DIVERSITY OF SOUTH AMERICAN AMBROSIA BEETLES (CURCULIONIDAE: SCOLYTINAE: XYLEBORINI) AND THEIR FUNGAL PARTNERS By Rachel Kathryn Osborn A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Entomology – Doctor of Philosophy Ecology, Evolutionary Biology and Behavior – Dual Major 2022 ABSTRACT DIVERSITY OF SOUTH AMERICAN AMBROSIA BEETLES (CURCULIONIDAE: SCOLYTINAE: XYLEBORINI) AND THEIR FUNGAL PARTNERS By Rachel Kathryn Osborn Ambrosia beetles from the tribe Xyleborini (Coleoptera: Curculionidae: Scolytinae) small, haplodiploid beetles that farm nutritional fungi on the walls of tunnels they excavate in the xylem of dead or nearly dead trees. These biological traits make them successful participants in worldwide wooded ecosystems and facilitate their human-mediated invasion beyond their native ranges. A minority of these introduced species are classified as pests because of the physical damage they cause to their plant hosts, or because they vector pathogenic fungi that infect ornamental, lumber, and forest trees. Most of the current knowledge on the diversity of xyleborine beetles and their fungi centers around species found in North America, Asia, and Europe. Little is known about the ambrosia partnerships in the Neotropics, which is concerning because South America is a strong trading partner with the US and the potential for new invasive Xyleborini to be imported from this area is significant. Continuing forest damage caused by invasive Xyleborini/fungi inspires robust research efforts to describe these symbionts and document their biological traits. Considerable efforts are required to enhance such endeavors in underrepresented regions such as South America and Africa. To increase understanding of the South American Xyleborini and their associated fungi, I compiled current knowledge of their historical and contemporary taxonomic records, biological records, and ecological studies. I also completed surveys throughout Ecuador to collect beetles and fungi. Molecular analysis of fungi isolated from Ecuadorian beetles reveals that several Coptoborus species associate with Fusarium fungi, including the ambrosia Fusarium Clade (AFC) that has previously been recovered from Euwallacea spp. and Xyleborus ferrugineus in Central America, Florida, California, Israel, and Asia. Examination of the morphology of some South American xyleborine specimens previously classified as Coptoborus spp. suggests a high similarity to Xyleborus spp. from Africa. Phylogenetic analysis of these South American and African beetles as well as morphological assessment of additional specimens necessitates the designation of a new genus Xenoxylebora gen. nov. containing species endemic to both continents. This unusual distribution demonstrates the ability of these ambrosia beetles to survive long-distance trans-oceanic dispersal. Copyright by RACHEL KATHRYN OSBORN 2022 This dissertation is dedicated to my father, Dr. Michael Vernon Osborn, who would be so proud. And to my husband, Davin Taddeo whose love and support made this work possible. v ACKNOWLEDGEMENTS I thank my advisor, Anthony Cognato for his patient mentorship and for always being on my side. I thank Greg Bonito, Jeff Conner, and Christina DiFonzo for their time and expertise as members of my guidance committee. I also thank Sarah Cognato and Gary Parsons for their mentorship and advice. Thank you to my friends at MSU who have given me the support and encouragement I needed to complete this work: Nikki Cavalieri, Dan Hulbert, Courtney Larson, Connie Rojas, Taylor Rupp, and Dan Turner. I thank my mother Lin Osborn, husband Davin Taddeo, and Jan and Russ Taddeo for their love and reassurance. I thank my collaborators from the Hulcr lab at the University of Florida, Gainesville, especially Violet Butterworth and Jiri Hulcr for their instruction, advice, and assistance in the field. Thank you to my Ecuadorian collaborators, Jessenia Castro, Cliff Keil, Malena Martínez, Jim McClarin, Maria Eugenia Ordoñez, and Jane Sloan for their help and generosity. Thank you to La Universidad Técnica Estatal de Quevedo, La Pontifica Universidad Católica del Ecuador, La Estación Yasuní, Yanayacu Biological Station, Yakusinshi Ecological Reserve for graciously allowing me to use their facilities for specimen collection, preparation, and examination. I also thank the following institutions for lending specimens used in this work: Canadian National Collection of Insects (Ottawa), Escuela Politécnica Nacional Instituto de Ciencias Biologicas (Quito), Michigan State University Albert J. Cook Arthropod Research Collection (East Lansing, Michigan), Musée Royal de l’Afrique Centrale (Tervuren), National Zoological Collection of Suriname (Paramaribo), Natural History Museum (London), Naturhistorisches Museum Wien (Vienna), Smithsonian National Museum of Natural History (Washington, DC), Universidad Nacional Mayor de San Marcos (Lima), and University of Kansas Biodiversity Institute & Natural History Museum (Lawrence, Kansas). vi Finally, I thank the many organizations who provided funding for this research: National Geographic, the Coleopterists’ Society, the Society of Systematic Biologists, and the Department of Entomology at Michigan State University. I am grateful for these organizations and for the people within them who believed in my research. vii TABLE OF CONTENTS LIST OF TABLES .......................................................................................................................... x LIST OF FIGURES ....................................................................................................................... xii INTRODUCTION ........................................................................................................................... 1 LITERATURE CITED .................................................................................................................... 5 CHAPTER 1: Diversity of xyleborine ambrosia beetles, their associated fungi, and the imperative of global collaboration................................................................................................. 10 ABSTRACT ...................................................................................................................... 10 INTRODUCTION ............................................................................................................. 10 Ecology of Xyleborini ........................................................................................... 12 History of research regarding harmful Xyleborini ................................................ 13 AMBROSIA FUNGI OF THE XYLEBORINI ................................................................ 15 CURRENT AND EMERGING PESTIFEROUS XYLEBORINE BEETLE/FUNGAL PARTNERSHIPS .............................................................................................................. 17 Xyleborus glabratus and Harringtonia lauricola .................................................. 17 Xylosandrus spp. and Ambrosiella spp. ................................................................. 18 Ambrosia Fusarium Clade and Euwallacea spp. + Coptoborus spp. ................... 25 Ambrosiodmus minor and Irpex subulatus ............................................................ 28 AMBROSIA BEETLE/FUNGAL RESEARCH IN REGIONAL CENTERS OF DIVERSITY ...................................................................................................................... 29 ENHANCING GLOBAL RESEARCH OF AMBROSIA SYMBIOSIS .......................... 31 APPENDIX ................................................................................................................................... 33 LITERATURE CITED .................................................................................................................. 47 CHAPTER 2: Ecuadorian Coptoborus beetles harbor Fusarium and Graphium fungi previously associated with Euwallacea ambrosia beetles ............................................................................... 66 ABSTRACT ...................................................................................................................... 66 INTRODUCTION ............................................................................................................. 67 METHODS ........................................................................................................................ 71 Material collection ................................................................................................. 71 Molecular data ....................................................................................................... 73 Phylogenetic analysis ............................................................................................ 74 RESULTS .......................................................................................................................... 77 Fungal identification via ITS ................................................................................. 77 Specific identification of ambrosia fungi via phylogenetic analyses .................... 78 DISCUSSION.................................................................................................................... 80 ACKNOWLEDGEMENTS .............................................................................................. 