolites. The ability to synthesize phytohormones, including indole-3-acetic acid (IAA), gibberellins, ethylene, cytokinins, jasmonic acid and abscisic acid, have been documented in numerous bacteria and fungi (Costacurta and Vanderleyden, 1995; Glick, 1995; Lindow et al., 1998; Robinson et al., 1998; Koga, 1995; Maor et al., 2004; Sirrenberg et al., 2007). Some of these microorganisms utilize a L-tryptophan-dependent pathway for IAA production, whereas others utilize a L-tryptophan independent pathway (Tudzynski and Sharon, 2002; Buchanan et al., 2005). My investigation of IAA production in infected endophytes found that some isolates were capable of IAA production when supplied with L-tryptophan and that the infected endophytes produced significantly higher levels of IAA than the cured counterpart, under identical growth conditions. Culture filtrate from the endophyte-bacterium complex significantly increased root growth in vitro in seedlings of tomato relative both to controls and to filtrate from the endophyte alone. These data suggest that this newly described microbial partnership can manipulate plant biochemistry with a positive effect.
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1.2 Explanation of dissertation format The goals of this dissertation are to explore the interactions of endophytic fungi with other microorganisms and their plant hosts, and to expand our knowledge of endophyte diversity, community structure, and ecological roles in nature. I introduce this dissertation research, methods and results as three appendices. In Appendix A, I examine the abundance, diversity, and species composition of endophytic fungi in pairs of related species of Cupressaceae from a mesic forest (central North Carolina, USA) and from a xeric desert environment (southern Arizona, USA), Utilizing a culture-based approach, I examined the fungal communities of two native species (Cupressus arizonica, AZ and Juniperus virginiana, NC) and a co-occurring, non-native species (Platycladus orientalis in both AZ and NC). My results lay the groundwork for addressing relative importance of locality, host species, and native/nonnative status in endophyte community structure. In Appendix B, I examine evidence of endohyphal bacteria associated with these fungal endophytes and began to elucidate the infection frequency, taxonomy, and phylogenetic relationships of these cryptic microorganisms. This work reports, for the first time, the occurrence of endohyphal bacteria in foliar endophytes and highlights their occurrence in four classes of Pezizomycotina. I recover novel lineages of endohyphal bacteria relative to previous studies and suggest that the association of these bacteria and their endophytic hosts is facultative, reflecting (1) loss of bacterial endosymbionts in culture over time, and (2) incongruent fungal- and bacterial phylogenies. These analyses
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set the stage for experimental work addressing the ecological importance of these previously unknown symbionts. The data presented in Appendix C are the results of in vitro and in planta experiments, conducted in collaboration with Malkanthi K. Gunatilaka, E. M. Kithsiri Wijeratne, and A. A. Leslie Gunatilaka, measuring phytohormone production in a focal endophytic fungus. This work demonstrates, for the first time, the ability of a foliar endophyte to produce indole-3-acetic-acid (IAA), a phytohormone with strong effects on plant cell growth and elongation. Moreover, we provide the first documentation of a role of an endohyphal bacterium in enhancing IAA production, and demonstrate a strong effect on seedling growth. In general, this work begins to answer the question: how do bacterial endosymbionts influence the outcome of plant-fungal symbioses?
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PRESENT STUDY
The methods, results and conclusions from this research are presented in the appended manuscripts. The following is a summary of the most important findings from each. 2.1
Geographic locality and host identity shape fungal endophyte communities in
cupressaceous trees Fungal endophytes, found in every major lineage of the most species-rich group of fungi (Ascomycota), are believed to represent a significant proportion of Earth’s fungal diversity. Using 95% sequence similarity as a proxy for species recognition, I report a high level of species richness in three species of Cupressaceae, an economically and ecologically important family of conifers. In addition to isolating over 400 strains of endophytes, I inferred the evolutionary relationships among these fungi for the first time using both Bayesian and parsimony analyses. Analyses of the nuclear ribosomal internal transcribed spacer regions (ITS rDNA) for 109 representative isolates showed that diversity was two times greater in plants from North Carolina vs. Arizona, and that endophytes recovered in Arizona were more likely to be host-generalists relative to those in North Carolina. Overall abundance, diversity, and taxonomic composition of these endophyte communities differed as a function of host identity, native vs. non-native status, and locality.
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2.2
Diverse bacterial endosymbionts inhabit living hyphae of phylogenetically
diverse fungal endophytes Here I report that phylogenetically diverse bacteria occur within living hyphae of endophytic fungi, including members of multiple major lineages of Ascomycota. Drawing from surveys of endophytes in healthy photosynthetic tissues of ecologically and economically important trees in five biogeographic provinces, I show that bacterial and fungal phylogenies are not congruent and do not reflect the evolutionary relationships of the host plants they both inhabit. Instead, both endophyte and bacterial endosymbiont species appear to be horizontally transmitted and are structured more by geography than by host identity.
2.3 IAA production by endophytic Pestalotiopsis neglecta is enhanced by an endohyphal bacterium
In this section, I demonstrate the effects of a representative endohyphal bacterium, identified using phylogenetic analyses as Luteibacter sp., on growth rates of endophytic Pestalotiopsis neglecta under various nutrient, temperature and pH conditions. I show that this fungal endophyte is capable of producing the phytohormone indole-3-acetic acid (IAA) in vitro. I report a significant difference in production of IAA for the infected vs. cured and that this same compound could positively influence the growth rate of tomato seedlings.
19 REFERENCES Arnold, A. E., Miadlikowska, J., Higgins, K. L., Sarvate, S. D., Gugger, P., Way, A., Hofstetter, V., Kauff, F. & Lutzoni, F. (2009). Hyperdiverse fungal endophytes and endolichenic fungi elucidate the evolution of major ecological modes in the Ascomycota. Systematic Biology 58: 283-297. Arnold, A.E. (2008). Endophytic fungi: Hidden components of tropical community ecology. Tropical Forest Community Ecology eds. Walter P. Carson & Stefan A. Schnitzer, Wiley-Blackwell West Sussex, UK. Arnold, A. E. (2007). Understanding the diversity of foliar endophytic fungi: Progress, challenges, and frontiers. Fungal Biology Reviews 21: 51-66. Arnold, A. E. & Lutzoni, F. (2007). Diversity and host range of foliar fungal endophytes: Are tropical leaves biodiversity hotspots? Ecology 88: 541-549. Arnold, A. E., Henk, D. A., Eells, R., Lutzoni, F. & Vilgalys, R. (2007). Diversity and phylogenic affinities of foliar fungal endophytes in loblolly pine inferred by culturing and environmental PCR. Mycologia 99:185-206. Arnold, A. E., Mejía, L., Kyllo, D., Rojas, E., Maynard, Z., Robbins, N. A. & Herre, E. A. (2003). Fungal endophytes limit pathogen damage in a tropical tree. Proceedings of the National Academy of Sciences USA 100: 15649–15654. Bertaux, J. et al. (2003). In situ identification of intracellular bacteria related to Paenibacillus spp. in the mycelium of the ectomycorrhizal fungus Laccarria bicolor S238N. Applied and Environmental Microbiology 69 (7): 4243-4248. Bianciotto, V., Bandi, C., Minerdi, D., Sironi, M., Tichy, H.V., and Bonfante, P. (1996). An obligate endosymbiont mycorrhizal fungus itself harbors oligately intracellular bacteria. Applied and Environmental Microbiology 62(8): 3005-3010. Bianciotto, V. at al. (2000). Detection and identification of bacterial endosymbionts in arbuscular mycorrhizal fungi belonging to the family Gigasporaceae. Applied and Environmental Microbiology 66 (10): 4503-4509. Bianciotto, V., Lumini, E., Bonfante, P. and Vandamme, P. (2003). ‘Candidatus Glomeribacter gigasporarum’ gen. nov., sp. nov., an endosymbiont of arbuscular mycorrhizal fungi. Int J Syst Evol Microbiol 53:121-124. Bidartondo, M. I. et al. (2004). Changing partners in the dark: isotopic and molecular evidence of ectomycorrhizal liaisons between forest orchids and trees. Proc. R. Soc. London B 271: 1799-1806.
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Lodge D.J., Fisher, P.J. and Sutton, B.C. (1996). Endophytic fungi of Manikara bidentata leaves in Puerto Rico. Mycologia 88(5): 733-738. Lu, G.Z, Cannon, P.F., Reid A., and Simmons, C.M. (2004). Diversity and molecular relationships of endophytic Colletotrichum isolates from the Iwokrama Forest Reserve, Guyana. Mycological Research 108: 53-63. Lumini, E. et al. (2007). Presymbiotic growth and sporal morphology are affected in the arbuscular mycorrhizal fungus Gigaspora margarita cured of its endobacteria. Cellular Microbiology 9(7): 1716-1729. MacDonald, R.M. and Chandler, M.R. (1981). Bacterium-like organelles in the vesiculararbuscular mycorrhizal fungus Glomus caledonius. New Phytologist 89: 241-246. Maor, R., Haskin, S., Levi-Kedmi, H., and Sharon, A. (2004). In planta production of indole-3-acetic acid by Colletotrichum gloeosporioides f. sp. aeschynomene. Applied and Environmental Microbiology 70(3):1852-1854. Mayo, K., Davis, R.E., and Motta, J. (1986). Stimulation of germination of spores of Glomus versiforme by spore-associated bacteria. Mycologia 78(3): 426-431. Minerdi, D. et al., (2008). Bacterial ectosymbionts and virulence silencing in a Fusarium oxysporum strain. Environmental Microbiology 10(7):1725-1741. Mirabal-Alonso, L. & Ortega-Delgado, E. (2007). Phosphate solubilizing bacteria isolated from the inside of Glomus mosseae spores from Cuba. First International Meeting on Microbial Phosphate Solubilization. Series: Developments in Plant and Soil Sciences , Vol. 102 Velazquez, E.; Rodriguez-Barrueco, C. (Eds.) Reprinted from Plant and Soil, Vol. 287, 1-84 Müller, M.M. and Hallaksela, A-M. (1998). Diversity of Norway spruce needle endophytes in various mixed and pure Norway spruce stands. Mycological Research 102(10): 1183-1189. Partida-Martinez, L.P. & Hertweck, C. (2005). Pathogenic fungus harbours endosymbiotic bacteria for toxin production. Nature 437:884-888. Petrini, O. (1996). Ecological and physiological aspects of host-specificity in endophytic fungi. In: Endophytic Fungi in Grasses and Woody Plants (S. C. Redlin & L. M. Carris, eds): 87–100. APS Press, St. Paul, Mn. Petrini, O. (1991). In: Microbial Ecology of Leaves, eds. Andrews, J. H. & Hirano, S. S. (Springer, New York), pp. 179–197. Robinson, M., Riov, J., and Sharon, A. (1998). Indole-3-acetic acid biosynthesis in Colletotrichum gloeosporioides f. sp. aeschynomene. Applied and Environmental Microbiology 64(12):5030-5032.
23 Rodrigues, K. F., Sieber, T. N., Gruenig, C. R. & Holdenrieder, O. (2004). Characterization of Guignardia mangiferae isolated from tropical plants based on morphology, ISSR-PCR amplifications and ITS1-5.8S-ITS2 sequences. Mycological Research 108: 45–52. Rodriguez, R.J., White, J.F., Arnold, A.E., and Redman, R.S. (2009). Fungal endophytes: Diversity and functional roles. New Phytologist 182:314-330. Ruiz-Lozano, J.M. and Bonfante, P. (1999). Identification of a putative P-transporter operon in the genome of a Burkholderia strain living inside the arbuscular mycorrhizal fungus Gigaspora margarita. Journal of Bacteriology 181:4106-4109. Shefferson, R. P., et al., (2005). High Specificity generally characterizes mycorrhizal associations in rare lady’s slipper orchids, genus Cypripedium. Molecular Ecology 14: 613-626. Sirrenberg, A. Göbel, C., Grond, S., Czempinski, N., Ratzinger, A., Karlovsky, P., Santos, P., Feussner, I. and Pawlowski, K. (2007). Piriformospora indica affects plant growth by auxin. Physiologia Plantarum 131:581-589. Spatafora, J. W., Sung, G. H., Johnson, D., et al. (2007). A five-gene phylogeny of Pezizomycotina. Mycologia 98(6): 1018-1028. Stanosz, G.R., Blodgett, J.T., Smith, D.R., and Kruger, E.L. (2001). Water stress and Sphaeropsis sapinea as a latent pathogen of red pine seedlings. New Phytologist 149: 531-538. Stone, J. K., Bacon, C. W. & White, J. F., Jr. (2000). An overview of endophytic microbes: Endophytism defined. In: Bacon, C. W., White, J. F., Jr. (Eds.) Microbial Endophytes. Marcel Dekker, New York. 3–29. Tudzynski, B. and Sharon, A. (2002). Biosynthesis, biological role and application of fungal phytohormones. The Mycota X. Industrial Applications. Springer-Verlag Berlin Heidelberg. Waller, F., B. Achatz, H. Baltruschat, J. Fodor, K. Becker, M. Fischer, T. Heier, R. Hückelhoven, C. Neumann, D. von Wettstein, P. Franken, and K-H. Kogel (2005). The endophytic fungus Piriformospora indica reprograms barley to salt-stress tolerance, disease resistance, and higher yield. Proc. Natl. Acad. Sci. USA 102:13386–13391.
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APPENDIX A
GEOGRAPHIC LOCALITY AND HOST INDENTITY SHAPE FUNGAL ENDOPHYTE COMMUNITIES IN CUPRESSACEOUS TREES
25 Dear Ms Hoffman, Thank you for your email. Please see our author pages http://www.elsevier.com/wps/find/authorsview.authors/copyright#whatrights , on author rights, which explains that you are able to use your paper as part of a dissertation. Best wishes Hannah Forder Journal Manager Pp Caroline Peters, Journal Manager -----Original Message----From:
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[email protected] Subject: permission to publish Please do not reply to this automated email. First Name Sender: Michele Last Name Sender: Hoffman Email Address:
[email protected] Subject: permission to publish Journal ID: 707043 Journal Title: Mycological Research Text of Email: Dear Sir or Madam, I wish to obtain permission from your journal to reprint the following publication as part of my forthcoming dissertation. Thank you for your time and consideration. Michele Hoffman Hoffman, M. T. and A. E. Arnold. 2008. Geographic locality and host identity shape fungal endophyte communities in cupressaceous trees. Myco. Res. 112:331â344.
mycological research 112 (2008) 331–344
26
journal homepage: www.elsevier.com/locate/mycres
Geographic locality and host identity shape fungal endophyte communities in cupressaceous trees Michele T. HOFFMAN, A. Elizabeth ARNOLD* Division of Plant Pathology and Microbiology, Department of Plant Sciences, 1140 East South Campus Drive, University of Arizona, Tucson, AZ 85721, USA
article info
abstract
Article history:
Understanding how fungal endophyte communities differ in abundance, diversity,
Received 13 March 2006
taxonomic composition, and host affinity over the geographic ranges of their hosts is key
Received in revised form
to understanding the ecology and evolutionary context of endophyte–plant associations.
28 August 2007
We examined endophytes associated with healthy photosynthetic tissues of three closely
Accepted 24 October 2007
related tree species in the Cupressaceae (Coniferales): two native species within their natural
Corresponding Editor: Barbara Schulz
ranges [Juniperus virginiana in a mesic semideciduous forest, North Carolina (NC); Cupressus arizonica, under xeric conditions, Arizona (AZ)], and a non-native species planted in each
Keywords:
site (Platycladus orientalis). Endophytes were recovered from 229 of 960 tissue segments
Ascomycota
and represented at least 35 species of Ascomycota. Isolation frequency was more than three-
Biodiversity
fold greater for plants in NC than in AZ, and was 2.5 (AZ) to four (NC) times greater for
Cupressaceae
non-native Platycladus than for Cupressus or Juniperus. Analyses of ITS rDNA for 109 repre-
Molecular phylogeny
sentative isolates showed that endophyte diversity was more than twofold greater in NC
Species richness
than in AZ, and that endophytes recovered in AZ were more likely to be host-generalists
Symbiosis
relative to those in NC. Different endophyte genera dominated the assemblages of each host species/locality combination, but in both localities, Platycladus harboured less diverse and more cosmopolitan endophytes than did either native host. Parsimony and Bayesian analyses for four classes of Ascomycota (Dothideomycetes, Sordariomycetes, Pezizomycetes, Eurotiomycetes) based on LSU rDNA data (ca 1.2 kb) showed that well-supported clades of endophytes frequently contained representatives of a single locality or host species, underscoring the importance of both geography and host identity in shaping a given plant’s endophyte community. Together, our data show that not only do the abundance, diversity, and taxonomic composition of endophyte communities differ as a function of host identity and locality, but that host affinities of those communities are variable as well. ª 2007 The British Mycological Society. Published by Elsevier Ltd. All rights reserved.
Introduction Recent studies have begun to elucidate the diversity and ubiquity of fungal endophytes associated with terrestrial plants, providing a framework for understanding the evolutionary context of these plant–fungus associations (reviewed by Stone et al. 2000; Schulz & Boyle 2005; Arnold & Lutzoni 2007).
Despite a strong foundation laid by a growing number of surveys, the degree to which endophytes differ in abundance, diversity, taxonomic composition, and host affinity over the geographic ranges of particular host lineages remains unclear. Resolving the importance of locality and host identity in shaping these aspects of endophyte communities is important for understanding fundamental aspects of endophyte symbioses.
* Corresponding author. E-mail address:
[email protected] 0953-7562/$ – see front matter ª 2007 The British Mycological Society. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.mycres.2007.10.014
332
Previous studies have indicated that the abundance, diversity, and species composition of endophytes can be strongly influenced by the locality in which a given plant occurs (e.g. Carroll & Carroll 1978; Petrini et al. 1982; Bills & Polishook 1992; Fisher et al. 1994; Hata & Futai 1996; Bayman et al. 1998; Arnold et al. 2001; Higgins et al. 2007). For example, different microhabitats vary in inoculum potential, with higher infection frequencies observed beneath forest canopies versus in forest clearings (Arnold & Herre 2003) or in more mesic microhabitats relative to proximate sites that are more xeric (Petrini et al. 1982; Petrini 1996). At larger geographic scales, endophytes are frequently more abundant in tropical regions than in boreal, arctic, or some temperate regions (Arnold & Lutzoni 2007), although local climate, short-term weather patterns, and some disturbance regimes can override latitudinal gradients (see Suryanarayanan et al. 2000; Arnold et al. 2007; Higgins et al. 2007). Similarly, diversity of endophytes differs at small scales as a function of land use history, vegetation cover, and other factors. For example, Gamboa & Bayman (2001) found higher endophyte diversity in leaves of Guarea guidonia in a forest preserve relative to a disturbed forest area in Puerto Rico. At larger scales, endophyte diversity varies as a function of latitude and annual rainfall (Arnold & Lutzoni 2007), although the contributions of co-varying factors, such as plant diversity, remain to be explored. Finally, the species composition of endophyte assemblages also differs as a function of localities. For example, Arnold et al. (2003) found distinctive endophyte communities associated with Theobroma cacao at multiple sites in Panama. In a study of endophytes and saprotrophs associated with closely related palms in Australia and Brunei Darussalam, Fro¨hlich & Hyde (1999) found that only 27 of 189 fungal species were shared between localities. Similarly, Fisher et al. (1995) found that only two of 25 species of endophytes recovered from Dryas octopetala (Rosaceae) in Switzerland also occurred in the same host in Norway. By comparing endophyte assemblages in a single host species or genus in multiple sites, these studies have shown that plants interact with distinctive endophyte communities across the geographic areas in which they grow. However, it is unclear whether plants growing in different localities experience not only a distinctive abundance, diversity, and species assemblage of endophytes, but also a different degree of relative host specificity. Do sympatric plant species differ in terms of the relative host specificity or host-generalism of the endophytes they harbour? Do different regions or environmental conditions select for more or less host-specific endophytes? To our knowledge, these questions have not been addressed previously. Most studies examining host affinities of endophytes have focused on distantly related host species that co-occur within particular geographic areas, and have yielded conflicting results. For example, a study involving endophytes of mistletoe and fir established that despite close physical proximity of these two plant taxa, endophyte communities remained exclusively associated with their particular host (Petrini 1996). Similarly, Suryanarayanan et al. (2000) found distinctive endophytes in Cuscuta reflexa relative to its host plants, and Arnold et al. (2000) found in lowland Panama that distantly related hosts, growing within metres of one another, bore distinctive
27
M. T. Hoffman, A. E. Arnold
endophyte communities. Yet Cannon & Simmons (2002) recovered similar endophyte communities from phylogenetically diverse angiosperm trees in a tropical forest reserve. Suryanarayanan et al. (2004) recovered the same endophyte species from diverse plant species in mangroves, dry deciduous forest, and other tropical forest types (Phyllosticta capitalensis), and Mohali et al. (2005) found that Lasiodiplodia theobromae occurs in multiple hosts and sites at a global scale. Suryanarayanan et al. (2005) found no evidence for host specificity among endophytes of cacti in the southwestern USA, yet Higgins et al. (2007) found that most genotypes of endophytes recovered from arctic and boreal plants were associated with only one host species. Thus, research to date remains in conflict with regard to the prevalence of host specificity among endophytes, and the potential for host specificity of endophyte communities to differ over the range of a plant taxon, or among species within a particular site, is not clear. To our knowledge, no study to date has concurrently assessed the importance of host identity and locality in shaping endophyte abundance, diversity, taxon composition, and relative host specificity by simultaneously sampling pairs of closely related plants in geographically distinct sites. Molecular tools are especially useful in this regard, providing a framework for explicitly comparing sterile endophytes and providing a tool to disentangle potentially cryptic species for which morphological characters are of limited use (Lacap et al. 2003; Arnold et al. 2007). In turn, phylogenetic analyses provide insight into the evolutionary associations of fungal lineages with hosts and localities, and can provide muchneeded insight into the degree to which particular clades of fungi are more or less likely to form host-specific associations with plants. Unfortunately, endophyte surveys are often restricted in terms of molecular phylogenetics due to the cost associated with sequencing informative loci for numerous and often phylogenetically diverse isolates (see Arnold et al. 2007; Higgins et al. 2007). In particular, there is a need to examine the ability of particular loci to infer topologies that are consistent with reconstructions provided by multi-locus analyses (e.g., Lutzoni et al. 2004; James et al. 2006) and to maximize the phylogenetic power provided by loci that can be sequenced rapidly and at reasonable cost. Using pairs of related species of the conifer family Cupressaceae from a mesic forest [central North Carolina (NC)] and from a xeric desert environment [southern Arizona (AZ)], we used a culture-based approach to examine the foliar endophyte communities of two native species (Cupressus arizonica, AZ and Juniperus virginiana, NC) and a co-occurring, non-native species (Platycladus orientalis in both AZ and NC). Our objectives were to: (1) examine the abundance, diversity, taxonomic composition, and host preferences of endophytic fungi of these cupressaceous hosts; (2) infer the relationships of endophytes associated with these trees in a broad phylogenetic context; (3) evaluate an accelerated phylogenetic search strategy for rapidly and robustly diagnosing these phylogenetic relationships using the LRORLR7 region of the LSU rDNA; and (4) generate hypotheses regarding the relative importance of geographic locality and host identity in structuring the endophyte communities associated with these ecologically and economically important trees.