85 APPENDIX ................................................................................................................................... 86 LITERATURE CITED ................................................................................................................ 125 viii CHAPTER 3: New xyleborine (Coleoptera: Curculionidae: Scolytinae) genus with an Afrotropical-Neotropical distribution .......................................................................................... 137 ABSTRACT .................................................................................................................... 137 INTRODUCTION ........................................................................................................... 138 METHODS ...................................................................................................................... 142 Taxon sampling ................................................................................................... 142 Molecular dataset................................................................................................. 143 Phylogenetic analyses .......................................................................................... 144 Biogeographical analyses .................................................................................... 145 Taxonomy ............................................................................................................ 146 Data availability................................................................................................... 147 RESULTS ........................................................................................................................ 148 Phylogenetic analysis .......................................................................................... 148 Biogeographical analyses .................................................................................... 149 Taxonomy ............................................................................................................ 150 Xenoxylebora Osborn, Smith & Cognato gen. nov. ................................ 150 Xenoxylebora addenda Osborn, Smith & Cognato sp. nov. ................... 152 Xenoxylebora calculosa Osborn, Smith & Cognato sp. nov................... 154 Xenoxylebora caudata (Schedl, 1957) comb. nov. ................................. 155 Xenoxylebora collarti (Eggers, 1932) comb. nov. .................................. 156 Xenoxylebora hystricosa Osborn, Smith & Cognato sp. nov. ................ 158 Xenoxylebora neosphenos (Schedl, 1976) comb. nov. ........................... 160 Xenoxylebora perdiligens (Schedl, 1937) comb. nov. ............................ 161 Xenoxylebora pilosa Osborn, Smith & Cognato sp. nov. ....................... 162 Xenoxylebora serrata Osborn, Smith & Cognato sp. nov. ..................... 164 Xenoxylebora sphenos (Sampson, 1912) comb. nov. ............................. 165 Xenoxylebora subcrenulata (Eggers, 1932) comb. nov. ......................... 167 Xenoxylebora sulcata Osborn, Smith & Cognato sp. nov. ..................... 167 Xenoxylebora syzygii (Nunberg, 1959) comb. nov. ................................ 169 Key to Xenoxylebora species (females only) ...................................................... 170 DISCUSSION.................................................................................................................. 172 ACKNOWLEDGEMENTS ............................................................................................ 175 APPENDIX ................................................................................................................................. 176 LITERATURE CITED ................................................................................................................ 202 ix LIST OF TABLES Table 1.1: Introduced Xyleborini, their native region, and region(s) of introduction. Data inferred from Wood and Bright 1992, Pennacchio et al. 2003, Rabaglia et al. 2006, Kirkendall and Ødegaard 2007, Wood 2007, Cognato and Rubinoff 2008, Kirkendall and Faccoli 2010, Haack and Rabaglia 2013, Gomez et al. 2018, Schiefer 2018 Lin et al. 2021, and Urvois et al. 2022 ............................................................................................................................................... 34 Table 1.2: Ambrosia fungal species associated with xyleborine beetles and their beetle partners. Inferred from von Arx and Hennebert 1965, Batra 1967, Scott and du Toit 1970, Brayford 1987, Gebhardt 2005, Harrington et al. 2008, Six et al. 2009, Harrington et al. 2010, Kasson et al. 2013, Mayers et al. 2015, O’Donnell et al. 2015, Simmons et al. 2016a, 2016b, Lin et al. 2017, Mayers et al. 2017, Na et al. 2018, Carrillo et al. 2019, Lynn et al. 2020, Nel et al. 2021, and Osborn et al. 2022a ........................................................................................................................ 