28
Endophytes of Cupressaceae
Materials and methods Host species The cypress family, Cupressaceae (Coniferales), is widely distributed in warm-temperate regions and contains 110–130 species of evergreen trees and shrubs in 25–30 genera (Fralish & Franklin 2002). Within the Cupressoideae, the genera Cupressus and Juniperus comprise 70–86 species, whereas Platycladus is monotypic (Platycladus orientalis). Cupressus arizonica (AZ cypress) is indigenous in the western United States and northern Mexico (Little 1950; USDA, NRCS 2004). Juniperus virginiana (Eastern red cedar) is distributed from Nova Scotia to Texas, and is the most widespread conifer in the eastern USA (Fralish & Franklin 2002). Platycladus orientalis is native to Korea, eastern Russia, and China. It has been extensively cultivated in southeastern Asia and has become naturalized in Florida (Little 1980; USDA, NRCS 2004). It is commonly planted as an ornamental in both AZ and NC.
Sample collection In March and April 2005, fresh, asymptomatic photosynthetic tissue was collected from healthy trees at the campus arboretum at the University of AZ (Tucson: Platycladus and Cupressus; four individuals/species) and at Duke Forest, Duke University (Durham, NC: Platycladus and Juniperus; four individuals/species). The University of AZ (32.231 N, 110.952 W, elevation 787 m; mean annual temperature ¼ 20.2 C) is located within the Sonoran Desert bioregion. This site is relatively xeric, receiving an average of 30.5 cm of precipitation annually. Durham, NC (36.001 N, 78.940 W, elevation 97 m; mean annual temperature ¼ 15.5 C) is relatively mesic, receiving an average of 109.2 cm of precipitation annually. At the University of AZ, all focal plants were within 250 m of one another, and were cultivated in mixed, open stands with a variety of native and non-native ornamental plants. Focal individuals were fully established and mature, and received minimal supplemental water through infrequent irrigation. At Duke University, all Juniperus individuals were located beneath an open canopy of Pinus taeda interspersed with Liquidambar styraciflua, Carya spp., Quercus spp., Platanus occidentalis, and Acer spp. At that site, all Platycladus individuals were located within 200 m of native J. virginiana and were planted adjacent to the forest edge. From each individual, we harvested mature photosynthetic tissues from three branches extending in three randomly chosen cardinal directions. Material was bagged for transport to the laboratory and was used in endophyte isolations within 4 h of collection.
Endophyte isolations Photosynthetic tissue from each sample was rinsed in running tap water for 30 s and then cut into 2 mm pieces before surface-sterilization. Tissue fragments were agitated in 95 % ethanol for 30 s, 10 % bleach (0.5 % NaOCl) for 2 min, and 70 % ethanol for 2 min (Arnold et al. 2007). Under sterile conditions, tissue segments were allowed to surface-dry before
333
plating on 2 % malt extract agar (MEA). To confirm the efficacy of surface-sterilization, a subset of tissue segments was pressed against the medium under sterile conditions for 30 s, and then removed. No mycelial growth was observed from these surface impressions. A total of 384 tissue segments (96 per individual tree) were plated for each species from AZ, and 96 tissue segments (24 per individual tree) were plated for each species from NC, corresponding to pilot data regarding differences in infection frequency (Arnold & Lutzoni 2007). Tissue segments were incubated on sealed plates under ambient light/dark conditions at room temperature (ca 21.5 C). Hyphal growth was monitored over an eight-week period. Using aseptic technique, fungi were transferred to axenic culture on 2 % MEA in 60 mm Petri plates and photographed after one week of growth. All samples were archived as living vouchers in sterile water at the Robert L. Gilbertson Mycological Herbarium at the University of AZ. Based on mycelial characteristics (Arnold et al. 2000), representatives of all unique morphotypes were selected for molecular analysis.
Genomic DNA extraction and PCR Total genomic DNA was extracted directly from pure cultures following Arnold et al. (2007). PCR was used to amplify the ITS rDNA and 5.8s gene (ca 600 bp) and LSU rDNA (ca 1.2 kb) following Higgins et al. (2007). Primers included ITS1F or ITS5 and ITS4 for ITS rDNA (White et al. 1990; Vilgalys & Hester 1990; Gardes & Bruns 1993), and LROR and LR7 for LSU rDNA (Vilgalys, unpubl.; Vilgalys & Hester 1990). Sigma Readymix REDTaq PCR reaction mix with MgCl2 (St. Louis, MO) was used for all PCR reactions. The 25 ml reaction mixture included 12.5 ml REDTaq, 1 ml of each primer (10 mM), 1 ml DNA template, and 9.5 ml PCR-quality water. Cycling reactions were run on an MJ Research PTC200 thermocycler (Waltham, MA) with the following protocol: 94 C for 3 min; 36 cycles of 94 C for 30 s, 54 C for 30 s, 72 C for 1 min; and 72 C for 10 min. SYBR Green I stain (Molecular Probes, Invitrogen; Carlsbad, CA) was used to detect DNA bands on a 1 % agarose gel. All products demonstrated single bands.
Sequencing and analyses PCR products were cleaned, quantified, and normalized at the GATC sequencing facility at the University of AZ. Sequencing was performed on the Applied Biosystems 3730XL DNA Analyser (Foster City, CA). The software applications phred and phrap (Ewing & Green 1998; Ewing et al. 1998) were used to call bases and assemble contigs, with automation provided by the ChromaSeq package (Maddison & Maddison: http:// mesquiteproject.org) implemented in Mesquite v. 1.06 (Maddison & Maddison: http://mesquiteproject.org). All base calls were verified by inspection of chromatograms in Sequencher version 4.5 (Gene Codes, Ann Arbor, MI). ITS rDNA sequences were obtained for 109 representative isolates (Supplementary Material Appendix 1). LSU rDNA sequences were obtained for 64 representative isolates (Supplementary Material Appendix 2) based on ITS rDNA genotype groups (described below). Sequence data generated for this study are available from GenBank under accession numbers
29
334
EF419892–EF420019 (ITS rDNA sequences) and EF420020– EF420083 (LSU rDNA sequences).
Endophyte richness, diversity, and similarity among hosts and sites ITS rDNA sequence data were used in BLAST searches of GenBank to provide preliminary identification at higher taxonomic levels and to guide taxon sampling for LSU rDNA analyses. To designate operational taxonomic units (OTU) based on ITS rDNA data, we used Sequencher to delimit groups corresponding to 90, 95, and 99 % ITS rDNA similarity without considering differences in sequence length (Arnold et al. 2007). Previous studies have used 90–97 % ITS rDNA sequence similarity as a proxy for species boundaries in Fungi (e.g. 97 %: O’Brien et al. 2005). One study has explicitly shown that groups based on 90 % ITS rDNA similarity are highly congruent with phylotypes based on a second locus (Arnold et al. 2007). Based on these studies, we used 90 % ITS rDNA genotype groups as a highly conservative proxy for species in all subsequent analyses. Species accumulation curves and bootstrap estimates of total species richness were inferred using EstimateS v. 7.5 (Colwell 2005: http://viceroy.eeb.uconn.edu/EstimateS). In addition to assessing richness (number of species) for each partition of the data, we also calculated diversity, which takes into account the abundance of different species. Diversity was measured using Fisher’s a (Fisher et al. 1943), which is robust for small sample sizes and thus accommodates a wide range of sampling effort (Higgins et al. 2007). All ITS rDNA genotypes (based on 90 % sequence similarity) that were recovered more than once were compared for all host-locality combinations. Jaccard’s index was used to assess similarity on the basis of presence/absence data only (see Arnold et al. 2000). The Morisita–Horn index was used to assess similarity on the basis of isolation frequency (see Gamboa & Bayman 2001; Arnold et al. 2003). Similarity indices were inferred using EstimateS v.7.5 (Colwell 2005: http://viceroy. eeb.uconn.edu/EstimateS).
M. T. Hoffman, A. E. Arnold
Manual alignment was performed using MacClade 4.08 (Maddison & Maddison 2005). Each alignment was based on LSU rDNA secondary structure as defined by Saccharomyces cerevisiae (Cannone et al. 2002). All ambiguously aligned regions were excluded. The alignment for the Dothideomycetes consisted of 4595 characters, of which 1208 were included; for the Sordariomycetes, 4605 characters, of which 1218 were included; for the Eurotiomycetes, 5443 characters, of which 2056 were included; for the Pezizomycetes, 4595 characters, of which 1208 were included. For each dataset, initial heuristic searches using parsimony as the optimality criterion were implemented in PAUP 4b10 (Swofford 2002), but these searches were ineffective in recovering optimal trees after up to five days. To improve the quality of searches, we initiated 30 sets of 200 searches for each dataset using PAUPRat (Sikes & Lewis 2001), which implements the ‘parsimony ratchet’ method (Nixon 1999) to efficiently recover shortest trees in datasets that are challenging for traditional heuristic searches (Sikes & Lewis 2001). The ‘filter trees’ command in PAUP 4b10 (Swofford 2002) was used to select all shortest trees from the resulting pools of trees (6001 trees per data set). The resulting set of shortest trees was then used to construct a strict consensus tree. Support was measured using a neighbor-joining bootstrap (1K replicates). To verify the quality of the topology given the PAUPRat approach, we conducted an additional set of analyses for each dataset using Bayesian MCMCMC. Based on comparisons of negative log likelihood values in Modeltest 3.7 (Posada & Crandall 1998), GTR þ I þ G was chosen as the best-fitting model for each dataset (Supplementary Material Appendix 4). Searches were implemented in MrBayes v. 3.1.1 (Huelsenbeck & Ronquist 2001) for 5M generations, initiated with random trees, four chains, and sampling every 500th tree. Extension of each analysis by up to 5M generations had no significant effect on negative log likelihood values. After elimination of the burn-in for each analysis, a majority rule consensus based on remaining trees was inferred.
Phylogenetic analyses
Results
Sixty-four LSU rDNA sequences for representative endophytes were organized at the class level on the basis of BLAST results and integrated into existing core alignments for the Dothideomycetes, Sordariomycetes, Eurotiomycetes, and Pezizomycetes. Core alignments included representative endophytes from boreal, temperate, and tropical sites (Arnold et al., in review; Arnold & Lutzoni 2007; Higgins et al. 2007; Supplementary Material Appendix 2) as well as named species of Ascomycota obtained from GenBank (Supplementary Material Appendix 3), and were subjected to preliminary analyses to ensure that the initial BLAST results were accurate (Higgins et al. 2007). Taxon sampling for the Dothideomycetes consisted of 26 named sequences and 78 endophytes (46 from this study); for the Sordariomycetes, 59 named sequences and 34 endophytes (15 from this study); for the Eurotiomycetes, 19 named sequences and three endophytes (one from this study); and the Pezizomycetes, 23 named sequences and four endophytes (two from this study).
From 960 tissue samples, 229 isolates of endophytic fungi were recovered in culture. Isolation frequency, defined as the percent of tissue segments bearing cultivable endophytes, was more than threefold greater for hosts from NC (56.3 %) than AZ (15.8 %), and was 2.5 to four times greater in Platycladus (AZ, 25.5 %; NC, 80.2 %) than in Cupressus (AZ; 6 %) or Juniperus (NC; 32.4 %; Table 1). Among 109 isolates sequenced for ITS rDNA, we recovered 35 unique ITS rDNA genotypes (90 % ITS rDNA sequence similarity; Fisher’s a ¼ 17.9). A total of 43 and 64 ITS rDNA genotypes based on 95 % and 99 % sequence similarity was recovered (Fisher’s a ¼ 26.2, and 65.1, respectively). Diversity of endophytes differed as a function of locality and host identity. Diversity of endophytes from hosts in NC was more than twofold greater than for hosts from AZ (Table 1). The total diversity of endophytes recovered from Juniperus and Cupressus was 1.4 times greater than from Platycladus (Table 1). In NC, diversity was ca twofold greater in
30
Endophytes of Cupressaceae
335
Table 1 – Number of tissue segments examined, isolates recovered, and isolates sequenced for ITS rDNA; diversity (Fisher’s alpha) of endophytes using functional taxonomic units based on 90, 95, and 99 % ITS rDNA sequence similarity; and number of singletons for each group based on 90 % ITS rDNA sequence similarity Host taxon/location
Segments plated
Isolates recovered
Isolates sequenced
Cupressus/AZ Juniperus/NC Platycladus/AZ Platycladus/NC
384 96 384 96
23 31 98 77
Cupressus & Juniperus Platycladus Cupressus & Platycladus (AZ) Juniperus & Platycladus (NC)
480 480 768 192
54 175 121 108
Fisher’s alpha
Singletons (90 % ITS rDNA)
90 % ITS rDNA
95 % ITS rDNA
99 % ITS rDNA
13 18 28 50
3.0 21.1 4.6 8.2
13.0 42.4 6.7 9.2
33.8 42.4 18.4 22.1
1 6 4 9
31 78 41 68
17.9 12.7 7.5 19.1
48.9 15.6 11.2 23.6
75.9 39.5 38.8 43.7
7 13 5 15
Hosts include Cupressus arizonica, Arizona (AZ); Juniperus virginiana, North Carolina (NC), and Platycladus orientalis in AZ and NC.
Juniperus than in Platycladus regardless of the stringency of ITS rDNA groups. In AZ, diversity in Cupressus was 1.6 times less than in Platycladus when species boundaries were based on 90 % ITS rDNA similarity. However, when species boundaries were defined at greater levels of ITS rDNA similarity (i.e. 95 and 99 % ITS rDNA congruence), diversity was ca twofold greater in Cupressus than in Platycladus (Table 1). Endophyte species richness also differed as a function of locality and host identity (Fig 1A–F). Greater species richness per sampling effort was recovered in NC versus AZ (Fig 1A– B), in Juniperus and Cupressus versus Platycladus (Fig 1C–D), and in Juniperus in NC relative to any other species/locality combination (Fig 1E–F). Overall, observed species richness fell within the 95 % confidence intervals for estimated richness, suggesting that our sample was effective in capturing the species richness of these endophyte communities (Fig 1G). However, the relatively steep slope of the accumulation curve indicates that numerous species of endophytes are yet to be recovered from these hosts as a whole. When genotype accumulation curves were compared for ITS rDNA groups based on different levels of sequence similarity, we found the largest differentiation between the 95 and 99 % groupings, with the most marked difference occurring at ca 98 % similarity (Fig 1H). This reflects that most of the endophytes recovered in this study are clustered in phylogenetically distinct groups that share high (98 %) ITS rDNA sequence similarity (see also Figs 2–5).
Relative importance of locality versus host identity Similarity indices based on both presence/absence data (Jaccard’s index) and isolation frequencies (Morisita–Horn index) and using 90 % ITS rDNA groupings suggest that endophyte communities in these hosts are structured more by locality than by host identity (Table 2). Highest similarity was observed among samples from different host species in the same site (i.e. Platycladus and Cupressus in AZ; Platycladus and Juniperus in NC). In contrast, samples from the same host species in different sites had low similarity values (i.e. Platycladus in NC and AZ). When compared across study sites, Juniperus and Cupressus did not share more endophytes with one another than with Platycladus.
Despite a large number of shared species between hosts within each site, the dominant endophyte taxon associated with each host and locality was distinct (Supplementary Material Appendix 1). In all cases, the most abundant taxa were Dothideomycetes. Cupressus (AZ) was dominated by an unidentified dothideomycete (90 % ITS rDNA genotype I). The highly diverse community of endophytes associated with Juniperus (NC) was not characterized by a dominant species, although three isolates (of 18 sequenced) represented a dothideomycete with phylogenetic affinity to Letendraea (90 % ITS rDNA genotype E; see below). Platycladus in NC was dominated by Phyllosticta sp. (90 % ITS rDNA genotype C; 21 of 68 isolates), and in AZ by Aureobasidium sp. and Phoma sp. (90 % ITS rDNA genotypes F and A, respectively, comprising ten and nine isolates of 41 sequenced).
Phylogenetic relationships of endophytes Results of phylogenetic analyses of endophytes from Cupressaceae, representative endophytes from other host species and sites, and representative Ascomycota for each class are shown in Figs 2–5. The majority of endophytes recovered in this study were placed with support in the Dothideomycetes (46 isolates) and Sordariomycetes (15 isolates). Only one endophyte (9147 from Platycladus in NC) was recovered as a member of the Eurotiomycetes. No endophytes were recovered among the Leotiomycetes, or from the lichen-dominated clades Lecanoromycetes, Lichinomycetes, and Arthoniomycetes. Two endophytes from Platycladus in AZ formed a clade within a well-supported lineage of Pezizomycetes. Phylogenetic analyses of the Dothideomycetes (Fig 2) using the parsimony ratchet yielded 780 best trees with tree lengths of 731 steps. BS support 70 % was observed for 12 nodes. Bayesian PP values 95 % were observed for 65 nodes, including all nodes with strong BS support. Many endophytes recovered in this study were reconstructed as part of larger clades of endophytic fungi from diverse hosts and sites that were strongly supported as sister to Botryosphaeria ribis (clade A), reconstructed in a clade containing Discosphaerina fagi and a variety of boreal forest endophytes (clade B), or strongly supported as part of a clade containing Phoma glomerata and a conifer endophyte from boreal forest (clades E and F). The
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Fig 1 – Genotype accumulation curves for fungal endophytes of Cupressus arizonica (AZ), Juniperus virginiana (NC), and Platycladus orientalis (AZ and NC). (A–G) Genotype groups are based on 90 % ITS rDNA sequence similarity, which is used here as a conservative proxy for species boundaries (following Arnold et al. 2007). (A) Species accumulation for endophytes of hosts in AZ. (B) Species accumulation for endophytes of hosts in NC. (C) Species accumulation of endophytes of native hosts: Cupressus (AZ) and Juniperus (NC). (D) Species accumulation of endophytes from non-native Platycladus (AZ and NC). (E) Comparison of species accumulation curves for each host species in AZ. (F) Comparison of species accumulation curves for each host species in NC. (G) Accumulation of ITS rDNA genotypes over the entire study. (H) Species accumulation and richness inferred using groups based on 90, 95, and 99 % ITS rDNA similarity.
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Fig 2 – Majority rule consensus tree based on Bayesian analyses of LSU rDNA data for the Dothideomycetes. Numbers above branches indicate branch support: Bayesian PP values 95 % are shown before the slash; NJ BS values 70 % are shown after the slash. Branches recovered in strict consensus tree from parsimony analyses are shown with bold black lines. Circles after endophyte culture numbers indicate native host species; triangles indicate non-native hosts. Letters indicate 90 % ITS rDNA genotype groups based on sequence similarity (Supplementary Material Appendix 1). Endophytic fungi from other studies are listed in Supplementary Material Appendix 2. All named taxa and GenBank identification numbers are shown in Supplementary Material Appendix 3. Dendrographa minor was included as an outgroup.
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Leotia lubrica Lulworthia fucicola Lulworthia grandispora Oxydothis frondicola Microdochium nivale Hyponectria buxi 4653 Picea mariana - Québec Arthrinium phaeospermum Apiospora sinensis 6722 Faramea occidentalis - Panama 9143 Platycladus orientalis - NC J 9089 Platycladus orientalis - AZ R 9126 Platycladus orientalis - NC J 9121 Platycladus orientalis - NC J Cryptosphaeria eunomia Fasciatispora petrakii Cainia graminis Seynesia erumpens Hypoxylon fragiforme 256 Laetia thamnia - Panama 2204 Pinus taeda - North Carolina 6717 Gustavia superba - Panama 9202 Juniperus virginiana - NC L 9201 Juniperus virginiana - NC L Astrocystis cocoes Rosellinia necatrix Xylaria acuta Xylaria hypoxylon 6728 Faramea occidentalis - Panama Halorosellinia oceanica Thyridium vestitum Gaeumannomyces graminis var graminis 6702 Trichilia tuberculata - Panama Diaporthe phaseolorum Valsa ambiens Cryptodiaporthe corni Cryphonectria cubensis Cryphonectria havanensis Melanconis marginalis Discula destructiva Gnomonia setacea Gnomoniella fraxini Plagiostoma euphorbiae Cephalotheca sulfurea Menispora tortuosa Camarops microspora Farrowia longicollea Farrowia seminuda Neurospora crassa Sordaria macrospora Sordaria fimicola Chaetomium globosum 9038 Platycladus orientalis - AZ P 9097 Cupressus arizonica - AZ T 9069 Platycladus orientalis - AZ H 9093 Platycladus orientalis - AZ H 9094 Platycladus orientalis - AZ H 9092 Platycladus orientalis - AZ S 9085 Platycladus orientalis - AZ H Pleurothecium recurvatum 6733 Swartzia simplex - Panama 412 Laetia thamnia - Panama 6756 Gustavia superba - Panama 6741 Theobroma cacao - Panama 6749 Theobroma cacao - Panama 6714 Gustavia superba - Panama 151 Laetia thamnia - Panama Microascus trigonosporus Ascosalsum cincinnatulum Magnisphaera stevemossago Ascosacculus heteroguttulatus Natantispora lotica Halosarpheia marina Sagaaromyces abonnis Corollospora maritima Varicosporina ramulosa Torrubiella luteorostrata Hydropisphaera erubescens Cordyceps khaoyaiensis 425 Laetia thamnia - Panama 4939 Picea mariana - Québec 416 Laetia thamnia - Panama 6708 Gustavia superba - Panama Balansia henningsiana Hypocrea citrina Verticillium epiphytum Emericellopsis terricola K 9154 Platycladus orientalis - NC K 9168 Platycladus orientalis - NC 4780 Dryas integrifolia - Iqaluit Hypomyces polyporinus Cordyceps irangiensis Cordyceps sinensis
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Fig 4 – Majority rule consensus tree based on Bayesian analyses of LSU rDNA data for the Eurotiomycetes. Numbers above branches indicate branch support: Bayesian PP values 95 % are shown before the slash; NJ BS values 70 % are shown after the slash. Branches recovered in strict consensus tree from parsimony analyses are shown with bold black lines. Triangle indicates non-native host. Letters indicate 90 % ITS rDNA genotype groups based on sequence similarity (Supplementary Material Appendix 1). Endophytic fungi from other studies are listed in Supplementary Material Appendix 2. All named taxa and GenBank identification numbers are shown in Supplementary Material Appendix 3. Peltula umbilicata was included as an outgroup.
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Fig 5 – Majority rule consensus tree based on Bayesian analyses of LSU rDNA data for the Pezizomycetes. Numbers above branches indicate branch support: Bayesian PP values 95 % are shown before the slash; NJ BS values 70 % are shown after the slash. Branches recovered in strict consensus tree from parsimony analyses are shown with bold black lines. Triangles indicate non-native hosts. Letters indicate 90 % ITS rDNA genotype groups based on sequence similarity (Supplementary Material Appendix 1). Endophytic fungi from other studies are listed in Supplementary Material Appendix 2. All named taxa and GenBank identification numbers are shown in Supplementary Material Appendix 3. The five taxa at the base of the tree were included as outgroups.
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Table 2 – Similarity of endophyte assemblages with regard to locality and host identity based on nonsingleton ITS rDNA genotypes (90 % sequence similarity) Platycladus Platycladus Juniperus Cupressus NC AZ NC AZ Platycladus NC Platycladus AZ Juniperus NC Cupressus AZ
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majority of endophytes within the Dothideomycetes were reconstructed with strong support as members of the wellsupported Pleosporales (clades C–G). One additional endophyte (H) was reconstructed with strong support as the sister taxon of Raciborskiomyces longisetosum. Clade C was reconstructed as sister to Preussia terricola, albeit without strong support. Clade D was reconstructed as sister to Letendraea helminthicola. Clades E and F were strongly supported as a clade with Phoma glomerata and included an endophyte from Picea mariana in Que´bec. Clade G was well supported as sister to Pyrenophora tritici-repentis. Throughout the Dothideomycetes, ITS rDNA genotype groups (based on 90 % ITS rDNA similarity) were largely congruent with LSU rDNA phylotypes. Phylotypes containing endophytes from cupressaceous hosts often reflected locality (AZ versus NC; e.g. clades A, B), but within-clade structure based on locality and host identity was suggested in several clades (e.g. clades D–G). Analyses of the Sordariomycetes (Fig 3) using the parsimony ratchet yielded 626 best trees with tree lengths of 1289 steps. BS support 70 % was observed for 25 nodes, and Bayesian PP 95 % was observed for 53 nodes. Twenty-four nodes were supported by both methods. Endophytes from cupressaceous hosts were reconstructed within the well-supported Xylariales (clades A–B), the Sordariales (clade C), and the Hypocreales (clade D). Clades A and B were reconstructed with support as sister to endophytes from Panama. Clade C was reconstructed as sister to the ubiquitous fungus Chaetomium globosum, and clade D as sister to Emericellopsis, albeit without strong support. ITS rDNA genotype groups were consistent with LSU rDNA phylotypes for the hypocrealean endophytes (clade D) and xylarialean endophytes in clade B. However, clades A (Xylariales) and C (Sordariales) were characterized by poor matches between ITS rDNA genotype groups and LSU rDNA phylotypes. Phylotypes reflected clustering of endophytes from different host taxa (clade C) and sites (clade A) although two well-supported phylotypes (clades B, D) contained endophytes from a single host species in a single site. For the Eurotiomycetes (Fig 4), the parsimony ratchet approach yielded 1005 best trees with tree lengths of 387 steps. BS support 70 % was observed for seven nodes, and Bayesian PP 95 % was observed for 13 nodes. Six nodes were supported
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by both measures. Endophyte 9147 was reconstructed with strong support as sister to Penicillium freii within the wellsupported Eurotiales. For the Pezizomycetes (Fig 5), the parsimony ratchet approach yielded 827 best trees with tree lengths of 561 steps. BS support 70 % was observed for 11 nodes; Bayesian PP 95 % was observed for 16 nodes. Ten nodes received significant support from both methods. Endophytes 9098 and 9084a from Platycladus in AZ were reconstructed as sister to one another with strong support, and are nested within a strongly supported clade within the Pezizales.
Discussion We used a culture-based approach, paired with sequencing of both a fast-evolving locus (ITS rDNA) and a phylogenetically informative locus (LSU rDNA), to assess the diversity and taxon composition of fungal endophytes among native and non-native conifers in two geographically distinct localities. We found that the abundance, diversity, species composition, and relative host affinity of endophyte communities differed as a function of locality and host species. Cultivable endophytes were present in up to 80.2 % of tissue segments examined, but their incidence differed more than 13-fold as a function of locality and host identity. Despite very conservative estimates of species boundaries, diversity ranged from low values (Fisher’s a ¼ 3: Cupressus, AZ) to values consistent with those of angiosperm trees in tropical forests (Fisher’s a ¼ 21.1: Juniperus, NC; Arnold & Lutzoni 2007). Most endophytes were placed in phylogenetic analyses within the Dothideomycetes and, to a lesser extent, the Sordariomycetes, but members of both the Eurotiomycetes and Pezizomycetes were also recovered. Isolates representing the Dothideomycetes were proportionally more common in AZ than NC (ca 88 versus ca 75 % of isolates with definitively identified ITS rDNA genotypes, respectively; Supplementary Material Appendix 1, Figs 2–5). The Sordariomycetes were proportionally more common in NC than AZ (ca 21 versus ca 10 %, respectively). Overall and for three host/locality combinations (Cupressus AZ, Platycladus AZ, and Platycladus NC), the majority of definitively identified isolates were placed in the Dothideomycetes. However, Juniperus from NC was characterized by a higher incidence of Sordariomycetes (ca 56 % of identified isolates; Supplementary Material Appendix 1). Although endophyte genotypes often occurred in more than one host species (Supplementary Material Appendix 1) and clades often contained representatives of multiple hosts and sites (Figs 2–5), the occurrence of numerous singleton species (Table 1, Fig 1) and relatively low similarity among hosts and localities (Table 2) suggest that each tree species in each locality has a signature community of endophytic symbionts. This was underscored by the observation that the dominant endophyte species associated with each host was distinct (Supplementary Material Appendix 1). The conservative species concept applied here likely underestimates richness and diversity (Table 1), and may overestimate similarity of communities among hosts and sites (Table 2). Diversity of endophytes in these temperate hosts thus appears to be surprisingly high, with a high degree of geographic turnover
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and distinctively structured communities in each host and locality. Fisher et al. (1993, 1994) found that endophyte assemblages of trees planted outside their native range are depauperate in terms of infection frequency, species richness, and prevalence of host-specific endophytes. Although our study did not examine endophytes of Platycladus within its native range, we did compare endophytes of Platycladus with those of two closely related, native hosts in a mesic forest and an arid desert. In both sites, isolation frequency was higher in Platycladus than in either native host. Moreover, Platycladus in NC harboured cultivable endophytes in a significantly higher percentage of tissue segments (80.2 %) than did seven representative, native species studied previously in Duke Forest, NC (mean S.D. ¼ 36.4 % 27 of tissue segments; Wilcoxon signed-rank test, P ¼ 0.031; Arnold & Lutzoni 2007). In contrast, Juniperus fell within the range of other native taxa in that forest (32.3 %; Wilcoxon signed-rank test, P ¼ 0.578). Elevated frequencies of endophyte infection in Platycladus relative to closely related, co-occurring plants could reflect three non-exclusive factors: (1) the presence of both local and introduced endophytes within these non-native trees; (2) host-specific rates of infection that are independent of the native/non-native status of these plants; and/or (3) the presence of more cosmopolitan/less host-specific endophytes, which might be more readily cultivated than those with greater host specificity. Further sampling is needed to address the first two scenarios, such that firm conclusions on this topic cannot be drawn from this study. However, support for the third scenario comes from previous studies indicating that opportunistic, hostgeneralist endophytes are especially common in plants outside their native ranges (e.g. Fisher et al. 1994). Under this scenario, endophytes of Platycladus should be characterized by a greater proportion of host-generalists than Juniperus and Cupressus. Even though all species harboured putatively opportunistic, widespread taxa (e.g. Phoma sp., Aureobasidium sp., Alternaria sp., Cladosporium sp., Xylaria sp.), these cosmopolitan genera were nearly five times more common in Platycladus than either native host (Supplementary Material Appendix 1). We also found that these cosmopolitan genera were three times more common in AZ than in NC. We hypothesize that desert endophytes may be under particularly strong selection to act as host-generalists: the benefit of opportunistically infecting hosts is likely high relative to the cost of prolonged exposure to intense heat, UV radiation, and desiccation typical of this region. High rates of host generalism among desert endophytes are further supported by similarity indices, which showed greater similarity in endophyte communities for different host species in AZ than in NC (Table 2). The most common endophytes recovered from our AZ site showed affinity to Aureobasidium sp., Phoma sp., and Alternaria sp. (Supplementary Material Appendix 1), consistent with a recent survey of non-host-specific cactus endophytes in AZ (Suryanarayanan et al. 2005). Thus our data suggest that relative host affinity of endophyte communities has the potential to change over the geographic range of host plant lineages, and may differ among host plant taxa as well. Arnold et al. (2007) investigated the use of a 600 bp region of LSU rDNA, paired with a phylogenetic backbone constraint, as
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a search strategy for parsimony analyses involving environmental samples (e.g. unknown endophyte cultures). That study concluded that additional data were needed to more adequately infer phylogenetic relationships for large datasets. In this study, we found that ca 1200 bp of LSU rDNA, coupled with the PAUPRat method, provided the tools for expediently recovering topologies within four classes of Ascomycota. These topologies were not in conflict with topologies resulting from Bayesian analysis (Figs 2–5). However, Bayesian methods consistently yielded greater numbers of significantly supported nodes than did the BS method employed here, which relied on NJ to rapidly assess branch support. Well-supported phylogenies are key to diagnosing the taxonomic affinity of unknown endophytes and are necessary for adequately addressing ecological questions for these sterile isolates. Future studies will benefit from multiple, concurrent analysis methods, especially with regard to linking traditional morphological species concepts to extensive molecular datasets. The occurrence of clades composed only of endophytes limits our ability to diagnose species boundaries using the phylogenetic criteria proposed in other studies (e.g. Arnold et al. 2007). However, well-supported, endophyte-containing clades provide a tool for assessing the frequency with which particular fungal lineages, rather than genotypes, are represented among different host species or sites. The prevalence of endophyte-only clades suggests that cupressaceous hosts harbour interesting and perhaps novel taxa of Ascomycota, but such statements can only be made with caution pending the population of public databases with additional sequences for known but unsequenced fungi, and ongoing morphological delimitation of the isolates recovered here. Such clades also are consistent with the diversification of fungi through the endophyte symbiosis. Intensive sampling of multiple host species within a given plant family represents an important but under-explored approach for investigating potential co-cladogenesis of endophytes and their plant hosts. Although the designation of ITS rDNA genotype groups based on percent similarity is a straightforward and rapid method for designating species boundaries, there is no threshold value of sequence similarity that is universally useful for distinguishing species of fungi. Percent divergence between species is likely to range widely among and within major lineages, different biological species of fungi frequently have identical ITS rDNA sequences (Lieckfeldt & Seifert 2000), and cryptic species may sometimes only be revealed through a phylogenetic approach based on other loci (Taylor et al. 2000). We found that our conclusions regarding the relative diversity of endophytes associated with Cupressus versus Platycladus were reversed when different ITS rDNA genotype similarities were used to designate species boundaries (Table 1). This observation, coupled with clade-specific differences in concordance between ITS rDNA genotypes and phylotypes (Figs 2–5), warrants continued conservatism in using sequence similarity as a proxy for species boundaries. With the exception of native fescue grass in AZ, the community structure, ecological roles, and evolution of fungal endophytes have not been well defined in desert environments, particularly with molecular techniques (Faeth & Sullivan 2003; but also see Suryanarayanan et al. 2005). How such
Endophytes of Cupressaceae
endophytes might benefit or negatively influence their host plant, particularly in times of severe stress, has not been determined. Similarly, the biotically rich forests of the mesic southeastern USA remain mostly unexplored in terms of their endophytic symbionts, and may harbour a diversity of endophytes similar in scale to that in tropical forests (Arnold & Lutzoni 2007). The relative costs and benefits of endophytic symbioses for particular plant species remain largely unresolved, and are particularly interesting in an evolutionary context given the ways in which endophyte communities differ in abundance, diversity, composition, and relative host specificity over the ranges of their host plants.
Acknowledgements We thank the University of Arizona’s Division of Plant Pathology and Microbiology and College of Agriculture and Life Sciences for supporting this work. Additional support from the National Science Foundation [DEB-0343953 to A.E.A. (and James W. Dalling); DEB-0200413 to A.E.A.] is gratefully acknowledged. We thank David Maddison for sharing pre-release versions of MacClade and Mesquite, and for advice with regard to phylogenetic analyses. We are grateful to Rebecca Porter, Mary Shimabukuro, Eric Janson, and Lindsay Higgins for technical assistance, and to Cara Gibson, Mali Gunatilaka, Ming-Min Lee, and especially Franc¸ois Lutzoni for helpful discussion.
Supplementary data Supplementary data associated with this article can be found in the online version, at doi:10.1016/j.mycres.2007.10.014.
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40
Appendix 1. Endophyte isolates (identification numbers, host taxa, and origin), ITS rDNA genotype groups based on 90%, 95%, and 99% sequence similarity, and top BLAST matches based on analyses conducted in AugustOctober, 2005. Isolates used for phylogenetic analyses of LSU rDNA data are marked with asterisks. Four isolates were included in LSUrDNA analyses but are not listed below because ITS rDNA sequences were not obtained (9079, 9084b, 9096, and 9098). Identification numbers correspond to vouchers maintained as living cultures at the University of Arizona.
Isolate
Host
Location
ITS rDNA ITS rDNA ITS rDNA genotype genotype genotype Blast top match (ITS):GenBank accession number, ID, and e-value (90%) (95%) (99%)
*9002 Platycladus
AZ
F
G
3L
gi|30026473|gb|AY225167.1| Aureobasidium pullulans isolate LB... 1053
*9005 Cupressus
AZ
A
2Q
R
gi|28974197|gb|AY183371.1| Phoma glomerata 18S ribosomal RNA ... 993
0.0
*9007 Platycladus
AZ
A
A
J
gi|59783265|gb|AY831558.1| Leptosphaerulina trifolii strain W... 805
*9008 Cupressus
AZ
A
A
A
gi|33302361|gb|AY337712.1| Phoma herbarum 18S ribosomal RNA g... 862
*9009a Platycladus
AZ
B
B
B
gi|32394890|gb|AY154712.1| Alternaria tenuissima strain IA287... 1063
*9009b Platycladus
AZ
F
G
3L
gi|30026473|gb|AY225167.1| Aureobasidium pullulans isolate LB... 1047
*9015 Platycladus
AZ
F
G
S
gi|37786120|gb|AY213639.1| Aureobasidium pullulans strain UWF... 991
*9018 Platycladus
AZ
A
A
A
gi|33302361|gb|AY337712.1| Phoma herbarum 18S ribosomal RNA g... 860
*9021 Platycladus
AZ
B
B
B
gi|32394890|gb|AY154712.1| Alternaria tenuissima strain IA287... 1082
0.0
*9026 Platycladus
AZ
B
B
B
gi|32394890|gb|AY154712.1| Alternaria tenuissima strain IA287... 1112
0.0
*9027 Platycladus
AZ
A
A
J
gi|59783265|gb|AY831558.1| Leptosphaerulina trifolii strain W... 837
*9028 Platycladus
AZ
F
G
K
gi|30026472|gb|AY225166.1| Aureobasidium pullulans isolate SK... 993
*9030 Platycladus
AZ
A
A
J
gi|59783265|gb|AY831558.1| Leptosphaerulina trifolii strain W... 839
*9031 Platycladus
AZ
F
G
K
gi|30026472|gb|AY225166.1| Aureobasidium pullulans isolate SK... 1017
0.0
*9036 Platycladus
AZ
F
G
K
gi|30026472|gb|AY225166.1| Aureobasidium pullulans isolate SK... 918
0.0
0.0
0.0 0.0
0.0 0.0 0.0
0.0 0.0
0.0
0.0
41
*9038 Platycladus
AZ
P
P
T
gi|30089120|emb|AJ458185.1|CNI458185 Chaetomium nigricolor 5.... 714
*9042 Platycladus
AZ
F
G
K
gi|30026472|gb|AY225166.1| Aureobasidium pullulans isolate SK... 993
0.0
*9051 Platycladus
AZ
M
M
U
gi|41058714|gb|AY465446.1| Dothideales sp. GS2N1c 18S ribosom... 646
*9054 Platycladus
AZ
A
2Q
M
gi|28974197|gb|AY183371.1| Phoma glomerata 18S ribosomal RNA ... 950
*9055 Platycladus
AZ
B
B
B
gi|32394890|gb|AY154712.1| Alternaria tenuissima strain IA287... 1114
*9057 Cupressus
AZ
I
H
L
gi|47420271|gb|AY546017.1| Fungal endophyte WMS23 internal tr... 813
0.0
*9058 Cupressus
AZ
I
H
L
gi|47420271|gb|AY546017.1| Fungal endophyte WMS23 internal tr... 813
0.0
*9059 Cupressus
AZ
I
H
L
gi|47420271|gb|AY546017.1| Fungal endophyte WMS23 internal tr... 813
*9060 Platycladus
AZ
F
G
V
gi|30026472|gb|AY225166.1| Aureobasidium pullulans isolate SK... 995
9064 Platycladus
AZ
A
2Q
3K
gi|28974197|gb|AY183371.1| Phoma glomerata 18S ribosomal RNA ... 997
*9065 Platycladus
AZ
A
2Q
M
gi|28974197|gb|AY183371.1| Phoma glomerata 18S ribosomal RNA ... 993
*9069 Platycladus
AZ
H
F
H
gi|29165262|gb|AY219880.1| Lecythophora sp. UBCtra1453C 18S r... 868
*9084a Platycladus
AZ
Q
Q
W
gi|25045988|gb|AF491556.1| Peziza varia KH-97-88 (C) 18S ribo... 1183
0.0
*9085 Platycladus
AZ
H
R
X
gi|28202120|gb|AY141180.1| Aureobasidium pullulans 18S riboso... 821
0.0
*9089 Platycladus
AZ
R
S
Y
gi|22773864|gb|AF405301.1| Bartalinia robillardoides 18S ribo... 854
*9092 Platycladus
AZ
S
T
Z
gi|71725083|dbj|AB231012.1| Lecythophora hoffmannii genes for... 880
*9093 Platycladus
AZ
H
F
H
gi|29165262|gb|AY219880.1| Lecythophora sp. UBCtra1453C 18S r... 868
0.0
*9094 Platycladus
AZ
H
F
2A
gi|29165262|gb|AY219880.1| Lecythophora sp. UBCtra1453C 18S r... 872
0.0
*9097 Cupressus
AZ
T
U
2B
gi|12583572|emb|AJ271575.1|TSU271575 Thielavia subthermophila... 1053
*9104 Cupressus
AZ
M
N
2C
gi|46243857|gb|AY510419.1| Preussia similis strain s19 intern... 942
*9106 Cupressus
AZ
M
N
2D
gi|41058713|gb|AY465445.1| Dothideales sp. GS5N1b 18S ribosom... 890
*9107 Cupressus
AZ
A
2Q
3K
gi|28974197|gb|AY183371.1| Phoma glomerata 18S ribosomal RNA ... 971
9115 Cupressus
AZ
I
V
2E
gi|47420271|gb|AY546017.1| Fungal endophyte WMS23 internal tr... 722
*9116 Cupressus
AZ
M
W
2F
emb|AJ972795.1| Monodictys sp. MA 4647 18S rRNA gene, 5.8S rR... 751
*9120 Cupressus
AZ
M
M
2G
gb|AY465446.1| Dothideales sp. GS2N1c 18S ribosomal RNA gene,... 607
*9121 Platycladus
NC
J
J
2H
gb|AF377289.1| Pestalotiopsis funereoides 18S ribosomal RNA g... 1035
0.0
0.0 0.0 0.0
0.0
0.0 0.0 0.0 0.0 0.0
0.0 0.0
0.0
0.0 0.0 0.0 0.0 0.0 1e-170
9122 Platycladus
NC
J
J
O
gb|AF377289.1| Pestalotiopsis funereoides 18S ribosomal RNA g... 1039
0.0
*9126 Platycladus
NC
J
J
O
gb|AF377289.1| Pestalotiopsis funereoides 18S ribosomal RNA g... 1037
0.0
*9128 Platycladus
NC
U
X
2I
gb|AF393724.2| Cladosporium tenuissimum strain ATCC 38027 18S... 961
0.0
*9129 Platycladus
NC
C
C
D
0.0
*9130 Platycladus
NC
C
C
D
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1154 gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1160 0.0
9133 Platycladus
NC
D
D
E
gb|AY786322.1| Botryosphaeria dothidea strain CBS 116743 18S ... 1041
0.0
42
*9134 Platycladus
NC
C
C
D
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1158
0.0
*9135 Platycladus
NC
C
C
D
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1154
0.0 0.0
9136 Platycladus
NC
C
C
D
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1116
*9137 Platycladus
NC
E
E
F
gb|AY561200.1| Foliar endophyte of Picea glauca sp. P3 intern... 1063
0.0
9138 Platycladus
NC
V
Y
2J
gb|DQ094165.1| Fungal sp. YX-41 internal transcribed spacer 1... 999
0.0
*9140 Platycladus
NC
E
E
F
gb|AY561200.1| Foliar endophyte of Picea glauca sp. P3 intern... 1072
0.0
9142 Platycladus
NC
C
C
C
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1154
*9143 Platycladus
NC
J
J
2K
gb|AF377289.1| Pestalotiopsis funereoides 18S ribosomal RNA g... 1037
*9144 Platycladus
NC
C
C
D
gi|12082793|gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ... 1217
*9145 Platycladus
NC
E
E
F
gb|AY561200.1| Foliar endophyte of Picea glauca sp. P3 intern... 1076
0.0 0.0 0.0
0.0
9146 Platycladus
NC
E
E
I
gb|DQ094168.1| Fungal sp. YX-47 internal transcribed spacer 1... 1041
0.0
*9147 Platycladus
NC
W
Z
2L
gb|AF125942.1| Penicillium sp. NRRL 28148 internal transcribe... 1033
0.0
9148 Platycladus
NC
E
E
I
gb|DQ094168.1| Fungal sp. YX-47 internal transcribed spacer 1... 1017
0.0
*9149a Platycladus
NC
E
E
F
gb|AY561200.1| Foliar endophyte of Picea glauca sp. P3 intern... 1055
0.0
*9149b Platycladus
NC
C
I
N
gi|12082793|gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ... 908
9150 Platycladus
NC
C
C
G
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1122
9151 Platycladus
NC
C
C
G
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1120
9153 Platycladus
NC
X
2A
2M
gb|AY063310.1| Fungal endophyte WMS14 internal transcribed sp... 648
*9154 Platycladus
NC
K
K
2N
gb|AY833034.1| Uncultured mycorrhizal ascomycete isolate 1 in... 387
9157 Platycladus
NC
A
A
2O
gb|AY337712.1| Phoma herbarum 18S ribosomal RNA gene, partial... 825
*9158 Platycladus
NC
A
A
J
gb|AY131201.1| Ascochyta lentis isolate MU AL1 18S ribosomal ... 811
9160 Platycladus
NC
C
C
D
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1164
0.0
9162 Platycladus
NC
C
C
G
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1120
0.0
9163 Platycladus
NC
C
I
2P
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 771
0.0
9165 Platycladus
NC
Y
2B
2Q
gb|AY265334.1| Scolecobasidium humicola strain B3343 internal... 315
8e-83
9167 Platycladus
NC
E
E
I
gb|DQ094168.1| Fungal sp. YX-47 internal transcribed spacer 1... 1033
0.0
*9168 Platycladus
NC
K
K
2R
gb|AY833034.1| Uncultured mycorrhizal ascomycete isolate 1 in... 387
2e-104
9169 Platycladus
NC
K
K
2S
dbj|AB067720.1| Cordyceps sinensis genes for 18S rRNA, ITS1, ... 464
1e-127
9171 Platycladus
NC
Z
2C
2T
emb|AJ390436.1|HAN390436 Nemania serpens 18S rRNA gene (parti... 969
9172 Platycladus
NC
C
C
D
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1158
9173 Platycladus
NC
C
C
D
gi|12082793|gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ... 1078
9174 Cupressus
AZ
F
G
2U
gb|AY225166.1| Aureobasidium pullulans isolate SK3 18S riboso... 934
0.0 0.0 0.0 0.0
2e-104 0.0
0.0
0.0 0.0
0.