38 Table 1.3: Harmful Xyleborini, their nutritional symbionts, associated pathogenic fungi, native region, year of first report in introduced regions, and significant plant hosts. Inferred from Blanford 1894a, Hagedorn 1908, Hoffmann 1941, Groschke 1953, Anderson 1974, Kessler et al. 1974, Hara and Beardsley 1979, Wood 1982, Weber and McPherson 1983, Nirenburg 1990, Pennacchio et al. 2003, Haack 2006, Rabaglia et al. 2006, Kirkendall and Ødegaard 2007, Cognato and Rubinoff 2008, Olivera et al. 2008, Eskalen et al. 2012, Garonna et al. 2012, Mendel et al. 2012, Stilwell et al. 2014, Egonyu et al. 2015, Mayers et al. 2015, Nageleisen et al. 2015, You et al. 2015, Flechtmann and Atkinson 2016, Simmons et al. 2016a, Gallego et al. 2017, Li et al. 2017, Kavčič 2018, Paap et al. 2018, Schiefer 2018, Carreras-Villaseñor et al. 2022, and Osborn et al. 2022a. ...................................................................................................... 42 Table 2.1: Fusarium and Graphium fungal isolates cultured from Ecuadorian Coptoborus beetles collected in Yasuní, Orellana, Ecuador and used in phylogenetic analyses ...................... 87 Table 2.2: Amplification primers used for gene sequencing ........................................................ 89 Table 2.3: PCR conditions used for gene sequencing .................................................................. 90 Table 2.4: Fusarium and Graphium fungal sequences obtained from GenBank and used in phylogenetic analysis. Sources: aO'Donnell et al. 2015, bKasson et al. 2013 cCarrillo et al. 2019, d Mendel et al. 2012, eNa et al. 2018, fZhang et al. 2006, gO'Donnell direct submission, hO'Donnll et al. 2007, iO'Donnell et al. 2010, jLynch et al. 2016, kCruywagen et al. 2010, lPaciura et al. 2008, mJacobs et al. 2003, nHulcr et al. 2007, oOkada et al. 2000, pTwizeyimana et at Direct Submission, qKolarik et al. unknown date, and rHameline et al. Direct Submission. *Outgroup. ** Fungus was isolated from xyleborine gallery. TEx-type strain. ?Identification of this beetle partner is questionable (O'Donnell et al., 2015) ............................................................................ 91 Table 2.5: BLAST identities of fungi isolated from Coptoborus spp. ....................................... 111 x Table 3.1: Specimens used for molecular phylogenetic reconstruction including voucher name, specific identification, collection location, publication source, and GenBank numbers ............. 177 Table 3.2: Primers used for PCR reactions................................................................................. 186 Table 3.3: Models of nucleotide substitution assigned by PartitionFinder 2.1.1 and ModelFinder, respectively, and used for phylogeny reconstruction by likelihood and Bayesian analyses. JC = Jukes Cantor model of evolution with equal base frequencies and equal substitution rates (Jukes and Cantor 1969); F81 = Felsenstein model with unequal base frequencies and equal rates of substitution (Felsenstein 1981); K80 = Kimura model with equal base frequencies and unequal rates of transition and transversion substitutions (Kimura 1980); HKY = Hasegawa model with unequal base frequencies and unequal rates of transition and transversion substitutions (Hasegawa et al. 1985); TN = Tamura Nei model with unequal rates of purine and pyrimidine rates and unequal rates of transition and transversion substitutions (Tamura and Nei 1993); TIM2e =Transition model with equal base frequencies and AC and CG substitution rates equal to AT and GT, respectively; TIM3e = Transition model with equal base frequencies and AC and AT substitution rates equal to CG and GT, respectively; GTR = general time reversible model with unequal base frequencies and unequal rates of substitution (Lanave et al. 1984, Rodríguez et al. 1990); I = variable nucleotide frequencies; F = nucleotide frequencies determined from the data; FQ = nucleotides with equal frequencies; G = gamma distributed rates of variation; R = FreeRate model of distributed rates of variation (Yang 1995, Soubrier et al. 2012) .................. 187 Table 3.4: Biogeographical states used for biogeography analyses ........................................... 189 xi LIST OF FIGURES Figure 1.1: Schematic representation of ambrosia associations between xyleborine and fungal genera. Cladogram of the Xyleborini summarized from Cognato et al. 2011, Cognato et al. 2018, and Johnson et al. 2018. Relationships between the beetles and fungi from von Arx and Hennebert 1965, Batra 1967, Scott and du Toit 1970, Brayford 1987, Gebhardt 2005, Harrington et al. 2008, Six et al. 2009, Harrington et al. 2010, Kasson et al. 2013, Mayers et al. 2015, O’Donnell et al. 2015, Simmons et al. 2016a, 2016b, Lin et al. 2017, Mayers et al. 2017, Na et al. 2018, Carrillo et al. 2019, Lynn et al. 2020, Nel et al. 2021, Osborn et al. 2022a ................... 46 Figure 2.1: Condensed Bayesian consensus tree of Fusarium strains reconstructed from ITS, EF1-a, RPB1, and RPB2 sequence data. Nodes are labeled with bootstrap values/posterior probabilities. Clades containing AFC strains associated with Euwallaceae spp. beetles have been collapsed. Fusarium sp. isolated from Coptoborus sp. for this study are written in bold. See Figure 2.2 for the complete tree .................................................................................................. 115 Figure 2.2: Complete Bayesian consensus tree of Fusarium strains reconstructed from ITS, EF1- a, RPB1, and RPB2 sequence data. Nodes are labeled with bootstrap values/posterior probabilities. Fusarium sp. isolated from Coptoborus sp. for this study are written in bold ..... 116 Figure 2.3: Bayesian consensus tree of Fusarium strains built from gene sequence data of EF1- a. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold ........................................................................ 117 Figure 2.4: Bayesian consensus tree of Fusarium strains built from gene sequence data of ITS. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold............................................................................................ 118 Figure 2.5: Bayesian consensus tree of Fusarium strains built from gene sequence data of RPB1. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold............................................................................................ 119 Figure 2.6: Bayesian consensus tree of Fusarium strains built from gene sequence data of RPB2. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold............................................................................................ 120 Figure 2.7: Condensed Bayesian consensus tree of Graphium strains reconstructed from ITS, and EF1-a sequence data ............................................................................................................ 121 Figure 2.8: Complete Bayesian consensus tree of Graphium strains reconstructed from ITS, EF1-a, and RPB2 sequence data. Nodes are labeled with bootstrap values/posterior probabilities. Graphium sp. isolated from Coptoborus sp. for this study are written in bold. T Denotes ex-type strains ........................................................................................................................................... 122 xii Figure 2.9: Bayesian consensus tree of Graphium strains built from gene sequence data of EF1- a. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold. T Denotes ex-type strains .............................. 123 Figure 2.10: Bayesian consensus tree of Graphium strains built from gene sequence data of ITS. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold. T Denotes ex-type strains .................................................. 124 Figure 3.1: Map of Scolytinae with native distributions across Africa and South America (Wood and Bright 1992, Jordal 2012, Hulcr et al. 2015, Gohli et al. 2016, Bright 2010, 2019, Atkinson 2021, Eliassen and Jordal 2021, Jordal 2021b, 2021c, 2021d, 2021e, 2021f). Each circled number represents the number of species from the corresponding group endemic or established in the indicated continent. The expanded side of each line indicates the continent containing a plurality of species. Two Premnobius species have cosmopolitan distributions and are therefore each counted once on each continent: P. cavipennis and P. ambitiosus. Xenoxylebora is depicted in blue; black indicates xyleborine genera, and grey indicated non-xyleborine groups .................. 195 Figure 3.2: Phylogenetic tree resulting from a Bayesian analysis of CO1, CAD, EF1-α and 28S. Nodes are labeled with posterior probability/bootstrap support. Posterior probabilities > 0.95 are considered strong clade support. Nodes within Xenoxylebora are additionally labeled with pie diagrams indicating the relative probabilities of origin in the Neotropics, Afrotropics, and Neotropics/Afrotropics. An outgroup Anisandrus sayi was used to root the tree ....................... 196 Figure 3.3: Maximum likelihood tree resulting from Nearest Neighbor Interchange search of CO1, CAD, EF1-α and 28S sequences using IQ-TREE version 2.1.3. Nodes are labeled with bootstrap support from 1000 pseudoreplications. Anisandrus sayi was used to root the tree ..... 