0.0
43
9176 Platycladus
NC
C
C
C
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1156
9177 Platycladus
NC
C
C
C
gi|12082793|gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ... 1245
0.0
0.0
9178 Platycladus
NC
C
I
N
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 908
0.0
9179 Platycladus
NC
G
O
Q
gb|AF502622.1| Leaf litter ascomycete strain its031 isolate 1... 820
9183 Juniperus
NC
A
2Q
2V
gi|28974197|gb|AY183371.1| Phoma glomerata 18S ribosomal RNA ... 1049
9184 Platycladus
NC
C
C
G
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1076
0.0
9187 Juniperus
NC
O
2D
2W
dbj|AB107890.1| Phomopsis sp. MAFF 665006 genes for ITS1, 5.8... 1021
0.0
9188 Juniperus
NC
O
2E
2X
gb|AY577815.1| Diaporthe phaseolorum strain E99382 18S riboso... 892
9189 Juniperus
NC
2A
2F
2Y
gb|AY208791.1| Stagonospora sp. Po41 internal transcribed spa... 839
9190 Juniperus
NC
D
D
E
gb|AY786322.1| Botryosphaeria dothidea strain CBS 116743 18S ... 987
9191 Platycladus
NC
2B
2G
2Z
gb|AY213667.1| Paecilomyces lilacinus strain UWFP 674 18S rib... 969
9193 Juniperus
NC
2C
2H
3A
gb|AY063297.1| Fungal endophyte WMS1 internal transcribed spa... 593
9194 Platycladus
NC
G
O
Q
gb|AF502622.1| Leaf litter ascomycete strain its031 isolate 1... 813
9195 Platycladus
NC
C
C
D
gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ribosomal RN... 1154
9198 Juniperus
NC
2A
2I
3B
gb|AY183366.1| Rhizosphaera kalkhoffii 18S ribosomal RNA gene... 829
emb|AJ539482.1|MCR539482 Morchella crassipes 18S rRNA gene (p... 646
0.0 0.0
0.0 0.0 0.0 0.0 2e-166
0.0 0.0 0.0
9199 Juniperus
NC
2D
2J
3C
*9200 Juniperus
NC
E
E
F
gb|AY561200.1| Foliar endophyte of Picea glauca sp. P3 intern... 1067
0.0
*9201 Juniperus
NC
L
L
P
emb|AJ390411.1|HAN390411 Biscogniauxia atropunctata 18S rRNA ... 1057
0.0
*9202 Juniperus
NC
L
L
P
emb|AJ390411.1|HAN390411 Biscogniauxia atropunctata 18S rRNA ... 1017
0.0
*9203 Juniperus
NC
E
E
F
gb|AY561200.1| Foliar endophyte of Picea glauca sp. P3 intern... 1061
9206 Platycladus
NC
2E
2K
3D
gb|AY971711.1| Fungal sp. 4.32 18S ribosomal RNA gene, partia... 607
9208 Juniperus
NC
2F
2L
3E
gb|AY315399.1| Xylariaceae sp. C8 internal transcribed spacer... 410
9209 Juniperus
NC
2G
2M
3F
gb|AF409982.1| Pestalotiopsis sp. EN 7 18S ribosomal RNA gene... 442
9211 Juniperus
NC
2H
2N
3G
gb|AY315396.1| Xylaria sp. F23 internal transcribed spacer 1,... 1322
9217 Juniperus
NC
E
E
F
9220 Juniperus
NC
J
J
3H
gb|AY687871.1| Pestalotia lawsoniae voucher PSH2000I-057 18S ... 1070
0.0
9247 Juniperus
NC
N
2O
3I
gb|AF260224.1|AF260224 Kabatina juniperi 18S ribosomal RNA, p... 1155
0.0
9248 Platycladus
NC
G
O
Q
gb|AF502622.1| Leaf litter ascomycete strain its031 isolate 1... 816
9249 Platycladus
NC
C
C
C
gi|12082793|gb|AF312009.1|AF312009 Phyllosticta spinarum 18S ... 1245
9283 Platycladus
NC
2I
2P
3J
dbj|AB041241.1| Guignardia mangiferae genes for 18S rRNA, ITS... 1274
0.0
0.0 1e-170 1e-111 4e-121
0.0
gb|AY561200.1| Foliar endophyte of Picea glauca sp. P3 intern... 1037
0.0
0.0 0.0 0.0
44
Appendix 2. GenBank accession numbers for LSU rDNA sequences of endophytic fungi obtained from other studies for phylogenetic analyses.
Endophyte isolate
Accession no.
4245 4250 4466 422 4653 6722 256 2204 6717 6728 6702 6733 412 6756 6741 6749 6714 151 425 4939 416 6708 4780 469 2834 2779 3300 1182 6730 6731 435b 2712 2722 462 4140 3358A 4221 3326 5007 1029
DQ979442 DQ979443 DQ979444 EF420091 DQ979449 EF420100 EF420088 EF420087 EF420099 EF420101 EF420096 EF420104 EF420089 EF420107 EF420105 EF420106 EF420098 EF420086 EF420092 DQ979453 EF420090 EF420097 DQ979452 EF420095 DQ979423 DQ979422 DQ979427 EF420085 EF420102 EF420103 EF420093 DQ979419 DQ979420 EF420094 DQ979440 DQ979433 DQ979441 DQ979429 DQ979455 EF420084
45
4109 4947A 4089 3275 3335 3353 3329 3377 5718 5095 3323 5654 2851 2706 5246
DQ979438 DQ979454 DQ979437 DQ979425 DQ979431 DQ979432 DQ979430 DQ979434 DQ979463 DQ979457 DQ979428 DQ979462 DQ979424 DQ979418 DQ979458
46
Appendix 3. Representative Ascomycota species included in phylogenetic analyses and GenBank identification numbers. Sequences for Coccidioides posadasii were obtained from TIGR (http://www.tigr.org).
Taxon
GenBank ID
Aleuria aurantia Ampelomyces quisqualis Aphanoascus fulvescens Apiospora sinensis Arthrinium phaeospermum Arthroderma curreyi Ascobolus carbonarius Ascobolus crenulatus Ascosacculus heteroguttulatus Ascosalsum cincinnatulum Ascosphaera apis Astrocystis cocoes Auxarthron zuffianum Balansia henningsiana Barssia oregonensis Bimuria novae-zelandiae Botryosphaeria ribis Byssochlamys nivea Byssothecium circinans Cainia graminis Camarops microspora Candida albicans Capronia mansonii Capronia pilosella Cephalotheca sulfurea Ceramothyrium carniolicum Chaetomium globosum Cheilymenia stercorea Coccidioides posadasii Cochliobolus heterostrophus Cordyceps irangiensis Cordyceps khaoyaiensis Cordyceps sinensis Corollospora maritima Cryphonectria cubensis Cryphonectria havanensis Cryptodiaporthe corni Cryptosphaeria eunomia var. eunomia Curvularia brachyspora Delphinella strobiligena Dendrographa leucophaea f. minor Dermatocarpon luridum Diaporthe phaseolorum Disciotis venosa Discosphaerina fagi Discula destructiva Dothidea insculpta Dothidea ribesia
45775583 31415592 33113349 27447246 27447247 33113367 45775606 45775607 34453082 34453080 11120650 27447238 33113353 45155172 45775581 15808399 11120642 33113391 15808400 20334356 27447236 671812 11120644 12025061 20334357 11120645 13469777 45775590 TIGR222929 45775574 13241779 13241765 29466997 22203655 22671402 22671403 22671407 27447241 12025063 15808401 47499217 52699696 1399144 45775596 15808402 22671422 52699697 15808403
47
Dothidea sambuci Emericellopsis terricola Eremascus albus Exophiala jeanselmei Fasciatispora petrakii Farrowia longicollea Farrowia seminuda Gaeumannomyces graminis var. graminis Glyphium elatum Gnomonia setacea Gnomoniella fraxini Gyromitra esculenta Gyromitra californica Halorosellinia oceanica Halosarpheia marina Helvella compressa Hydropisphaera erubescens Hypocrea citrina Hypomyces polyporinus Hyponectria buxi Hypoxylon fragiforme Leotia lubrica Letendraea helminthicola Lojkania enalia Lulworthia fucicola Lulworthia grandispora Magnisphaera stevemossago Melanconis marginalis Menispora tortuosa Microascus trigonosporus Microdochium nivale Microxyphium citri Morchella elata Morchella esculenta Natantispora lotica Neolecta vitellina Neurospora crassa Otidea onotica Oxydothis frondicola Paracoccidioides brasiliensis Peltula umbilicata Penicillium freii Peziza proteana Peziza quelepidotia Peziza succosa Phaeosphaeria avenaria Phoma glomerata Plagiostoma euphorbiae Pleurothecium recurvatum Preussia terricola Pyrenophora tritici-repentis Pyrenula cruenta Pyrenula pseudobufonia Raciborskiomyces longisetosum Rosellinia necatrix Saccharomyces cerevisiae Sagaaromyces abonnis Sarcoscypha coccinea Sarcosphaera crassa Schizosaccharomyces pombe Setomelanomma holmii Setosphaeria monoceras
45775610 1698507 11120651 4206347 27447243 13469782 13469784 16565889 17104831 16565895 16565884 52699700 45775602 27447237 34453085 45775584 45155165 45775578 32261014 27447249 27447244 45775573 15808405 15808406 22203665 22203666 34453095 22671436 45775611 1399149 2565289 11120643 45775594 12025081 34453084 12025084 13469785 17154899 27447250 2271524 15216700 52699706 45775588 52699707 17154944 45775613 31415593 22671445 45775614 45775615 45775601 12025090 52699710 15808410 27447239 172409 34453078 45775576 45775597 288696 28144315 15808411
48
Seynesia erumpens Sordaria fimicola Sordaria macrospora Stylodothis puccinioides Sydowia polyspora Taphrina communis Thyridium vestitum Torrubiella luteorostrata Trematosphaeria heterospora Varicosporina ramulosa Verpa conica Verrucaria pachyderma Verticillium epiphytum Valsa ambiens Westerdykella cylindrica Xylaria acuta Xylaria hypoxylon
12025093 45155203 37992293 11120648 45775604 52699720 45775600 13241778 15808412 22203671 45775595 15216679 8777749 16565896 11120649 45775605 45775577
Appendix 4. Model parameters for Bayesian analyses for each focal class of Ascomycota. Class
Selected model
- Ln likelihood
Dothideomycetes Sordariomycetes Eurotiomycetes Pezizomycetes
GTR+I+G GTR+I+G GTR+I+G GTR+I+G
8463.17251 5924.5000 3613.9267 4519.40130
Proportion of invariant sites 0.420231 0.565827 0.634904 0.522561
Gamma shape parameter 0.526946 0.4267 0.55459 0.639625
Estimated Dothideomycetes base frequencies = A:0.266879 C:0.217424 G:0.307280 T:0.208417 Estimated Pezizomycetes base frequencies = A:0.287876 C:0.185319 G:0.288132 T:0.238673 Estimated Eurotiomecetes base frequencies = A:0.273327 C:0.208930 G:0.305662 T:0.212081 Estimated Sordariomycetes base frequencies = A:0.276040 C:0.210782 G:0.300050 T:0.213129
49
APPENDIX B
DIVERSE BACTERIAL ENDOSYMBIONTS INHABIT LIVING HYPHAE OF PHYLOGENETICALLY DIVERSE FUNGAL ENDOPHYTES
50
51
Diverse bacteria inhabit living hyphae of phylogenetically diverse fungal endophytes
Michele T. Hoffman and A. Elizabeth Arnold
Division of Plant Pathology and Microbiology School of Plant Sciences 1140 E. South Campus Drive University of Arizona Tucson, AZ 85721 USA
For correspondence: AE Arnold, Tel: 520.621.7212; Fax: 520.621.9290; Email:
[email protected]
Running head: Endohyphal bacteria of endophytic fungi
Contains 3 tables, 5 figures, 4 appendices
52
Abstract Both the establishment and outcomes of plant-fungal symbioses can be influenced by abiotic factors, the interplay of fungal and plant genotypes, and additional microbes associated with fungal mycelia. Recently, bacterial endosymbionts have been documented in soilborne Glomeromycota and Mucoromycotina, and in at least one species each of mycorrhizal Basidiomycota and Ascomycota. Here, we show for the first time that phylogenetically diverse endohyphal bacteria occur in living hyphae of diverse foliar endophytes, including representatives of four classes of Ascomycota. We examined 414 isolates of endophytic fungi, isolated from photosynthetic tissues of six species of cupressaceous trees in five biogeographic provinces, for endohyphal bacteria using microscopy and molecular techniques. Viable bacteria were observed within living hyphae of endophytic Pezizomycetes, Dothideomycetes, Eurotiomycetes, and Sordariomycetes from all tree species and biotic regions surveyed. A focus on 29 fungal/bacterial associations revealed that bacterial and fungal phylogenies were incongruent with each other and with taxonomic relationships of host plants. Overall, eight families and 15 distinct genotypes of endohyphal bacteria were recovered; most were Proteobacteria, but a small number of Bacillaceae also were found, including one that appears to occur as an endophyte of plants. Frequent loss of bacteria following subculturing suggests a facultative association. Our study recovered distinct lineages of endohyphal bacteria relative to previous studies, is the first to document their occurrence in foliar endophytes representing four of the most species-rich classes of fungi, and highlights for the first time their diversity and phylogenetic relationships
53
with regard both to the endophytes they inhabit and the plants in which these endophyte-bacterial symbiota occur.
Keywords: Ascomycota, endohyphal bacteria, Bayesian analysis, Burkholderiales, Cupressaceae, diversity, fungal endophytes, phylogeny, FISH
54
Introduction Traits related to the establishment and outcome of plant-fungal symbioses can reflect not only abiotic conditions and the unique interactions of particular fungal and plant genotypes (50, 51, 57, 60, 63, 68), but also additional microbes that interact intimately with fungal mycelia (4, 12, 43). For example, mycorrhizosphere-associated actinomycetes release volatile compounds that influence spore germination in the arbuscular mycorrhizal (AM) fungus Gigaspora margarita (Glomeromycota) (14). Levy et al. (35) describe Burkholderia spp. that colonize spores and hyphae of the AM fungus Gigaspora decipiens and are associated with decreased spore germination. Diverse "helper" bacteria have been implicated in promoting hyphal growth and the establishment of ectomycorrhizal symbioses (23, 26, 58, 71). Minerdi et al. (44) found that a consortium of ectosymbiotic bacteria limited the ability of the pathogen Fusarium oxysporum to infect and cause vascular wilts in lettuce, with virulence restored to the pathogen when ectosymbionts were removed. In addition to interacting with environmental and ectosymbiotic bacteria, some plant-associated fungi harbor bacteria within their hyphae (first noted as 'bacteria-like organisms' of unknown function) (39). These bacteria, best known from living hyphae of several species of Glomeromycota and Mucoromycotina, can alter fungal interactions with host plants in diverse ways (see 12, 32, 52). For example, the vertically transmitted bacterium Candidatus Glomeribacter gigasporarum colonizes spores and hyphae of the arbuscular mycorrhizal (AM) fungus Gigaspora gigasporarum (9, 10). Removal of the bacterial partner from the fungal spores suppresses fungal growth and development,
55
altering the morphology of the fungal cell wall, vacuoles, and lipid bodies. (38). In turn, the discovery of phosphate-solubilizing bacteria within Glomus mossae spores (45), coupled with the recovery of a P-transporter operon in Burkholderia sp. from Gigaspora margarita (55), suggest a competitive role in phosphate acquisition and transport by these bacteria within the AM symbiosis. Within the Mucoromycotina, Partida-Martinez and Hertweck (52) reported that a soilborne plant pathogen, Rhizopus microsporus, harbors endosymbiotic Burkholderia that produce a phytotoxin (rhizoxin) responsible for the pathogenicity of the fungus. These examples, coupled with the discovery of bacteria within hyphae of ectomycorrhizal Dikarya (Tuber borchii; Ascomycota; Laccaria bicolor and Piriformospora indica; Basidiomycota) (5, 6, 7, 59), suggest that the capacity to harbor endohyphal bacteria is widespread among fungi. To date, however, endocellular bacteria have been documented only from fungi that occur in the soil and rhizosphere (12, 32). Here we report for the first time that phylogenetically diverse bacteria occur within living hyphae of foliar endophytic fungi, including members of four classes of filamentous Ascomycota. We use a combination of light and fluorescent microscopy to visualize bacterial infections within living hyphae of representative strains. Then, drawing from surveys of endophytes from asymptomatic foliage of cupressaceous trees in five biogeographic provinces, we provide a first characterization of the phylogenetic relationships, host associations, and geographic distributions of endohyphal bacteria associated with focal fungal endophytes.
56
Methods Conifers (Pinophyta or Coniferae), comprising ca. 630 species, form the dominant tree component in major biomes such as boreal forests, temperate tree savannas, and temperate rainforests (20, 21, 22). The most widely distributed family of conifers, the Cupressaceae (cypresses and relatives), includes 30 genera and ca. 142 species in seven proposed subfamilies (25). Eleven genera are placed in the subfamily Cupressoideae, including three examined in this study (Cupressus, Juniperus, and Platycladus). We collected foliar endophytes from six species of Cupressoideae, as available, in five localities (Table 1): semi-deciduous forest in the piedmont of North Carolina, USA (NC), and four distinct regions of Arizona, USA (AZ) selected on the basis of distinctive biogeographic histories and the occurrence of desired plant species. These sites included the Chuska Mountains (CHU) of northeastern AZ, with petran montane coniferous forest featuring significant elements of the Rocky Mountain flora; the Mogollon Rim (MOG) and central mountains, including Mingus Mountain and the Bradshaw Mountains, which lie at the interface of the Great Basin coniferous woodlands, interior chaparral, and petran montane forest; the Sky Island archipelago (SKY) of southeastern AZ, including the Santa Catalina and Chiricahua Mountains, characterized by madrean evergreen forest; and the Campus Arboretum at the University of Arizona (UA), a semi-urban, cultivated setting within the Arizona upland province of the Sonoran Desert.