197 Figure 3.4: Diagnostic characters for Xenoxylebora. (3.4.3) type 1 antennal club. (3.4.4) slender protibia, outer edge weakly rounded, posterior face flat and unarmed. (3.4.5) sutural tubercles on the declivital apex ........................................................................................................................ 198 Figure 3.5: Dorsal, lateral, and declivital aspects of Xenoxylebora addenda holotype (3.5.6– 3.5.8); Xenoxylebora calculosa holotype (3.5.9–3.5.11); Xenoxylebora caudata paratype (3.5.12– 3.5.14); Xenoxylebora collarti (3.5.15–3.5.17); Xenoxylebora hystricosa holotype (3.5.18– 3.5.20) .......................................................................................................................................... 199 Figure 3.6: Dorsal, lateral, and declivital aspects of Xenoxylebora neosphenos (3.6.21–3.6.23); Xenoxylebora perdiligens (3.6.24–3.6.26); Xenoxylebora pilosa holotype (3.6.27–3.6.29); Xenoxylebora serrata holotype (3.6.30–3.6.32); Xenoxylebora sphenos (3.6.33–3.6.35) .......... 200 Figure 3.7: Dorsal, lateral, and declivital aspects of Xenoxylebora subcrenulata holotype (3.7.36–3.7.38); Xenoxylebora sulcata holotype (3.7.39–3.7.41); Xenoxylebora syzygii paratype (3.7.42–3.7.44) ............................................................................................................................ 201 xiii INTRODUCTION As global ecological homogeny increases, the economic and ecological impact of invasive species continues to increase in the United States (Seebens et al. 2017). Bark and ambrosia beetles from the weevil subfamily Scolytinae are especially likely to be introduced to the United States from exotic locations. All life stages of these small beetles are spent inside woody plants, which makes them easily overlooked as they cross boarders via international trade networks inside ornamental plants of wood packing material. (Rabaglia et al. 2019, Lantschner et al. 2020). Most bark and ambrosia beetles prefer recently deceased host trees because these plant hosts cannot defend themselves against the tunneling beetles (Hubbard 1897). Such trees are also possess optimal physical conditions for brood development, provide protection from predators, and are unlikely to house competing fungi or wood-boring animals. Only a small number of scolytine species attack living trees and are thus destructive to forests and food crop and ornamental trees (Smith and Hulcr 2015). Ambrosia beetles differ from bark beetles in that they burrow into the xylem of their host trees and rely on the growth of symbiotic fungi that they farm inside these galleries for nutrition. Bark beetles tunnel under the bark and feed on the phloem of their plant hosts. Ambrosia beetles have the potential to vector devastating plant diseases if they are partnered with phytopathogenic fungi (Hulcr and Dunn 2011, Carrillo et al. 2014). For instance, Coptoborus ochromactonus Smith and Cognato has caused significant damage to trees in balsa plantations in Ecuador beginning in the 1990s and was recently discovered to associate with a putatively lethal strain of Fusarium fungus (Stilwell et al. 2014, Osborn et al. 2022). 1 The beetle-fungus mutualism also makes ambrosia beetles more efficient at expanding their geographical range with (Rassati et al. 2016, Lantschner et al. 2020) and without (Gohli et al. 2016) human mediation. Their reliance on fungi for nutrition affords them a greater variety of possible host taxa than bark beetles which have a more constrained host breadth (Francke- Grosmann 1967, Bright 1968). Ambrosia beetles require host trees that have adequate moisture and nutrition for their fungal symbionts, and which are free from contaminating fungi that might outcompete the ambrosia fungi (Freeman et al. 2016, Cavaletto et al. 2021, Nuotclà et al. 2021). Outside of these habitat requirements, ambrosia beetles can successfully colonize a variety of trees. The host range available to a given bark beetle is smaller because their phloem-feeding lifestyle places them in direct interaction with the chemistry of their hosts (Kirisits 2004). Thus, when bark beetles are introduced to novel environments, they are unlikely to survive unless they encounter host trees that meet their specific nutritional needs. Ambrosia beetles carry their fungal partners with them as they disperse to new trees. They readily establish in new locales because growth requirements of many ambrosia fungi can be fulfilled by a phylogenetically diverse group of trees. This invasion potential is enhanced in the ambrosia beetle tribe Xyleborini because these genera have a haplodiploid mating system. Females lay fertilized eggs which develop into female offspring, and unfertilized eggs which become males. The male offspring stay inside the ambrosia gallery and mate with their sisters. Therefore, when a female xyleborine is introduced to a new ecosystem, she needs only to find a suitable place to plant a new ambrosia fungal garden since she was already fertilized in her natal gallery (Jordal et al. 2001, Peer and Taborsky 2005). 2 These biological traits allowed xyleborine ambrosia beetles to establish new populations on distant continents. Lowered Allee Effect thresholds likely contributed to the success of the first individuals as they arrived in the Neotropics from the Nearctic region 23 million years ago (Jordal et al. 2000, Liebhold and Kean 2018, Lantschner et al. 2020). These early xyleborines experienced a rapid rate of speciation, resulting in the hyperdiversity of xyleborines present in the South America today (Jordal et al. 2000, Gohli et al. 2017). This radiation, in turn, has no doubt fueled further dispersal and colonization of ambrosia beetles throughout the Americas, and has caused perpetual challenges to scientists as they catalogue species and untangle taxonomic relationships. This is particularly true of the South American members of the genus Xyleborus (Jordal et al. 2000, Cognato et al. 2011). About 40 xyleborine species have established invasive populations beyond their native geographic ranges (Rabaglia et al. 2006, Kirkendall and Faccoli 2010, Haack and Rabaglia 2013, Gomez et al. 2018, Lin et al. 2021). This includes several Xylosandrus native to Asia that now infest a broad range of trees, damaging forests, lumber plantations, nurseries, and food crop trees all around the world (Atkinson et al. 2000, Kirkendall and Ødegaard 2007, Egonyu et al. 2015, Reding et al. 2015, Flechtmann and Atkinson 2016, Dzurenko et al. 2021, Carreras-Villaseñor et al. 2022, Urvois et al. 2022). Xyleborus glabratus Eichhoff, which is also native to Asia, was first discovered in the United States in 2002 and has since spread throughout the Southern states where it attacks live individuals of several tree species, especially those in the Lauraceae family (Haack 2006). The symbiotic relationship between X. glabratus and the pathogenic fungus Harringtonia lauricola (Harrington, Fraedrich and Aghayeva) (= Raffaelea lauricola) exemplifies the danger of virulent fungi paired with invasive ambrosia beetles. Xyleborus glabratus and H. lauricola continue to cause significant damage to avocado trees, ornamental 3 plants, and forest ecosystems from Florida to Texas (Inch et al. 2012, Ploetz et al. 2017, Wingfield et al. 2017). Losses caused by Xyleborine invaders and their fungi from the Eurasia have inspired extensive research regarding their distributions, evolutionary histories, and biology. However, there is less information about Neotropical xyleborine beetles and their fungi (Wood 2007, Smith et al. 2017). The United States increasingly imports food, plants, and other raw materials from South America, and therefore risks importing invasive ambrosia beetles from our southern neighbors – and potentially harmful fungi (Marini et al. 2011, Meurisse et al. 2019, Lantschner et al. 2020). This dissertation examines current knowledge on the taxonomy, evolutionary history, and ecology of the xyleborine-fungal symbiome and adds to this knowledge through a survey of endemic ambrosia beetles and their fungi from Ecuador. I use the knowledge and material from this research to address four questions: (i) what fungi were associated with xyleborine species in previous research? (ii) which fungi are carried by various Coptoborus species collected in Ecuador? (iii) what new taxonomic boundaries can be established to better-reflect the evolutionary past of Neotropical xyleborine genera? And (iv) do new generic boundaries need to be created? 4 LITERATURE CITED 5 LITERATURE CITED Atkinson TH, JL Foltz, and RC Wilkinson. 1988. Xylosandrus crassiusculus (Motschulsky), an Asian ambrosia beetle recently introduced into Florida (Coleoptera: Scolytidae). Entomology Circular. 310: 1–4. Bright DE. 1968. Review of the tribe Xyleborini in America north of Mexico (Coleoptera: Scolytidae). The Canadian Entomologist. 100: 288–1323. Carreras-Villaseñor N, JB Rodríguez-Haas, LA Martínez-Rodríguez, AJ Pérez-Lira, E Ibarra- Laclette, E Villafán, AP Castillo-Díaz, LA Ibarra-Juárez, ED Carrillo-Hernández, and D Sánchez-Rangel. 2022. Characterization of two Fusarium solani species complex isolates from the ambrosia beetle Xylosandrus morigerus. Journal of Fungi. 8: 231. Carrillo JD, RE Duncan, JN Ploetz, AF Campbell, RC Ploetz, and JE Peña. 2014. Lateral transfer of a phytopathogenic symbiont among native and exotic ambrosia beetles. Plant Pathology. 