57
Endophyte collection Fresh, asymptomatic photosynthetic tissue was collected from at least three healthy, mature individuals of each focal species in each locality during the growing seasons of 2004-2007. From each tree, healthy, mature photosynthetic tissue was collected from three haphazardly selected branches at the outer canopy ca. 1-2 m above ground. Material was transferred to the laboratory for processing within 6-12 hours of collection. Tissue samples were washed in running tap water and then cut into 2mm segments, corresponding to infection domains for individual fungi (3). Segments were surface-sterilized by rinsing in 95% ethanol for 30 seconds, 10% Clorox® (0.6% sodium hypochlorite) for two minutes, and 70% ethanol for two minutes (1), allowed to surfacedry under sterile conditions, and plated on 2% malt extract agar (MEA), which encourages growth by a diversity of endophytes (24). In sum, 384 tissue segments (96 per individual) were plated per species at each location, with the exceptions of Juniperus deppeana in the Chiricahua Mountains (144 segments), and Platycladus orientalis and Juniperus virginiana from North Carolina (96 segments/species). Plates were incubated at room temperature and inspected for hyphal growth daily for 18 weeks. Emerging fungi were isolated on 2% MEA, archived as living vouchers in sterile water, and deposited at the Robert L. Gilbertson Mycological Herbarium at the University of Arizona (ARIZ; accession numbers available from MH). Seven isolates of bacteria obtained directly from surface-sterilized plant tissue (i.e.,
58
bacterial endophytes) were accessioned as sterile cultures and maintained in sterile 80% glycerol at -80°C.
PCR amplification and sequencing Total genomic DNA was extracted directly from axenic fungal cultures following Arnold et al. (2). In addition, DNA was extracted a second time from mycelium of five isolates using the QIAGEN® DNeasy® Plant Mini kit to ensure that none of our records of endohyphal bacteria represented contamination of reagents in our standard DNA extraction protocol. Our two extraction methods were consistent in terms of DNA extraction quality, sequence quality, and resulting diagnoses of bacterial infection. For each fungal isolate, we used PCR to amplify the nuclear ribosomal internal transcribed spacers and 5.8s gene (ITSrDNA), and when possible, the first 600bp of the large subunit (LSUrDNA) as a single fragment (ca. 1000-1200bps in length) using primers ITS1F and ITS4 or LR3 (27, 56, 67, 70). Each 25µl reaction included 12.5 µl of Sigma ReadymixTM REDTaqTM, 1 µl of each primer (10 µM), 9.5 µl of PCR-quality water, and 1 µl of DNA template. Cycling reactions were run with MJ Research PTC200TM thermocyclers and consisted of 94°C for 3 minutes; 36 cycles of 94°C for 30 seconds, 54°C for 30 seconds, 72°C for 1 minute; and 72°C for 10 minutes. The presence or absence of bacteria within the surrounding matrix was determined initially using light microscopy. Fungal isolates were examined after one week of growth in pure culture on 2% MEA using a Leica DM4000B with bright field imaging (400X; NA = 0.75). Once visual examination ruled out non-endohyphal
59
bacteria (i.e., contaminants in the medium or microbes on hyphal surfaces), total genomic DNA extracted from fresh mycelia was examined using PCR primers specific to bacterial 16S rDNA: 10F and 1507R (1497 bp) or 27F and 1429R (1402 bp) (34, 46). PCR reaction mixes and cycling parameters were as described above, except that annealing temperatures were 58°C (10F and 1507R) or 55°C (27F and 1429R). SYBR® Green I stain (Molecular Probes, Invitrogen) was used to detect DNA bands on 1.5% agarose gels. Positive PCR products were cleaned, quantified, and normalized at the University of Arizona Genomic Analysis and Technology Core facility (GATC), followed by bidirectional sequencing with PCR primers (5 µM) using an Applied Biosystems 3730XL DNA Analyzer. PCR products of insufficient concentration for sequencing were cloned using Agilent Technologies StrataClone™ cloning kits following the manufacturer’s instructions. Water was used in place of template for negative controls. Positive clones were amplified and sequenced using primers M13F and M13R. The software applications phred and phrap (18, 19) were used to call bases and assemble bidirectional reads into contiguous consensus sequences, with automation provided by ChromaSeq (40) implemented in Mesquite v. 2.6 (41). Base calls were verified by inspection of chromatograms in Sequencher™ v 4.5 (Gene Codes Corp., Ann Arbor, MI). ITSrDNA and partial LSUrDNA sequence data have been submitted to GenBank under accession numbers GQ153054-GQ153264 or were published previously by the authors (28), and bacterial 16S rDNA sequences under accession numbers HM046622-HM046627 and HM117722-117749.
60
Diversity and taxonomic placement of endophytes Previous studies have used 90-97% ITS rDNA sequence similarity to estimate species boundaries for environmental samples of fungi (97%, O’Brien et al. (49); 90%, Hoffman & Arnold (28); 95%, Arnold et al. (1). Empirical estimates of percent sequence divergence between sister species for four endophyte-containing ascomycetous genera (Botryosphaeria, Colletotrichum, Mycosphaerella, and Xylaria) indicated that groups based on 95% ITS rDNA sequence similarity conservatively estimate species (66). We used Sequencher™ v 4.5 (Gene Codes Corp., Ann Arbor, MI.; default settings except for the requirement of 20 character minimum overlap; see Arnold et al. (3)) to estimate fungal OTU at 95% ITSrDNA sequence similarity, and bacterial OTU at 97% 16S sequence similarity (62). Taxonomic placement for fungi at the level of order and above was estimated by BLAST comparisons with the curated ITSrDNA database for fungi maintained by the Alaska Fungal Metagenomics Project (FMP; http://www.borealfungi.uaf.edu/) and phylogenetic analysis (below). Bacterial taxonomy was estimated by BLAST comparisons with GenBank and the Ribosomal Database Project (RDP; release 9) classifier program (Naive Bayesian rRNA Classifier Version 2.0, July 2007) (15, 69), combined with phylogenetic analyses (below). All statistical analyses were performed in JMP 7.0 (SAS Institute Inc. Cary, NC).
61
Live-Dead stain Live-Dead stain was used to confirm the presence and viability of endohyphal bacteria for fungal isolates identified as containing bacteria on the basis of light microscopy and PCR (above). We treated fresh mycelia in sterile water with 2µl of Molecular Probe Live-Dead fluorescent stain for 15 min and prepared slides with VECTASHIELD® HardSet™ 1400 medium (Vector Laboratories, Inc.) to prevent photobleaching. A Leica DM4000B microscope with Luminera camera and 100W mercury arc lamp was used for fluorescent imaging with a Chroma Technology filter set 35002 (480nm excitation/520nm emission) and 1000X APO oil objective (NA=1.40-0.70; 0.13mm working distance) at room temperature. Visible fluorescence of bacterial nucleic acids within living hyphae that was not consistent with fungal mitochondrial or nuclear DNA, coupled with positive PCR results and a lack of extrahyphal or ectosymbiotic bacteria, provided evidence of viable endohyphal bacteria (47, 64).
Fluorescent in situ hybridization (FISH) To rule out misinterpretation of fluorescence results from Live-Dead analyses we used FISH with a probe specific for eubacteria to confirm endocellular infection in two focal isolates (9084b and 9143). In each case, we examined hyphae from cultures grown on 2% MEA and on 2% MEA amended with antibiotics, which removed evident bacterial infections (Hoffman and Arnold, in prep.). Fresh mycelium was harvested and suspended in 1000ul 1X phosphate buffered saline (PBS). After centrifugation at 8000rpm for 5 minutes, PBS buffer was replaced
62
with fresh 4% formaldehyde solution and incubated for 1.5 hours at 4°C. Samples then were centrifuged at 8000rpm for 5 minutes and the pellet resuspended in PBS buffer twice prior to storage in absolute ethanol at -20°C. Fixed mycelium was collected to 0.5 ml tubes and dehydrated in a series of ethanol/PBS solutions beginning with 50%, 70%, and finally 95% ethanol. Dehydrated mycelium was incubated for 90 minutes at 46°C in 10ul of hybridization buffer (HB) using 40% formamide hybridization stringency (800ul formamide, 800ul DEPC water, 500ul EDTA) with 2ul of EUB338 probe (10uM), a universal 16S rDNA oligonucleotide probe (labeled with 5' TAMRA fluorochrome tag; 5' GCTGCCTCCCGTAGGAGT 3'; EX559nm/EM583nm; Integrated DNA Technologies, Inc.). Each sample was rinsed in 100ul wash buffer (460ul 5M NaCl, 1ml 1M Tris, 50ul SDS, DEPC water, fill to 50ml) and warmed to 46°C twice. This method was repeated using [1] antibiotic-cured isolates of each strain, which showed no evidence of endohyphal bacteria; and [2] hybridization buffer and PBS only, but no EUB338 probe, to control for autoflorescence. Mycelium was mounted on gelatin-coated glass slides using DAPI as a counter-stain (42) and examined with either an Olympus BX61 with a mercury arc lamp or a Leica DMI 6000 confocal system equipped with a 543nm laser (63X oil; 1024x1024 format; xyz acquisition; line average =1; frame average = 4). Leica software LAS-AF v 1.8.2 was used to capture images.
Phylogenetic analyses Based on positive 16S rDNA PCR results and the lack of any extrahyphal bacteria in our axenic cultures, endohyphal bacteria were observed in 75 of 414 fungal isolates. Twenty-
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one infected isolates representing a broad diversity of Pezizomycotina, and 49 isolates in which infections were never observed and that represented a broad array of ITS rDNA genotype groups, were sequenced for 1200 bps of LSU rDNA following Arnold et al. (3) (primers LROR or LR3R and LR7; protocols as described above). Additional LSU rDNA sequence data have been submitted to GenBank under accession numbers HM117750HM117756. These 70 sequences were integrated into core alignments of 157 representative Ascomycota, partitioned by class into Sordariomycetes, Dothideomycetes, and Eurotiomycetes (28) using Mesquite v 2.6 (41). Details of taxon sampling are given in Table 2 and Appendix 1. Because our sample of Pezizomycetes contained only one isolate with strong evidence of endohyphal bacteria, we did not address phylogenetic relationships of pezizomycetous endophytes here. Alignment for each class, based on LSU rDNA secondary structure defined by Saccharomyces cerevisiae (13), was performed using ClustalW with manual adjustment in Mesquite. All ambiguously aligned regions were excluded, with matrix details given in Appendix 2. Alignments are available online at http://arnoldlab.net/alignments.html. Phylogenetic relationships for members of each class were inferred using parsimony and Bayesian Metropolis-coupled Markov chain Monte Carlo (MCMCMC) analyses. For the former, five sets of 200 heuristic searches with random stepwise addition and tree bisection-reconnection (TBR) were implemented using PAUPRat, which incorporates the ‘parsimony ratchet’ method (28, 48, 61) in PAUP* 4b10 (65). The ‘filter trees’ command in PAUP* 4b10 was used to select all shortest trees, which
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were used to assemble strict consensuses. Bootstrap support was determined using 1000 replicates and the default settings for nonparametric NJ analysis as implemented in PAUP* 4b10. Clades with ≥70% bootstrap support were considered strongly supported. Details of parsimony searches are given in Appendix 2. For Bayesian analyses, Modeltest 3.7 (54) was used to select the appropriate model of evolution; in each case, GTR+I+G was selected using the Akaike Information Criterion. Analyses were executed in Mr. Bayes 3.1.2 (31) for four runs of up to 6 million generations each, initiated with random trees, four chains, and sampling every 1000th generation. Likelihoods converged to a stable range for each data set and all trees prior to convergence were discarded as burn-in. Clades with > 95% posterior probability values were considered to have significant support. Twenty-nine bacterial 16S DNA sequences, derived from total genomic DNA extractions of fungal endophyte mycelia with no evidence of extrahyphal bacteria (Table 2), and seven 16S sequences from bacterial endophytes obtained directly from plant material (Table 3), were incorporated into a core alignment of 30 sequences of named bacterial taxa obtained from the Ribosomal Database Project 9 website (Cole et al. (15); http://rdp.cme.msu.edu) using seqmatch (type strains, isolates, >1200 bp, good quality; ca. 1400bp), together with eight additional sequences from GenBank that represented endosymbionts of fungi isolated in previous studies (Appendix 3) (9, 10, 53). Data were aligned using NAST and screened for chimeric sequences (minimum sequence length = 300 bp) (http://greengenes.lbl.gov/) (17). Ambiguous regions were excluded, resulting in a matrix of 6300 characters (http://arnoldlab.net/alignments.html).
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For the bacterial data set, a heuristic search using maximum parsimony with random step-wise additions and TBR branch swapping was implemented in PAUP* 4b10, resulting in 167 optimal trees of 3668 steps. Support was assessed using a nonparametric neighbor-joining bootstrap (1000 replicates). Bayesian MCMCMC analysis was implemented in MrBayes v. 3.1.2 for 2.5 million generations, initiated with random trees, four chains, and sampling every 1000th tree, using GTR+I+G based on evaluation in Modeltest 3.7 (54). After elimination of the first 1,400 trees as burn-in, the remaining 1,100 trees were used to infer a majority rule consensus. These results were complemented by phylogenetic inference in ARB v. 7.12.07 (37) using the AxML maximum likelihood (ML) method (FastDNAML; olsen/felsenstein (F84); no filter: all positions included in phylogenetic analysis; -ln likelihood = 22430.51). Branch support was assessed using parametric ML bootstrapping (100 replicates) and Bayesian posterior probabilities.
Results Cultivable endophytes were isolated from healthy foliage of all plant species and from every study site (Table 1), yielding 414 isolates from 4,560 tissue segments. ITS rDNA data for all isolates were assembled at 95% sequence similarity to yield 113 OTU (putative species; Fisher’s alpha = 51.2). Representatives of 5 classes and approximately 10 orders, 13 families, and 28 genera were recovered. The most common orders differed among host species and sites, but the entire dataset comprised a high frequency of Pezizales (Pezizomycetes), Capnodiales, Pleosporales, Botryosphaeriales,
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Dothideales (Dothideomycetes), Helotiales (Leotiomycetes), and Xylariales (Sordariomycetes) (Appendix 4). Host specificity, diversity, and geographic distributions of these fungi will be addressed in a forthcoming paper (Hoffman & Arnold, in prep.).
Endohyphal bacteria Endohyphal bacteria initially were observed in 75 of 414 endophyte isolates (18%; determined by positive PCR results using 16S primers and no evidence of extrahyphal bacteria), including isolates from all four classes of Ascomycota recovered in our surveys (Pezizomycetes, Dothideomycetes, Eurotiomycetes, and Sordariomycetes), isolates from all three host genera (Cupressus, Juniperus, and Platycladus), and isolates from plants in all biogeographic regions (Table 2). After 47 of these isolates were subcultured, however, they later were screened as negative for bacteria. In general, length of time in culture (after ca. 2 weeks) appeared to adversely affect the detectability or persistence of bacterial infections. Based on examination with light microscopy and Live-Dead stain, no isolate scored as positive for endohyphal bacteria had evident extrahyphal or ectosymbiotic bacteria. Live-Dead stain indicated that bacteria within hyphae were viable and confirmed viability of the hyphae that housed them (data not shown). Fluorescence in situ hybridization (FISH) for isolates 9084b and 9143 further confirmed the presence of endohyphal bacteria within living mycelia. Isolate 9084b is shown with the EUB338 fluorescent probe mounted in anti-fading mount medium only (Fig. 1A). Isolate 9143 is
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shown with the EUB338 probe and DAPI counter-stain (Fig. 1B), illustrating the presence of small numbers of bacterial cells in comparison to significant amounts of total genomic DNA contained in the hyphal structure.
Phylogenetic inferences and taxonomic placement Phylogenetic analyses confirmed BLAST-level taxonomy at the class level for fungal endophytes in the Dothideomycetes, Eurotiomycetes, and Sordariomycetes (Figs. 3-5). No pattern regarding the structure of endophyte lineages as a function of host plant taxonomy was evident (i.e., the sister relationship of Cupressus and Juniperus, and their sister relationship to Platycladus, is not reflected in the phylogenetic relationships of their fungal associates). Isolates containing bacterial associates were spread broadly across endophyte-containing clades in each class, indicating a phylogenetically widespread capacity to harbor bacteria. Taxonomic placement of endohyphal bacteria initially was assessed using BLAST comparisons in GenBank, typically yielding matches to unidentified or nameless environmental samples. The RDP classifier (69) placed these bacteria into two phyla, Proteobacteria (including Alphaproteobacteria, Betaproteobacteria, and Gammaproteobacteria) and Firmicutes. Overall, we recovered five orders and 10 families, putatively identified as Sphingomonadaceae (Alphaproteobacteria), Burkholderiaceae, Comamonadaceae, and Oxalobacteraceae (Betaproteobacteria), Moraxellaceae, Xanthomonadaceae, Pasteurellaceae, and Enterobacteriaceae (Gammaproteobacteria), and Bacillaceae and Paenibacillaceae (Firmicutes) (Table 2).
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Based on 97% 16S rDNA identity, these bacteria represented 15 OTU. Of these, nine were found only once. Three of the remaining six OTU were found in fungi from multiple genera of Cupressaceae, four were found in fungi from more than one biogeographic region, and all were found in fungi representing multiple genera. Four genotypes each were found in fungi representing different classes of Pezizomycotina (genotypes A, B, C, E) (Table 2). Proteobacteria were observed in association with fungi of the Dothideomycetes, Eurotiomycetes, and Sordariomycetes, and in endophytes from all three plant genera representing hosts in AZ and NC. Firmicutes were found in Eurotiomycetes and one member of the Pezizomycetes. Nominal logistic analysis provided no evidence for a significant effect of host genus (Cupressus, Juniperus, or Platycladus), region (AZ vs. NC), or fungal class on the incidence of major bacterial groups (Alpha-, Beta-, and Gammaproteobacteria; Firmicutes) (P >0.05 for all effects). Phylogenetic relationships of endohyphal bacteria from fungal endophytes were not congruent with those of the endophytes they inhabited (Figs. 2-5), nor with relationships among host plants (data not shown). The bacterial lineages recovered here were distinct from those recognized previously as endosymbionts of fungi (Fig. 2).
Comparison of bacterial endophytes and endohyphal bacteria Only one bacterial genotype was represented both among bacterial endophytes (isolated from surface-sterilized plant tissue only) and putative endohyphal bacteria (genotype A, Bacillales; Tables 2 and 3). This genotype was isolated directly from foliage of J.
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osteosperma (Chuska Mountains) and C. arizonica and J. deppeana (Sky Islands), and was amplified from genomic DNA from a fungal culture recovered from J. deppeana (Sky Islands). Two additional endophytes recovered here also were Bacillaceae, representing genotypes that never were found as endohyphal bacteria (genotypes H, J) (Tables 2, 3). One bacterial endophyte represented the Enterobacteriaceae (putative Erwinia) and was not found as a bacterial endosymbiont within fungal hyphae.
Discussion Recent studies have highlighted the potential of endosymbiotic microbes, including bacteria and viruses, to shape the ecological roles of fungi (e.g., 43, 52). Endohyphal bacteria have been found previously in living hyphae of plant-associated Glomeromycota, Mucoromycotina, and up to three taxa of mycorrhizal Dikarya. Our study is the first to compare their phylogenetic relationships with diverse fungal hosts, and the first to document their occurrence in ascomycetous endophytes of foliage representing four of the most species-rich classes of Ascomycota (Pezizomycetes, Sordariomycetes, Dothideomycetes, and Eurotiomycetes), which together include numerous plant pathogens, saprotrophs, and pathogens and parasites of animals (31). Coupled with previous studies demonstrating the presence of bacteria in living mycelia of mycorrhizal or soil-borne fungi (6, 7, 9, 52), our data provide strong evidence that the ability of fungi to harbor endohyphal bacteria is phylogenetically widespread. Our survey data show that endohyphal bacteria-endophyte associations occur in regions with divergent biogeographic histories and markedly different environmental
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conditions (e.g., diverse, aridland montane systems vs. piedmont forest in southeastern North America) and in fungi associated with three genera of plants. Screening of a small number of isolates obtained in previous studies (Figs. 3-5) confirmed that endohyphal bacteria occur in endophytes from additional biogeographic provinces and host plant lineages: infections were observed in endophytes isolated from Pinaceae in boreal forest (endophyte 4466 from Picea mariana in Canada) and two families of angiosperms from a tropical forest (endophyte 6722 from Faramea occidentalis, Rubiaceae, and endophyte 6731 from Swartzia simplex, Fabaceae, recovered at Barro Colorado Island, Panama). Because infections either were lost or became more difficult to detect following subculturing, and because of potential limitations of our microscopy, screening methods and primer selection, our results likely underestimate the incidence of endohyphal bacteria in these plant-associated fungi. Examination with Live-Dead stain provided evidence that bacteria within fungal hyphae were viable, and showed that such bacteria occur within living fungal tissues. FISH confirmed these results by diagnosing the presence of bacteria using 16S rDNA specific tags (Fig. 1). In turn, our phylogenetic inferences show that endohyphal bacteria include a greater phylogenetic diversity – both within and beyond the Proteobacteria – than was previously known (23, 33, 53; Fig. 2). In contrast to previous studies, which have focused primarily on bacteria associated with a particular fungal species or closely related group of species, we addressed the diversity of bacteria associated with a phylogenetically broad sample of ecologically similar fungi. Our analyses provide no evidence of co-cladogenesis: fungal
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and bacterial trees were obviously incongruent, and neither matched the phylogenetic relationships among the plant species surveyed here. Our phylogenetic results are consistent with the hypothesis of horizontal transmission both for fungal endophytes, as expected (1, 28), and for the bacterial symbionts examined here (Figs. 2-5). Using cultivation media infused with antibiotics we have found that many of these fungi can be cured of their endosymbionts (Hoffman & Arnold, in prep.), and we currently are investigating methods to re-infect cured mycelia. These attempts will shed light on our hypotheses regarding the facultative nature of these associations, their transmission modes, and their costs and benefits with regard to the fungi they inhabit. Notably, some endophytes were unable to grow when transferred to media containing antibiotics (Hoffman & Arnold, in prep.), suggesting that the relationship may be more intimate or tend toward greater obligacy in some pairs of fungi/bacteria than others. For one isolate (9143), we have successfully isolated the bacterial partner from the fungus and confirmed its identity using 16S rDNA sequencing (Hoffman et al., in prep.). Attempts to reinfect the endophyte with this bacterium have been ineffective to date. Discovering the diversity and ecological roles of fungal endophytes encompasses a trove of future research that will be important for understanding plant ecology and evolution. So too does elucidation of the incidence, diversity, and ecological importance of their endohyphal bacteria, which appear to be common but previously overlooked inhabitants of plant-symbiotic fungi. Evidence from a variety of studies (reviewed by 12, 32) indicates that endohyphal bacteria have the capacity to alter the outcome of plant-fungal interactions in a phylogenetically diverse array of
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symbioses. Experimental assessment of such effects will provide key evidence to understanding the degree to which bacterial associates influence the nature of endophytic symbioses. In turn, incorporating ancestral state reconstructions and further sampling of both endohyphal bacteria (of fungi) and bacterial endophytes (of plants) will elucidate the degree to which bacteria have transitioned from endophytic to fungalendosymbiotic, or the reverse – or have followed a distinctive evolutionary trajectory.
Acknowledgments We thank the Division of Plant Pathology and Microbiology, School of Plant Sciences, and College of Agriculture and Life Sciences at the University of Arizona, and the Arizona-Nevada Academy of Sciences, for supporting this work. Additional support from the National Science Foundation (NSF-0626520 and 0702825 to AEA; NSF-IGERT to MTH) is gratefully acknowledged. We thank D.R. Maddison for sharing pre-release versions of Mesquite and Chromaseq; M. Gunatilaka, M. Shimabukuro, D. Grippi, M. J. Epps, and C. Weeks-Galindo for lab assistance and helpful discussion; J. U’Ren, F. Lutzoni, E. Gaya, J. Miadlikowska, and F. Santos-Rodriguez for isolation of samples from the Chiricahua Mountains; and B. Klein and undergraduate students at Diné College for isolation of samples from the Chuska Mountains on the Navajo Nation. We are especially grateful to H. Van Etten and R. Palanivelu for access to microscopy facilities, A. Estes for EUB338 probes and helpful discussion about FISH, and J. L. Bronstein for comments on the manuscript. This paper represents a portion of the doctoral dissertation research of M.H. in Plant Pathology and Microbiology at the University of Arizona.
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80
Table 1. Locations and characteristics of sampling sites, and plant species surveyed for endophytes in each location.
Habitats
Latitude, Longitude
Elevation (m)
Mean annual temperaturea (°C)
Mean annual precipitation a (mm)
Temperate mixed forest
36.00°N, 78.94°W
97
15.5
1092
J. virginiana L., P. orientalis (L.) Franco
Ponderosa pine forest
36.16°N, 108.91°W
2200
10.4
253
J. osteosperma (Torr.) Little, J. scopulorum Sarg.
Iron Springs, AZ (IS)
Pinyon-juniper woodland
34.58°N, 112.60°W
1851
11.8
482
C. arizonicab Greene, J. deppeana Stead
Mingus Mtn, AZ (MM)
Ponderosa pine-pinyon-juniper woodland
34.70°N, 112.13°W
2354
15.3
483
J. deppeana Stead
Strawberry, AZ (ST)
Ponderosa pine-pinyon-juniper woodland
34.40°N, 111.49°W
1707
13.2
541
J. deppeana Stead
Mt Lemmon, AZ (MTL)
Madrean evergreen woodland
32.44°N, 110.78°W
1600
9.2
751
C. arizonicab Greene, J. deppeana Stead
Chiricahua Mtns AZ (CHI)
Madrean evergreen woodland
32.03°N, 109.38°W
1371
11.7
549
J. deppeana Stead
Cultivated trees, semi-urban setting
32.23°N, 110.95°W
787
20.2
305
C. arizonicab Greene, J. virginiana L., P. orientalis (L.) Franco
Site
Plant species
Piedmont, Central NC (DUKE) Duke Forest, NC (NC) Petran forest, NE AZ (CHU) Chuska Mtns, AZ (CHU) Mogollon Rim, Central AZ (MOG)
Sky Islands , SE AZ (SKY)
Campus Arboretum, SE AZ (UA) Univ. of Arizona (UA)
a b
Climate information obtained from http://www.wrcc.dri.edu/coopmap/ Recently designated change from Cupressus arizonica Greene to Callitropsis arizonica (Greene) is acknowledged (36).
81
Table 2. Taxonomic classification for 29 fungal endophytes and associated endohyphal bacteria. Bacterial taxonomic classifications are based on BLAST results and Ribosomal Database project (RDP) classifier program (Naive Bayesian rRNA Classifier Version 2.0, July 2007) (69). Fifteen distinct 16S rDNA OTU, based on 97% sequence similarity, are designated in groups A-R. Asterisks indicate one genotype found also as a bacterial endophyte in plant tissue, such that its status as only an endosymbiont of fungi is not clear (Genotype A; see Table 3). Fungal identifications were based on comparisons with the FMP curated database in March 2009 and are approximate at the genus and species levels, but strongly supported at the class level based on phylogenetic analyses (Figs. 3-5).
Fungal class
16S genotype
Top 16S BLAST/GenBank acc.
Bacterial lineage (based on RDP classifier)
Dothidea sambuci AY930108.1
Dothideomycetes
K
AZ: UA
Alternaria mali AY154683.1
Dothideomycetes
L
P. orientalis
AZ: UA
Phoma glomerata AY183371.1
Dothideomycetes
M
9055
P. orientalis
AZ: UA
Alternaria mali AY154683.1
Dothideomycetes
C
9060
P. orientalis
AZ: UA
Aureobasidium pullulans AY225166.1
Dothideomycetes
E
9094
P. orientalis
AZ: UA
Lecythophora sp. AY219880.1
Sordariomycetes
E
9096
P. orientalis
AZ: UA
Hormonema sp. AF182378.1
Dothideomycetes
N
9106
C. arizonica
AZ: UA
Preussia africana DQ865095.1
Dothideomycetes
C
9107
C. arizonica
AZ: UA
Phoma glomerata AY183371.1
Dothideomycetes
F
9112
C. arizonica
AZ: UA
Lecythophora sp. AY219880.1
Sordariomycetes
P
9120
C. arizonica
AZ: UA
Monodictys sp. AJ972795.1
Sordariomycetes
E
9122
P. orientalis
NC: Duke
Pestalotiopsis caudata EF055188.1
Sordariomycetes
C
Acinetobacter sp. FJ422393 Acinetobacter junii NR_026208 Uncultured Sphingomonas EU341143 Uncultured bacterium clone DQ984599 Uncultured bacterium clone EU236303 Uncultured bacterium clone EU236303 Oxalobacteraceae sp. DQ490310 Uncultured bacterium clone DQ984599 Uncultured Haemophilus sp. EU071476 Variovorax paradoxus DQ257419 Uncultured bacterium clone EU236303 Uncultured bacterium clone DQ984599
Gammaproteobacteria, Pseudomonadales, Moraxellaceae Gammaproteobacteria, Pseudomonadales, Moraxellaceae Alphaproteobacteria, Sphingomonadales, Sphingomonadaceae Gammaproteobacteria, Xanthomonadales, Xanthomonadaceae Betaproteobacteria, Burkholderiales, Burkholderiaceae Betaproteobacteria, Burkholderiales, Burkholderiaceae Betaproteobacteria, Burkholderiales, Oxalobacteraceae Gammaproteobacteria, Xanthomonadales, Xanthomonadaceae Gammaproteobacteria, Pasteurellales, Pasteurellaceae Betaproteobacteria, Burkholderiales, Comamonadaceae Betaproteobacteria, Burkholderiales, Burkholderiaceae Gammaproteobacteria, Xanthomonadales, Xanthomonadaceae
Isolate
Host plant
Site
2611
J. scopulorum
AZ: CHU
9026
P. orientalis
9054
Top BLAST match
82
9128
P. orientalis
NC: Duke
Cladosporium oxysporum AJ300332.1
Dothideomycetes
E
9133
P. orientalis
NC: Duke
Botryosphaeria dothidea AY640254.1
Dothideomycetes
C
9135
P. orientalis
NC: Duke
Phyllosticta spinarum AF312009.1
Dothideomycetes
C
9140
P. orientalis
NC: Duke
Microdiplodia sp. EF432267.1
Dothideomycetes
R
9143
P. orientalis
NC: Duke
Pestalotiopsis caudata EF055188.1
Sordariomycetes
C
9145
P. orientalis
NC: Duke
Microdiplodia sp. EF432267.1
Dothideomycetes
D
9147
P. orientalis
NC: Duke
Penicillium citreoni AY373908.1
Eurotiomycetes
E
11123
J. virginiana
AZ: UA
Pithya cupressina U66009.1
Pezizomycetes
G
11164
J. deppeana
AZ: MTL
Eurotiomycetes
A*
11259
J. deppeana
AZ: MTL
Sordariomycetes
B
11272
J. deppeana
AZ: MOG
Eurotiomycetes
B
11035
J. deppeana
AZ: MTL
Phaeomoniella chlamydospora AY772236.1 Biscogniauxia. mediterranea AJ390413.1 Phaeomoniella chlamydospora AB278179.1 Pestalotiopsis besseyi AY687313.1
Sordariomycetes
A*
9084b
P. orientalis
AZ: UA
Aureobasidium pullulans DQ680686.1
Dothideomycetes
F
9106 clA
C. arizonica
AZ: UA
Preussia africana DQ865095.1
Dothideomycetes
E
9106 clB
C. arizonica
AZ: UA
Preussia africana DQ865095.1
Dothideomycetes
O
9126 clB
P. orientalis
NC: Duke
Pestalotiopsis caudata EF055188.1
Sordariomycetes
Q
9149b
P. orientalis
NC: Duke
Phyllosticta spinarum AF312009.1
Dothideomycetes
D
Uncultured bacterium clone EU236303 Uncultured bacterium clone DQ984599 Uncultured bacterium clone DQ984599 Pantoea agglomerans FJ756346.1 Uncultured bacterium clone DQ984599 Pantoea oleae AF130967 Uncultured bacterium clone EU236303 Bacillus foraminis AJ717382 Bacillus subtilis FJ549011 Paenibacillus sp. AM906086 Paenibacillus sp. AM906086 Bacillus atrophaeus FJ194961 Uncultured bacterium clone EF509196 Uncultured bacterium clone EU236303 Acinetobacter sp. FJ267573 Pantoea agglomerans EU598802.1 Pantoea oleae AF130967
Betaproteobacteria, Burkholderiales, Burkholderiaceae Gammaproteobacteria, Xanthomonadales, Xanthomonadaceae Gammaproteobacteria, Xanthomonadales, Xanthomonadaceae Gammaproteobacteria, Enterobacteriales, Enterobacteriaceae Gammaproteobacteria, Xanthomonadales, Xanthomonadaceae Gammaproteobacteria, Enterobacteriales, Enterobacteriaceae Betaproteobacteria, Burkholderiales, Burkholderiaceae Firmicutes, "Bacilli", Bacillales, Bacillaceae Firmicutes, "Bacilli", Bacillales, Paenibacillaceae Firmicutes, "Bacilli", Bacillales, Paenibacillaceae Firmicutes, "Bacilli", Bacillales, Paenibacillaceae Firmicutes, "Bacilli", Bacillales, Paenibacillaceae Gammaproteobacteria, Pasteurellales, Pasteurellaceae Betaproteobacteria, Burkholderiales, Burkholderiaceae Gammaproteobacteria, Pseudomonadales, Moraxellaceae Gammaproteobacteria, Enterobacteriales, Enterobacteriaceae Gammaproteobacteria, Enterobacteriales, Enterobacteriaceae
83
Table 3. Bacterial endophytes isolated from foliage of focal species of Cupressaceae, collection sites, 16S rDNA OTU based on 97% sequence similarity when compared among themselves and with the endohyphal bacteria listed in Table 2, top RDP classifier match and accession information, and bacterial lineage.
Isolate
Plant species
Location
16S genotype
RDP Classifier match
Bacterial lineage
11124
J. deppeana
AZ: MTL
H
Bacillus megaterium FJ527650
Firmicutes, "Bacilli", Bacillales, Bacillaceae
11136
C. arizonica
AZ: MTL
A
Bacillus subtilis FJ549011
Firmicutes, "Bacilli", Bacillales, Bacillaceae
11180
J. osteosperma
AZ: CHU
A
Bacillus sp. FJ514812
Firmicutes, "Bacilli", Bacillales, Bacillaceae
11219
C. arizonica
AZ: MTL
A
Bacillus subtilis NR_024931
Firmicutes, "Bacilli", Bacillales, Bacillaceae
11290
J. deppeana
AZ: MTL
A
Bacillus subtilis FJ549011
Firmicutes, "Bacilli", Bacillales, Bacillaceae
11351
C. arizonica
AZ: MOG
I
Erwinia persicina EU681952
Gammaproteobacteria, Enterobacteriales, Enterobacteriaceae
11394
J. virginiana
AZ: UA
J
Bacillus licheniformis EU071553
Firmicutes, "Bacilli", Bacillales, Bacillaceae
84
Figure legends Figure 1. Fluorescent in situ hybridization (FISH) microscopy showing hyphae of two isolates of endophytic fungi harboring endohyphal bacteria. A (isolate 9084b, Dothideomycetes) shows TAMRA fluorophore at site of internal structure in hyphal cells. B (isolate 9143, Sordariomycetes) shows TAMRA fluorophore with DAPI counterstain (blue), highlighting the fungal nuclear and mitochondrial DNA in addition to the bacterial 16S rDNA (yellow/green). Scale bars: A, 10µm for image A and 25µm for image B.
Figure 2. Phylogenetic relationships of 29 endohyphal bacteria, 7 bacterial endophytes (paired circles), and 38 named taxa based on Bayesian analysis of 16S ribosomal DNA sequences. Branch support values indicate parsimony bootstrap bootstrap proportions (>70%; before slash) and Bayesian posterior probabilities (>95%; after slash). Branches in bold indicate >70% neighbor-joining bootstrap proportions. Named bacterial sequences and GenBank accession numbers are listed in Appendix 3; genotype groups for fungi are indicated by letters. Sequence data representing previously described bacterial endosymbionts of Glomeromycota and Mucoromycotina (9, 10, 52) are marked with solid squares.
Figure 3. Phylogenetic relationships of fungal endophytes with representative Dothideomycetes based on majority rule consensus tree from Bayesian analysis. Branch support values indicate parsimony bootstrap proportions (>70%; before slash) and Bayesian posterior probabilities (>95%; after slash). Taxon labels in bold indicate endophytes isolated in this study and screened for bacterial symbionts. Isolate names are
85
followed by plant species and collection site. Endophytes in which endohyphal bacteria were observed indicate the GenBank accession number for the top 16S BLAST match, taxonomic placement, and 16S rDNA genotype group. Closed circles indicate infections that were observed initially but were lost during cultivation. The open circle (isolate 6731) represents an infected endophyte from another study (1).
Figure 4. Phylogenetic relationships of endophytes with representative Eurotiomycetes based on majority rule consensus tree from Bayesian analysis of 25 LSU rDNA sequences. Branch support values indicate parsimony bootstrap proportions (>70%; before slash) and Bayesian posterior probabilities (>95%; after slash). Taxon labels in bold indicate endophytes isolated in this study and screened for bacterial symbionts. Isolate names are followed by plant species and collection site. Endophytes in which endohyphal bacteria were observed indicate the GenBank accession number for the top 16S BLAST match, taxonomic placement, and 16S rDNA genotype group. The closed circle (isolate 11164) indicates an infection was lost during cultivation. The open circle (isolate 4466) represents an infected endophyte from another study (1).
Figure 5. Phylogenetic relationships of endophytes with representative Sordariomycetes based on majority rule consensus tree from Bayesian analysis of 95 LSU rDNA sequences. Branch support values indicate parsimony bootstrap proportions (>70%; before slash) and Bayesian posterior probabilities (>95%; after slash). Taxon labels in bold indicate endophytes isolated in this study and screened for bacterial symbionts. Isolate names are followed by plant species and collection site. Endophytes in which
86
endohyphal bacteria were observed indicate the GenBank accession number for the top 16S BLAST match, taxonomic placement, and 16S rDNA genotype group. Closed circles indicate infection was lost during cultivation. The open circle (isolate 6722) represents infected endophyte from another study (1).
87
Figure 1.
100/98
Figure 2.
96/99 91/100
90/-
0.10
Candidatus Glomeribacter gigasporarum * AJ251633.1 n Candidatus Glomeribacter gigasporarum * X897272 n 100/Candidatus Glomeribacter gigasporarum * AJ251634.1 n 100/Candidatus Glomeribacter gigasporarum * AJ251635.1 n 98/100 Burkholderia sp 20577 * AJ938141.1 n 100/- Burkholderia rhizoxinica * AJ938142.1 n 100/98 Burkholderia sp 30887 * AJ938143.1 n 100/Burkholderia sp 69968 * AJ938144.1 n 97/100 Burkholderia fungorum * AF215705 98/100 Burkholderia sordidicola * AF512827 Burkholderia cepacia ATCC25416T * AB334766 99/100 99/100 Burkholderia pseudomallei ATCC23343 * CP000573 96/100 Ralstonia solanacearum ATCC11696 * EF016361 Ralstonia mannitolilytica * AJ270258 100/9147 Penicillium * P. orientalis NC * EU236303 * E 99/100 9094 Lecythophora * P. orientalis UA * EU236303 * E 100/100 9060 Aureobasidium * P. orientalis UA * EU236303 * E 100/100/9106clA Preussia * C. arizonica UA * EU236303 * E 100/9120 Monodictys * C. arizonica UA * EU236303 * E 98/9128 Cladosporium * P. orientalis NC * EU236303 *E 99/9096 Hormonema * P. orientalis UA * DQ490310 * N 94/100 9112 Lecythophora * C. arizonica UA * DQ257419 * P 98/100 Nitrosomonas sp. BF16c52 * AF386747 96/100 Xanthomonas campestris LMG568T * AM920689 97/100 Xanthomonas oryzae LMG5047 * NR_026319 100/9133 Botryosphaeria * P. orientalis NC * DQ984599 * C 94/100 9135 Phyllosticta * P. orientalis NC * DQ984599 * C 100/9106 Preussia * C. arizonica UA * DQ984599 * C 100/- 9143 Pestalotiopsis * P. orientalis NC * DQ984599 * C 100/- 9055 Alternaria * P. orientalis UA * DQ984599 * C 98/9122 Pestalotiopsis * P. orientalis NC * DQ984599 * C 99/Pseudomonas syringae * AJ308316 93/9026 Alternaria * P. orientalis UA * FJ596644 * L 99/2611 Dothidea * J. scopulorum CHU * FJ422393 * K 95/9106clB Preussia * C. arizonica UA * FJ267573 * O 91/100 9084b Aureobasidium * P.orientalis UA * EF509196 * F 95/9107 Phoma* C. arizonica UA * EU071476 * F 98/100 9145 Microdiplodia * P. orientalis NC * AF130967 * D 98/9149b Phyllosticta * P. orientalis NC * AF130967 * D 100/Escherichia coli ATCC11775T * NR_024570 100/Erwinia amylovora ATCC15580 * U80195 99/11351 C. arizona IS * EU681952 * I == 100/Pantoea agglomerans ATCC27155 * FJ357815 9126clb Pestalotiopsis * P. orientalis NC * EU598802.1 * Q 98/9140 Microdiplodia * P. orientalis * NC EF432267.1 * R 100/100 9054 Phoma * P. orientalis UA * EU341143 * M 100/100 Sphingomonas mali IFO10550 * NR_026374 91/100 Caulobacter leidyia ATCC15260 * NR_025324 98/98 Sphingomonas phyllosphaerae * AY563441 96/100 Howardella ureilytica * DQ925472 96/100 Lactovum miscens * AJ439543 98/100 11259 Biscogniauxia * J. deppeana MTL * AM906086 * B 98/11272 Phaeomoniella * J. deppeana IS * AM906086 * B 98/100 Paenibacillus graminis * AJ223987 97/Cohnella phaseoli * EU014872 96/100 11123 Pithya * J. virginiana UA * AJ717382 * G 99/99 11124 J. deppeana MTL * FJ542812 * H == 100/- Bacillus anthracis * AM747220 98/100 Bacillus cereus ATCC31293 * FJ601653 99/99 100/100 Bacillus mycoides * AF155956 11394 J. virginiana UA * EU071553 * J == 100/100 Bacillus licheniformis * AM913932 98/100 11180 J. osteosperma CHU * FJ514812 * A == 100/100 11035 Pestalotiopsis * J. deppeana MTL * FJ194961 * A 100/99 11219 C. arizonica MTL * NR_024931 * A == 100/99 11136 C. arizonica MTL * FJ549011 * A == 100/- 11164 Phaeomoniella * J. deppeana MTL * FJ549011 * A 11290 J. deppeana MTL * FJ549011 * A == 90/100 Desulfomicrobium orale * NR_025378 Desulfomicrobium norvegicum * NR_025407 86/100 Campylobacter helveticus * NR_025948 Campylobacter jejuni subsp. jejuni * CP000768 Bacteroides ovatus ATCC8483T * X83952 Bacteroides fragilis ATCC25285T * CR626927 94/100
88
Figure 3.
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Dendrographa minor Botryosphaeria ribis 9133 Platycladus orientalis NC * DQ984599 (Xanthomonadaceae) * B 469 Laetia thamnia - Panama 2834 Dryas integrifolia - Quebec 2779 Dryas integrifolia - Quebec 3300 Dryas integrifolia - Quebec 1182 Magnolia grandiflora - North Carolina 6730 Swartzia simplex - Panama 6731 Swartzia simplex - Panama * O 435b Laetia thamnia - Panama 9149b Platycladus orientalis UA * AF130967 (Enterobacteriaceae) * C 9130 Platycladus orientalis NC 9144 Platycladus orientalis NC 9135 Platycladus orientalis NC * DQ984599 (Xanthomonadaceae) * B 9134 Platycladus orientalis NC 9129 Platycladus orientalis NC Microxyphium citri Raciborskiomyces longisetosum 9128 Platycladus orientalis NC * EU236303 (Burkholderiaceae) * D 2712 Huperzia selago - Quebec 2722 Huperzia selago - Quebec 462 Laetia thamnia - Panama 4140 Dryas integrifolia - Quebec 3358A Dryas integrifolia - Quebec 4221 Dryas integrifolia - Quebec 3326 Picea mariana - Quebec 5007 Picea mariana - Quebec Sydowia polyspora 1029 Magnolia grandiflora - North Carolina DC2611 Juniperus scopulorum CHU * FJ422393 (Moraxellaceae) * H * 4109 Huperzia selago - Quebec Delphinella strobiligena 4947A Picea mariana - Quebec 4089 Picea mariana - Quebec Dothidea insculpa Dothidea sambuci Dothidea ribesia Stylodothis puccinioides 9096 Platycladus orientalis UA * DQ490310 (Oxalobacteraceae) * K Discosphaerina fagi 3275 Dryas integrifolia - Quebec 3335 Dryas integrifolia - Quebec 3353 Dryas integrifolia - Quebec 9042 Platycladus orientalis UA 9060 Platycladus orientalis UA * EU236303 (Burkholderiaceae) * D 9079 Platycladus orientalis UA 9084b Platycladus orientalis UA * EF509196 (Pasteurellaceae) * F 9002 Platycladus orientalis UA 9009b Platycladus orientalis UA 9028 Platycladus orientalis UA 9031 Platycladus orientalis UA 9036 Platycladus orientalis UA 3329 Picea mariana - Quebec 3377 Picea mariana - Quebec Lojkania enalia Trematosphaeria heterospora 5095 Picea mariana - Quebec Westerdykella cylindrica Preussia terricola 9106 Cupressus arizonica UA * DQ984599 (Xanthomonadaceae) * B 9116 Cupressus arizonica UA 9120 Cupressus arizonica UA * EU236303 (Burkholderiaceae) * D 9104 Cupressus arizonica UA 9051 Platycladus orientalis UA 5718 Dryas integrifolia - Quebec Byssothecium circinans 3323 Dryas integrifolia - Quebec Bimuria novae zelandiae Letendraea helminthicola 9203 Juniperus virginiana NC 9200 Juniperus virginiana NC 11251 Cupressus arizonica MTL 9149a Platycladus orientalis UA 9145 Platycladus orientalis NC * AF130967 (Enterobacteriaceae) *C 9140 Platycladus orientalis NC * FJ756346.1 (Enterobacteriaceae) * O 9137 Platycladus orientalis NC Phoma glomerata 9158 Platycladus orientalis NC 9054 Platycladus orientalis NC * EU341143 (Sphingomonadaceae) * J 9107 Cupressus arizonica UA * EU071476 (Pasteurellaceae) * F 9065 Platycladus orientalis UA 5654 Picea mariana - Quebec 9005 Cupressus arizonica UA 9008 Cupressus arizonica UA 9007 Platycladus orientalis UA 9018 Platycladus orientalis UA 9015 Platycladus orientalis UA 9027 Platycladus orientalis UA 9030 Platycladus orientalis UA Curvularia brachyspora Cochliobolus heterostrophus Setoshaeria monoceras Pyrenophora tritici repentis 9055 Platycladus orientalis UA * DQ984599 (Xanthomonadaceae) * B 9026 Platycladus orientalis UA * NR_026208 (Moraxellaceae) * I 9021 Platycladus orientalis UA 9009a Platycladus orientalis UA 9057 Cupressus arizonica UA 9059 Cupressus arizonica UA 9058 Cupressus arizonica UA Setomelanomma holmii Ampelomyces quisqualis Phaeosphaeria avenaria 2851 Huperzia selago - Quebec 2706 Huperzia selago - Quebec 5246 Huperzia selago - Quebec
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Figure 4.
Peltula umbilicata Arthroderma curreyi 100/100
Auxarthron zuffianum 100/-
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4466 Picea mariana * Quebec * O 100/100
11247 Juniperus deppeana MIN
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11272 J. deppeana IS * AM906086 (Paenibacillaceae) * A 11164 J. deppeana MTL* FJ549011 (Paenibacillaceae) * E
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Capronia mansonii 422 Laetia thamnia * Panama
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Figure 5.
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Leotia lubrica Lulworthia fucicola Lulworthia grandispora Thyridium vestutum Gaeumannomyces graminis var graminis 6702 Trichilia tuberculata - Panama Diaporthe phaseolorum Valsa ambiens Cryptodiaporthe corni Cryphonectria cubensis Cryphonectria havanensis Melanconis marginalis Discula destructive Gnomonia setacea Gnomoniella fraxini Plagiostoma euphorbiae Cephalotheca sulfurea Menispora tortuosa Camarops microspora Farrowia longicollea Farrowia seminude Chaetomium globosum Neurospora crassa Sordaria macrospora Sordaria fimicola 9038 Platycladus orientalis UA 9097 Cupressus arizonica UA * 9069 Platycladus orientalis UA 9093 Platycladus orientalis UA 9094 Platycladus orientalis UA * EU236303 (Burkholderiaceae) * D 9092 Platycladus orientalis UA 9085 Platycladus orientalis UA Oxydothis frondicola Microdochium nivale Hyponectria buxi 4653 Picea mariana - Quebec Arthrinium phaeospermum Apiospora sinensis 6722 Faramea occidentalis - Panama * O 9143 Platycladus orientalis NC * DQ984599 (Xanthomonadaceae) * B 9126 Platycladus orientalis NC * DQ984599 (Xanthomonadaceae) * N 9121 Platycladus orientalis NC 9089 Platycladus orientalis UA 11035 Juniperus deppeana SKY * FJ194961 (Paenibacillaceae) * E Cryptosphaeria eunomia Fasciatispora petrakii Cainia graminis Seynesia erumpens Hypoxylon fragiforme 256 Laetia thamnia - Panama 2204 Pinus taeda - NC 6717 Gustavia superba - Panama 11122 Cupressus arizonica SKY 9202 Juniperus virginiana NC 9201 Juniperus virginiana NC Astrocystis cocoes Rosellinia necatrix Xylaria acuta Xylaria hypoxylon 6728 Faramea occidentalis - Panama Halorosellinia oceanica Pleurothecium recurvatum 6756 Gustavia superba - Panama 6741 Theobroma cacao - Panama 6733 Swartzia simplex - Panama 412 Laetia thamnia - Panama 6749 Theobroma cacao - Panama 6714 Gustavia superba - Panama 151 Laetia thamnia - Panama Microascus trigonosporus Ascosalsum cincinnatulum Magnisphaera stevemossago Sagaaromyces abonnis Corollospora maritima Varicosporina ramulosa Ascosacculus heteroguttulatus Natantispora lotica Halosarpheia marina 425 Laetia thamnia - Panama Torrubiella luteorostrata 416 Laetia thamnia - Panama 6708 Gustavia superba - Panama 4939 Picea marina - Quebec Balansia henningsiana Hydropisphaera erubescens Cordyceps khaoyaiensis 4780 Dryas integrifolia - Iqaluit Emericellopsis terricola 9154 Platycladus orientalis NC 9168 Platycladus orientalis NC Hypocrea citrina Verticillium epiphytum Hypomyces polyporinus Cordyceps irangiensis Cordyceps sinensis
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Appendix 1A. GenBank accession numbers for LSU rDNA sequences of endophytic fungi for phylogenetic analyses. All endophyte sequences were obtained and published by Higgins et al., 2007.
Endophyte isolate
Accession no.
151 256 412 416 422 425 435b 462 469 1029 1182 2204 2706 2712 2722 2779 2834 2851 3275 3300 3323 3326 3329 3335 3353 3358A 3377 4089 4109 4140 4221 4466 4653 4780 4939 4947A 5007 5095 5246
EF420086 EF420088 EF420089 EF420090 EF420091 EF420092 EF420093 EF420094 EF420095 EF420084 EF420085 EF420087 DQ979418 DQ979419 DQ979420 DQ979422 DQ979423 DQ979424 DQ979425 DQ979427 DQ979428 DQ979429 DQ979430 DQ979431 DQ979432 DQ979433 DQ979434 DQ979437 DQ979438 DQ979440 DQ979441 DQ979444 DQ979449 DQ979452 DQ979453 DQ979454 DQ979455 DQ979457 DQ979458
93
5654 5718 6702 6708 6714 6717 6722 6728 6730 6731 6733 6741 6749 6756
DQ979462 DQ979463 EF420096 EF420097 EF420098 EF420099 EF420100 EF420101 EF420102 EF420103 EF420104 EF420105 EF420106 EF420107
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Appendix 1B. Representative Ascomycota species used in phylogenetic analyses with GenBank identification numbers. Sequences for Coccidioides posadasii were obtained from TIGR (http://www.tigr.org).
Taxon Ampelomyces quisqualis Aphanoascus fulvescens Apiospora sinensis Arthrinium phaeospermum Arthroderma curreyi Ascosacculus heteroguttulatus Ascosalsum cincinnatulum Ascosphaera apis Astrocystis cocoes Auxarthron zuffianum Balansia henningsiana Bimuria novae-zelandiae Botryosphaeria ribis Byssochlamys nivea Byssothecium circinans Cainia graminis Camarops microspora Capronia mansonii Capronia pilosella Cephalotheca sulfurea Ceramothyrium carniolicum Chaetomium globosum Coccidioides posadasii Cochliobolus heterostrophus Cordyceps irangiensis Cordyceps khaoyaiensis Cordyceps sinensis Corollospora maritima Cryphonectria cubensis Cryphonectria havanensis Cryptodiaporthe corni Cryptosphaeria eunomia var. eunomia Curvularia brachyspora Delphinella strobiligena Dendrographa leucophaea f. minor Dermatocarpon luridum Diaporthe phaseolorum Discosphaerina fagi Discula destructiva Dothidea insculpta Dothidea ribesia Dothidea sambuci Emericellopsis terricola Eremascus albus Exophiala jeanselmei
Accession no. 31415592 33113349 27447246 27447247 33113367 34453082 34453080 11120650 27447238 33113353 45155172 15808399 11120642 33113391 15808400 20334356 27447236 11120644 12025061 20334357 11120645 13469777 TIGR222929 45775574 13241779 13241765 29466997 22203655 22671402 22671403 22671407 27447241 12025063 15808401 47499217 52699696 1399144 15808402 22671422 52699697 15808403 45775610 1698507 11120651 4206347
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Fasciatispora petrakii Farrowia longicollea Farrowia seminuda Gaeumannomyces graminis var. graminis Glyphium elatum Gnomonia setacea Gnomoniella fraxini Halorosellinia oceanica Halosarpheia marina Hydropisphaera erubescens Hypocrea citrina Hypomyces polyporinus Hyponectria buxi Hypoxylon fragiforme Leotia lubrica Letendraea helminthicola Lojkania enalia Lulworthia fucicola Lulworthia grandispora Magnisphaera stevemossago Melanconis marginalis Menispora tortuosa Microascus trigonosporus Microdochium nivale Microxyphium citri Natantispora lotica Neurospora crassa Oxydothis frondicola Paracoccidioides brasiliensis Peltula umbilicata Penicillium freii Phaeosphaeria avenaria Phoma glomerata Plagiostoma euphorbiae Pleurothecium recurvatum Preussia terricola Pyrenophora tritici-repentis Pyrenula cruenta Pyrenula pseudobufonia Raciborskiomyces longisetosum Rosellinia necatrix Sagaaromyces abonnis Setomelanomma holmii Setosphaeria monoceras Seynesia erumpens Sordaria fimicola Sordaria macrospora Stylodothis puccinioides Sydowia polyspora Thyridium vestitum Torrubiella luteorostrata Trematosphaeria heterospora Varicosporina ramulosa
27447243 13469782 13469784 16565889 17104831 16565895 16565884 27447237 34453085 45155165 45775578 32261014 27447249 27447244 45775573 15808405 15808406 22203665 22203666 34453095 22671436 45775611 1399149 2565289 11120643 34453084 13469785 27447250 2271524 15216700 52699706 45775613 31415593 22671445 45775614 45775615 45775601 12025090 52699710 15808410 27447239 34453078 28144315 15808411 12025093 45155203 37992293 11120648 45775604 45775600 13241778 15808412 22203671
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Verrucaria pachyderma Verticillium epiphytum Valsa ambiens Westerdykella cylindrica Xylaria acuta Xylaria hypoxylon
15216679 8777749 16565896 11120649 45775605 45775577
Appendix 2. Details of phylogenetic analyses for three classes of fungi.
Parsimony analyses
Bayesian analyses
Class
Sequences (this study/ named taxa)
Included characters (total)
Length of shortest trees
Trees excluded as burn-in
Trees in majority rule consensus
Dothideomycetes
107 (49/58)
1208 (4595)
761
3585
5415
Eurotiomycetes
25 (4/21)
1056 (4443)
418
3000
3400
Sordariomycetes
95 (17/78)
1218 (4605)
1313
5550
8890
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Appendix 3. Representative species of bacteria used in phylogenetic analyses with GenBank identification numbers.
Taxon
Bacillus anthracis Bacillus cereus sp. Bacillus sp. L164 Bacillus mycoides Bacteroides fragilis Bacteroides ovatus ATCC 8483T Burkholderia cepacia ATCC 25416T Burkholderia fungorum Burkholderia pseudomallei ATCC 23343 Burkholderia rhizoxinica Burkholderia sordicola Burkholderia sp 20577 Burkholderia sp 30887 Burkholderia sp 69968 Campylobacter helveticus Campylobacter jejuni subsp jejuni Candidatus Glomeribacter gigasporarum Candidatus Glomeribacter gigasporarum Candidatus Glomeribacter gigasporarum Candidatus Glomeribacter gigasporarum Caulobacter leidyia strain ATCC 15260 Cohnella phaseoli Desulfomicrobium norvegicum Desulfomicrobium orale Erwinia amylovora ATCC15580 Escherichia coli ATCC11775T Howardella ureilytica strain Lactovum miscens Nitrosomonas sp. BF16c52 Paenibacillus graminis Pantoea agglomerans ATCC27155 Pseudomonas syringae Ralstonia mannitolilytica Ralstonia solanacearum ATCC 11696 Sphingomonas mali IFO10550 Sphingomonas phyllosphaerae Xanthomonas campestris pv. campestris Xanthomonas oryzae strain LMG 5047
GenBank Accession
AM747220 FJ601653 AM913932 AF155956 CR626927 X83952 AB334766 AF215705 CP000573 AJ938142.1 AF215705 AJ938141.1 AJ938143.1 AJ938144.1 NR_025948 CP000768 X897272 AJ251633.1 AJ251634.1 AJ251635.1 NR_025324 EU014872 NR_025407 NR_025378 U80195 NR_024570 DQ925472 AJ439543 AF386747 AJ223987 FJ357815 AJ308316 AJ270258 EF016361 NR_026374 AY563441 AM920689 NR_026319
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Appendix 4. Percentages of taxonomic orders of Pezizomycotina obtained in each biogeographic region based on FMP curated ITS database results. Remaining taxa not shown here include mitosporic fungi, fungi of uncertain placement, unidentifiable fungi, and a small number of isolates from lineages other than the Pezizomycotina, such that columns may not sum to 100%. Percentage of isolates by taxonomic orders
SKY
MOG
CHU
UA
DUKE
Botryosphaeriales Capnodiales Chaetothyriales Coniochaetales Diaporthales Dothideales Eurotiales Halosphaeriales Helotiales Hypocreales Myriangiales Pezizales Pleosporales Sordariales Xylariales
11.1% 11.1% 0 0.7% 0.7% 7.4% 0 0 5.9% 0 0 25.2% 5.2% 0.7% 18.5%
9.9% 1.4% 1.4% 4.2% 0 8.5% 0 0 5.6% 0 0 21.1% 11.3% 1.4% 12.7%
1.6% 1.6% 0 3.2% 3.2% 4.8% 9.7% 3.2% 12.9% 3.2% 0 27.4% 8.1% 0 8.1%
0 11.3% 1.6% 6.5% 0 12.9% 0 3.2% 0 0 0 4.8% 35.5% 3.2% 1.6%
44.0% 1.2% 0 0 2.4% 0 1.2% 0 2.4% 4.8% 1.2% 1.2% 4.8% 0 17.9%
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APPENDIX C
IAA PRODUCTION BY ENDOPHYTIC PESTALOTIOPSIS NEGLECTA IS ENHANCED BY AN ENDOHYPHAL BACTERIUM
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IAA production by endophytic Pestalotiopsis neglecta is enhanced by an endohyphal bacterium Michele T. Hoffman1, Malkanthi K. Gunatilaka1, E. M. Kithsiri Wijeratne2, A. A. Leslie Gunatilaka2 and A. Elizabeth Arnold1
1. College of Agriculture and Life Sciences Division of Plant Pathology and Microbiology School of Plant Sciences 1140 E. South Campus Drive, Forbes 303 University of Arizona Tucson, AZ 85721 USA
2. Southwest Center for Natural Products Research and Commercialization Office of Arid Lands Studies 250 E. Valencia Road University of Arizona Tucson, Arizona 85706 USA For correspondence: AE Arnold, Tel: 520.621.7212; Fax: 520.621.9290; Email:
[email protected]
Running head: Endohyphal bacterium enhances IAA production by fungal endophyte Contains 5 figures, and 2 appendices
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Abstract A variety of plant-associated microbes produce phytohormones with pervasive effects on plant growth. Specifically, indole-3-acetic acid (IAA) production has been recorded among diverse plant pathogens, mycorrhizal and rhizosphere endophytes, and in endophytic bacteria and yeasts from healthy, above-ground plant tissues. Here, we show for the first time that a filamentous foliar endophyte, Pestalotiopsis neglecta, produces IAA, and that its production is significantly enhanced when the endophyte hosts an endohyphal bacterium (Luteibacter sp.). When cultivated axenically, the bacterium does not produce IAA in culture. Both P. neglecta and the P. neglecta/Luteibacter complex appear to rely on L-tryptophan for IAA synthesis. Culture filtrate from the endophytebacterium complex significantly increased root growth in vitro in seedlings of tomato relative both to controls and to filtrate from the endophyte alone. Our results reveal that this endophyte, recovered from a coniferous host (Platycladus orientalis), has the potential to strongly influence growth of a distantly related plant, demonstrate the previously unknown role of an endohyphal bacterium in enhancing phytohormone production by a plant-symbiotic fungus, and provide a new lens with which to view the considerable genotypic, species-level, and functional diversity of foliar endophytic fungi. Keywords: bacterial endosymbionts, IAA, auxins, Pestalotiopsis, Luteibacter, Xanthomonadales, fungal endophytes, phylogeny
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Introduction Benefits conferred to plants by mycorrhizal fungi, including enhanced uptake of nutrient and water, are well-documented and generally represent an intuitively straightforward alignment of plant- and fungal fitness (Parniske, 2008; Bonfante and Anca, 2009; Kobayashi and Crouch, 2009; Hoeksema et al., 2010). In contrast, the potential contributions of foliar endophytes to plant physiology are both less obvious and less thoroughly studied. Such contributions are especially difficult to resolve in the case of endophytes associated with woody plants, which frequently comprise an exceptional richness of highly localized infections at the scale of only a few millimeters of leaf area (Carroll and Petrini, 1983; Lodge et al., 1996; Arnold et al., 2000; Arnold, 2007). These Class 3 endophytes – a term used to refer to the species-rich, phylogenetically diverse, horizontally transmitted, tissue-specific, and localized endophytes known from all major lineages of land plants (Rodriguez et al., 2009) – are a fundamental feature of plant biology, occurring in plants from the tundra to tropical rainforests (Higgins et al., 2007; Arnold and Lutzoni, 2007). The benefits or costs they extend to their hosts, the nature of their interactions with host tissues on the intimate scales of cells and molecules, and the mechanisms by which they escape, tolerate, or prevent induction of host defenses represent three major questions in the study of these ubiquitous plant symbionts (Arnold et al., 2003; Waller et al., 2005; Arnold and Engelbrecht, 2007). Over recent decades, a large number of studies have documented the ability of diverse microbes to synthesize phytohormones such as gibberellins, ethylene, cytokinins, jasmonic acid, abscisic acid, and especially indole-3-acetic acid (IAA), often with profound effects on plant growth, tissue differentiation, and reproduction (Costacurta and
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Vanderleyden, 1995; Glick, 1995; Lindow et al., 1998; Robinson et al., 1998; Koga, 1995; Maor et al., 2004; Sirrenberg et al., 2007; Spaepen et al., 2007). Most of these studies have focused on plant-pathogenic microbes, mycorrhizal fungi, rhizosphere endophytes, and endophytic bacteria and yeasts from above-ground tissues (Gruin, 1959; Sosa-Morales et al., 1997; Nassar et al., 2005; van der Lelie et al., 2009; Chagué et al., 2009; Xin et al., 2009). Together, they reveal that microbial production of IAA is phylogenetically widespread, encompassing both bacteria (e.g., Erwinia herbicola; Yang et al., 2006; Pantoea agglomerans; Barash and Manulis-Sasson, 2009) and diverse lineages of fungi (e.g., Mucoromycotina, Basidiomycota, and some Ascomycota; Strack et al., 2003; Bajo et al., 2008; Reineke et al., 2008). In many cases, IAA production by plant-associated microbes strongly influences plant physiology in ways that are relevant to plant-microbe interactions. For example, production of IAA by some mycorrhizal fungi stimulates production of plant biomass and promotes disease resistance (Bonfante and Anca, 2009). Sirrenberg et al. (2007) noted an increase in Arabidopsis root growth following exposure to diffusible IAA in filtrate from liquid cultures of the rhizosphere fungus Piriformospora indica (Sebacinales, Basidiomycota). Microbial IAA production can inhibit plant hypersensitive responses, limiting the production of plant-defense compounds such as chitinase and glucanase enzymes (Tudzynski and Sharon, 2002). Here, we show for the first time that Pestalotiopsis neglecta, a filamentous foliar endophyte recovered from Platycladus orientalis (Cupressaceae), produces IAA in vitro, and that its production is enhanced significantly when the endophyte hosts an endohyphal bacterium (Luteibacter sp.). Culture filtrate from the endophyte-bacterium complex
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significantly increased root growth in seedlings of a distantly related plant (tomato) relative both to controls and to filtrate from the endophyte alone. Our results highlight the previously unknown role of an endohyphal bacterium in enhancing phytohormone production by a plant-symbiotic fungus and provide a new lens with which to view the remarkable genotypic, species-level, and functional diversity of foliar endophytic fungi.
Methods Identification of fungal endophyte 9143 As part of a larger study, an endophytic fungus (9143) was collected from surfacesterilized foliage of a mature, healthy individual of Platycladus orientalis (Cupressaceae) in Durham, NC (Hoffman and Arnold, 2008). The isolate was archived as a living voucher in sterile water at the Robert L. Gilbertson Mycological Herbarium (ARIZ) at the University of Arizona. Previous phylogenetic analyses (Hoffman and Arnold, 2010) confirmed ordinal placement of this isolate (Xylariales) but were insufficient to identify it to species. After observing its bacterial endosymbiont and IAA production (below), we identified the isolate on the basis of conidial morphology (Steyaert, 1953) and then confirmed our designation using phylogenetic analyses. For morphological characterization, we examined conidia of cultures grown on 2% malt extract agar media for 7 days (Appendix 1). For phylogenetic analyses, we examined ITS rDNA sequence data for 9143 (obtained previously; see Hoffman and Arnold, 2008) in the context of 59 publicly available ITS rDNA sequences for Pestalotiopsis spp. (Jeewon et al, 2003, 2004; Wei et al., 2007; Watanabe et al, 2010). We manually aligned these 60 sequences using Mesquite version 1.06 (Maddison and
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Maddison, 2005). Taxon sampling included the breadth of close relatives of P. neglecta, representing our estimation of taxonomic placement based on morphology. Phylogenetic relationships were inferred using parsimony and Bayesian Metropolis-coupled Markov chain Monte Carlo (MCMCMC) methods. For maximum parsimony (MP), a heuristic search with random stepwise addition and tree bisectionreconnection (TBR) was implemented in PAUP* 4b10 (Swofford, 2002). Support was assessed using parsimony bootstrap (1000 replicates). Bayesian MCMCMC analysis was implemented in MrBayes v. 3.1.2 (Huelsenbeck and Ronquist; 2001) for 2 sets of 3 million generations each, initiated with random trees, four chains, and sampling every 1000th tree, using GTR+I+G based on sequence evaluation in Modeltest 3.7 (Posada and Crandall, 1998). After elimination of the first 500 trees as burn-in, the remaining 5002 trees were used to infer a majority rule consensus. Sequence data for this isolate are deposited in GenBank (ITS data used for species-level identification, as well as LSUrDNA data from Hoffman and Arnold, 2010; EF419899.1, EF420070.1).
Identification of endohyphal bacterium Using light microscopy and PCR with 16S rDNA primers, we determined that endophyte 9143 harbored an endohyphal bacterium, which we identified previously only as a member of the Gammaproteobacteria (Hoffman and Arnold, 2010). To place this bacterium at the species level, we sampled more densely within the phylum, with a special focus on Xanthomonaceae, using 36 16S rDNA sequences obtained by BLAST matches to our 16S rDNA sequence (obtained using 10F/1507R primers; HM117737). Manual alignment was conducted in Mesquite v. 1.06 (Maddison and Maddison, 2005)
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and the resulting data set analyzed using parsimony and Bayesian MCMCMC methods. Relationships were inferred using Felsenstein non-parametric distance methods, saving 10 best trees in each replicate set of 100 in PAUP* 4b10 (Swofford, 2002). Using maximum parsimony (MP), a heuristic search with random stepwise addition and tree bisection-reconnection (TBR) was implemented in PAUP*. Support was then assessed using parsimony bootstrap (1000 replicates). Bayesian analysis was implemented in MrBayes v. 3.1.2 (Huelsenbeck and Ronquist, 2001) on the CIPRES teragrid portal (Miller et al., 2009; http://www.phylo.org/sub_sections/portal) for 2 runs of 10 million generations, initiated with random trees, four chains, and sampling every 1000th tree, using GTR+I+G based on evaluation in Modeltest 3.7 (Posada and Crandall, 1998). After elimination of the first 440 trees as burn-in, the remaining 19,122 trees were used to infer a majority rule consensus.
Production of bacterium-free strain To eliminate the bacterium from the endophyte, we grew isolate 9143 on 2% MEA amended with 40µg/ml of antibiotic ciprofloxacin. Infection status (infected, cured) was confirmed using light microscopy (400X), which ruled out external contaminants, followed by DNA extraction and 16S rDNA PCR (Hoffman and Arnold, 2010). Infected and cured cultures of 9143 were stored at -80°C in 80% glycerol. We found no difference between infected and cured isolates of 9143 in terms of growth rate on 2% MEA or water agar at 22°C, nor with regard to pH of the growth medium (Appendix 2).
Measurement of indole compound production in vitro
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Small samples of mycelium from infected and cured isolates were plated on 2% MEA and allowed to grow for 7 d. Small pieces were removed from the resulting cultures using a 0.5mm cork borer and used to inoculate three sterile flasks containing 80ml of Czapek Dox broth (CDB; Hymedia; pH 7.2) augmented with L-tryptophan (5mM). For each treatment set, one additional flask containing only CDB was used as a negative control. All flasks were checked for contamination (turbidity) after agitating at 120rpm at 26°C for 24 h. Contaminated cultures were discarded and replaced. After 72 hours of agitation, three 1ml aliquots of broth were removed from each flask and treated with colorimetric Salkowski reagent (0.01 M Fe (NO3)3; 7.0 M HCIO4) prior to standard spectophotometric measurements at 530nm (Salkowski, 1885; Ehmann, 1977; Glickmann and Dessaux, 1995). Each 1ml extract was combined with 500µl of Salkowski reagent, incubated at room temperature for 30 minutes, and then measured once on a Whatman spectrophotometer (530nm). CDB was used as a blank between readings. The mean of the three readings was calculated for each culture, concentrations inferred using an IAA standard curve (below), and analyzed with a t-test in JMP® 8.0 (SAS Institute Inc., Cary, NC.). The IAA standard curve was inferred using commercial IAA (MW 175.19, Sigma; 1mM), prepared in a volumetric flask using 95% molecular grade methanol. Fourteen 1.5ml Eppendorf™ tubes were prepared in proportional ratios with 25-1950µl of IAA and 50-1975µl of reagent. After incubating in the dark for 30 minutes, each sample was measured twice. A blank calibration control using 1ml water plus 500µl of reagent was as a blank between readings.
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Identification of the indole compound as IAA using chromatography We used thin layer chromatography (TLC) and high performance liquid chromatography (HPLC) to identify the indole compound that was observed on the basis of colorimetric reaction with Salkowski reagent. For chromatographic analysis, two cultures each of infected and cured mycelia were grown for two weeks in 500mL of sterile CDB supplemented with 5mM L-tryptophan. One additional culture (9143-infected) was grown in sterile CDB without L-tryptophan (1.0 L) to determine whether fungus 9143 uses a tryptophan-dependent pathway for IAA production. Each culture was filtered using Whatman No.1 filter paper and the filtrate (1.0 L) extracted three times with 500mL of ethyl acetate (EtOAc). The combined EtOAc layer was washed three times with 500mL of H2O, dried over anhydrous Na2SO4, and evaporated under reduced pressure to yield the EtOAc extract. Normal phase TLC analysis of the EtOAc extract was performed on aluminumbacked plates coated with 0.20 mm layer of silica gel 60 F254 (E. Merck, Darmstadt). Spots were visualized by inspection of plates under UV light (254 nm) and after spraying with Van Urk-Salkowski reagent (Ehmann, 1977) followed by heating. Commercially available IAA (Sigma) was used as an authentic sample (Eluant: 8% MeOH in dichloromethane, Rf 0.3). Analytical reversed phase TLC investigations were performed on aluminum-backed plates coated with 0.20 mm layer of silica gel 60 RP-18 F254S (E. Merck, Darmstadt; Eluant: 20% MeCN in H2O, Rf 0.5). Analytical HPLC analysis of the EtOAc extract was performed using a Kromasil 5µm C-18 column (4.6 x 250 mm) on a Shimadzu HPLC system equipped with DGU14A degasser, LC-10ADvp pump, SPD-M10Avp diode array detector and SCL-10Avp
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system controller utilizing Shimadzu LC-MS solution software. Samples were redissolved in MeOH (2.0 mg/mL) and injections (10µL) were made with Shimadzu SIL10ADvp auto injector. The mobile phase consisted of H2O/ MeCN/HCOOH (69.75:30:0.25) with a flow rate of 1.2 mL min-1.
Seedling assays Cured and infected isolates of 9143 were agitated for 15 days in 200ml CDB + 5µM Ltryptophan. Cultures then were vacuum-filtered (0.44 mm nylon filter) and the liquid retained in sterile 50ml tubes. The pH was measured from an aliquot of each (pH 5.7 for 9143-infected; pH 4.2 for 9143-cured; pH 7.1 for CDB alone; Denver Instruments pH meter) and the solutions brought to pH 7.0-7.1, as needed, by amendment with 0.5M NaOH. Approximately 25ml of this filtrate was further filter-sterilized using 0.2mm syringe filters into a sterile container and used in seedling assays. Tomato seeds (ACE 55; The Home Depot®) were surface-sterilized by agitating in 50% bleach for 12-15 minutes, rinsed in sterile water, and placed on sterile filter paper in sterile 60mm Petri dishes (ca. 20 seeds/dish). Three milliliters of sterile water was added to each dish before sealing with Parafilm®. Seeds were allowed to germinate at 25°C for 5 days. Sets of 10 apparently healthy seedlings were chosen haphazardly and transferred under sterile conditions to new 60mm plates containing sterile filter paper. Each set of seedlings was watered with 3.5ml of sterile water. Four plates/treatment (40 seedlings/treatment) then received 50µl per seedling of one of five treatments: (a) CDB from 9143-infected; (b) CDB from 9143-cured; (c) CDB + 5µM L-tryptophan; (d) CDB + IAA (0.1mg/ml; pH 7.0); or (e) sterile water.
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Plates were sealed with Parafilm® and incubated under fluorescent lights at room temperature for 5 d (12 h light/dark cycle). Seedlings then were harvested and stretched to full length by mounting to paper with transparent cellophane tape. Shoot and root lengths to longest points were measured for each seedling using calipers. Data were analyzed in JMP® 8.0 using ANOVA after normalizing all measurements to the results of the CDB + IAA treatment (treatment d).
Isolation of the endohyphal bacterium Incubating the isolate 9143 on 2% MEA at 36°C for 7 d (Appendix 1) resulted in emergence of bacterial growth from the apparently axenic endophyte culture. We extracted DNA from the bacterium following Hoffman and Arnold (2010) and sequenced an 1100bp portion of the 16SrDNA region of using primers 27F /1429R (Lane 1991). The sequence of this bacterial isolate (BAC182) was identical to that of the bacterium sequenced directly from endophyte 9143. BAC182 was grown overnight in sterile LB broth and vouchered in sterile glycerol at -80°C. Using the above methods we found no evidence of IAA production by axenic cultures of this bacterium (data not shown).
Results Phylogenetic analyses were used in conjunction with morphology to positively identify endophyte 9143 as Pestalotiopsis neglecta (Fig. 1, Appendix 1). Multisetulate conidia are fusiform shaped concolorous cells, lacking knobbed appendages. Phylogenetic analysis placed this fungal isolate as sister to one of three identified P. neglecta strains from Watanabe et al. (2010). Our results are congruent with their analysis, placing two other putative “P. neglecta” sequences in a separate subclade. Reclassification of the
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taxonomic placement of Pestalotiopsis spp. has been recommended (see Jeewon et al., 2004; Hawksworth 2005; Wei et al., 2007), with greater emphasis placed on morphological characteristics combined with rigorous phylogenetic methods. Phylogenetic analyses of 16S rDNA confirmed placement of fungal endophyte 9143 endohyphal bacterium in the Xanthomonadaceae, with strong support as a member of the genus Luteibacter (Luteibacter sp.; Fig. 2). Chromogenic testing indicated that isolate 9143 produced an indole compound in vitro, with enhanced production in the presence of the endohyphal bacterium (Fig. 3). At the end of the 14 d cultivation period, mean concentration of the indole compound in broth from 9143-infected (104.8µg/ml) was 78µg/ml greater than that produced by 9143cured (Fig. 3). Strong differences in indole compound concentration were observed throughout the 14 d cultivation period (repeated measures ANOVA, F1, 4 = 358.7; p < 0.0001; Fig. 3). TLC and HPLC identified the compound as indole-3-acetic acid (IAA) and confirmed that endophyte 9143 produced significantly more IAA when the endohyphal bacterium was present vs. absent (Fig. 4). No IAA was produced when 9143-infected and cured isolates were grown in CDB without tryptophan, suggesting a tryptophandependent pathway for IAA production (data not shown). Growth of tomato seedlings, measured as total seedling length, shoot length, root length, and root:shoot ratio, differed significantly as a function of treatment (Fig. 5). Seedlings treated with filtrate from 9143-cured did not differ in any of these measures relative to the CDB control (Fig. 5, post-hoc Tukey-Kramer test, alpha = 0.05). Seedling
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length was significantly greater following treatment with 9143-infected, reflecting a significant enhancement of root elongation (Fig. 5, Tukey-Kramer tests, alpha = 0.05).
Discussion Endohyphal bacteria have been found previously in living hyphae of plant-associated Glomeromycota, Mucoromycotina, and several ectomycorrhizal Dikarya (Tuber borchii; Ascomycota; Laccaria bicolor and Piriformospora indica; Basidiomycota) (Bianciotto et al., 1996; Barbieri et al., 2000; Bertaux et al., 2003; Levy et al., 2003; Partida-Martinez and Hertweck, 2005). Recently, we reported their presence in foliar endophytic fungi representing four classes of Pezizomycotina, and demonstrated that they are both geographically widespread and phylogenetically diverse (Hoffman and Arnold, 2010). The present study is the first to (1) definitively identify the partners in a close ecological relationship between a foliar endophyte and an endohyphal bacterium, (2) show that a species of Pestalotiopsis (P. neglecta) is capable of producing a phytohormone, (3) demonstrate that a filamentous foliar endophyte can produce indole-3-acetic-acid (IAA), (4) show that IAA produced by this endophyte is dependent on the L-tryptophan pathway, and (5) demonstrate that an endohyphal bacterium (Luteibacter sp.) enhances this IAA production, with exogenous culture filtrate of the endophyte with its endohyphal symbiont yielding significantly enhanced growth in roots of a model plant. Together, our results reveal that products of an endophyte from a coniferous host has the potential to strongly influence growth of a distantly related plant (see also Rodriguez et al., 2009), demonstrate the previously unknown role of an endohyphal bacterium in altering phytohormone production by a plant-symbiotic fungus, and provide a new lens with
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which to view the considerable genotypic, species-level, and functional diversity of foliar endophytic fungi. In the presence of endohyphal Luteibacter sp. and L-tryptophan, P. neglecta cultivated in CDB produced ca. 100µg/ml of IAA. This amount is likely biologically significant in the host, as demonstrated both by our seedling assays and by previous studies with plant-pathogenic and rhizosphere fungi. IAA production observed here was within the range produced by Ustilago maydis mutant strains (75-262µg/ml) as measured using the same colorimetric reagents (Sosa-Morales et al., 1997). Using more precise methods (GC-MS and HPLC-ESI_MS/MS), culture filtrate from Piriformospora indica measured ca. 0.16µmol of IAA after 4.5 weeks of growth (Sirrenberg et al., 2007), exceeding that demonstrated here. Isolates of Colletotrichum were found to generate 232µg/ml of IAA using TLC and GC-MS (Robinson et al., 1998; Chagué et al., 2009). Although our study does not yet address bacterial-fungal-plant interactions during the endophyte symbiosis per se, it provides a first estimation of one way in which an apparently facultative bacterial endosymbiosis can influence interactions between an endophytic fungus and its host. Further experiments involving the establishment of this endophytic fungus within host tissues, and exploration of relative benefits for hosts more closely related to that from which the fungus was first isolated, represent important future directions. The strong response we observed in root growth as a function of filtrate from cultures containing both the endophyte and its endohyphal symbionts suggests that we should evaluate the potential for this foliar endophyte to colonize root tissues, and/or examine how fungi localized in foliage might influence growth in other tissues. We also are interested to discover whether IAA production by the endophyte or
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endophyte/endohyphal bacteria complex camouflages the fungus within the host, making it less detectable by plants and thus enhancing its ability to grow without inducing a hypersensitive response. Recent work has shown that even closely related or conspecific fungi can harbor different endohyphal bacteria (Hoffman and Arnold, 2010). These facultative associations are by definition flexible and appear to be gained and lost readily (Hoffman and Arnold, 2010). Our study provides the first evidence that the differences in bacterial associates of endophytes might play a key role in determining the nature of endophytehost associations, and may account for some of the diversity and ecological plasticity observed in endophyte-plant interactions (Arnold et al., 2009). More generally, increasing but still limited exploration of ectohyphal bacteria, endohyphal bacteria, and mycoviruses have begun to illustrate the powerful but often overlooked ways in which microbes associated with fungal hyphae can influence the outcome of plant-fungus interactions (Marquez et al., 2007, Bonfante and Anca, 2009; Kobayashi and Crouch, 2009). Advancing our knowledge of endophyte interactions with plant hosts, other extrinsic microorganisms, and most recently, diverse endohyphal bacteria, will help define the functional biology of these diverse and ubiquitous symbionts.
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Acknowledgments We thank the Division of Plant Pathology and Microbiology, the Department of Plant Sciences, the College of Agriculture and Life Sciences at the University of Arizona. Additional support from the National Science Foundation (NSF-0626520 and 0702825 to AEA; NSF-IGERT to MTH) is gratefully acknowledged. We thank D.R. Maddison for sharing pre-release versions of Mesquite and Chromaseq; R. Ryan, M. J. Epps, J. U’Ren, M. del Olmo Ruiz and A. Woodard for lab assistance and helpful discussion. We are especially grateful to H. VanEtten for access to chemistry facilities and J. L. Bronstein, L. S. Pierson, III, and M. J. Orbach for comments on the manuscript. This paper represents a portion of the doctoral dissertation research of MTH in Plant Pathology and Microbiology at the University of Arizona.
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Figure legends
Figure 1. Majority rule consensus tree based on Bayesian analyses of 60 ITSrDNA sequences representing Pestalotiopsis (with Seiridium as outgroup). Support value before the slash represents maximum-parsimony bootstrap (>70%); number after the slash represents Bayesian branch support > 90%.
Figure 2. Majority rule consensus tree (using P. syringae as outgroup) based on Bayesian analyses of 37 proteobacterial 16S rDNA sequences with affinity for Luteibacter spp. in BLAST analyses. Support value before the slash represents maximum-parsimony bootstrap >70%, number after the slash represents Bayesian branch support > 90. Branches in bold indicate neighbor-joining bootstrap support >70%.
Figure 3. Spectrophotometric absorbances at 530nm following treatment of supernatant from CDB cultures of 9143-infected and 9143-cured with Salkowski reagent, indicating production of an indole compound (identified as IAA; Fig. 4). Each point represents the average of three readings (two-tailed t4 = 20.99; p < 0.0001), converted to concentration amounts using the IAA standard curve (y =0.0108x-0.0049). Error bars indicate standard deviation.
Figure 4. HPLC analysis (retention time and co-injection) results for (A) IAA standard, and extracts from (B) 9143-cured and (C) 9143-infected. Elution time was 12.54 minutes for the IAA standard, 12.89 for 9143-cured, and 12.95 by 9143-infected sample. , IAA
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comprised 16% of the total sample from 9143-cured, and 29% from 9143-infected (Fig. 2C) based on the area ratio under the peak.
Figure 5. Growth responses of tomato seedlings to pH-consistent supernatant from 9143infected cultures in CDB; supernatant from 9143-cured in CDB; and control treatments (CDB alone, water). (A) Total seedling length, (B) root length, (C) root to shoot ratio, (D) shoot length. Data were normalized to measurements for plants that received CDB + 0.1mg/ml IAA treatment (data not shown). Different superscripts within each panel indicate statistically different means.
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Figure 1.
126
Figure 2.
127
Figure 3.
9143+ 9143-
IAA concentration (µg/ml)
Days
128
Figure 4.
(A) Indole-3-acetic acid standard
(B) 9143-cured
(C) 9143-infected
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Figure 5.
A. a
B.
F3,156 = 30.2; p< 0.0001
F3,156 = 32.8; p< 0.0001
a b
b
b
b
c
9143+
9143-
CDB
c
H2 O
9143+
9143-
CDB
H2 O
Seedling length (mm)
C.
D. F3,156 = 3.2; p< 0.0001
a ab
F3,156 = 19.3; p< 0.0001
a
ab
ab b
c
9143+
9143-
CDB
b
H2 O
Treatments
9143+
9143-
CDB
H2 O
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Appendix I
Pestalotiopsis spore morphology Pestalotiopsis neglecta asexual spore morphology description coincides with the morphology of conidia from endophyte 9143 (Fig A). These spores are fusiform, four-septate, a fuliginous brown in color, with end cells hyaline. The apical end is short with two or three spreading setulae, approximately 22um long. The basal end contains a pedicel about 4-7um long (Steyaert, 1953).
Figure A. Conidia from 9143-uninfected culture (400X) showing fusiform cells with 4 septae.
Figure B. Results of growth assays over 14 days for 9143-infected (black diamond) and 9143-cured (open square) on 2% MEA at pH4.5 (panel A), pH8.0 (panel B), and pH=6.8 (standard 2% MEA;
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panel C) at 22°C. Panel D shows growth on water agar at 22°C. 9143 did not grow at 36°C. Error bars indicate standard error of the three replicates performed for all experiments.
A.
B.
C.
D.