63: 54–62. 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Bark and Ambrosia Beetles of South America (Coleoptera: Scolytidae) Monte L Bean Science Museum, Brigham Young University, Provo, Utah. 900 p. 9 CHAPTER 1: Diversity of xyleborine ambrosia beetles, their associated fungi, and the imperative of global collaboration ABSTRACT Ambrosia beetles from the tribe Xyleborini are part of forest ecosystems on every continent except Antarctica. Because of their small size, haplodiploid mating structure, and protected lives inside the sapwood of woody plants, they have a unique ability to expand into new regions via inadvertent human transport. A small number of invasive xyleborines cause significant damage to forests, lumber concerns, and agricultural systems. The most dangerous of these transmit pathogenic fungi capable of causing disease in or killing living trees. The relationships between these fungi and their beetle vectors range from mutualistic symbiosis to facultative association and many are not understood. Unstable taxonomies, convergent morphologies and the difficulty of obtaining and isolating ambrosia fungi accross their entire global ranges make comprehensive surveys of ambrosia fungi difficult to achieve. Ambrosia fungi from Asia, Europe and North America are fairly well documented, however we have yet to sufficiently document those from Africa, and South America. Worldwide cooperation to improve and standardize scientific study of the ambrosia symbiome is needed to better understand these economically impactful organisms. INTRODUCTION Forest biomes, with their abundance of trees and other flora, are prominent ecosystems on every continent except Antarctica. As a biological system, forests experience a constant cycle of death, 10 decomposition, and renewal which provides a steady supply of dead trees as a ubiquitous source of carbon and nutrients for terrestrial environments (Ulyshen 2016, Klemm et al. 2005). Although cellulose, hemicellulose, and lignin that form plant cell walls are indigestible to most organisms, many animals rely on these molecules to provide much of their energetic needs. The difficult task of converting plant polysaccharides into shorter sugars is dominated by bacteria, fungi, and some protists (Flint et al. 2008). Many mammals rely on plant digesting microbes as endosymbionts (Martins et al. 2014), and several groups of insects independently evolved strategies involving microbes to gain access to the nutrients within plant cell walls. For example, most termites employ a highly structured community of bacteria, archaea, and co-evolved protozoa that allow them to degrade plant cell walls more efficiently than any other herbivorous animal (Brune and Okhuma 2011). Several groups of insects use plant-degrading fungi to convert plant material into a nutritional food source. The relationships between cellulolytic fungi and these insects take various forms ranging from obligate mutualism with attine ants and macrotermitine termites, to commensal associations with the bee species Scaptotrigona depilis (Moure), silphid and lymexylid beetles, and inchoate interactions with Euops attelabid beetles, and Doubledaya erotilid beetles (Kobayashi et al. 2008, Paludo et al. 2019, Biedermann and Vega 2020, Toki and Aoki 2021). Bark and ambrosia beetles from the curculionid subfamily Scolytinae contain species with a variety of relationships with wood-degrading fungi. Many bark beetles transmit spores and maintain an opportunistic relationship with them as cohabitates of the same host plants (Six 2003, Harrington 2005). They supplement their diets with fungi and transport them in specialized organs called mycangia. However, they do not require these fungi to survive. Scolytine species that maintain a mutualistic symbiosis with fungi are called ambrosia beetles. Wood-degrading 11 fungi are the sole food source for adults and larvae of these beetles. Ambrosia fungi benefit from this association because they are protected from parasites and competing fungi (Batra 1967, Kirisits et al. 2004, Biedermann and Vega 2020) and given access to suitable growing substrates with appropriate nutrient and moisture levels (Rassati et al. 2016, Nuotclá et al. 2021). Independent phylogenetic analyses found that the ambrosia fungal feeding evolved at least 12 times in Scolytinae and once in the weevil subfamily Platypodinae (Jordal and Cognato 2012, Gohli et al. 2017, Johnson et al. 2018, Pistone et al. 2018). Some ambrosia beetle lineages are quite diverse, and research suggests that fungus farming is one factor that contributes to their species richness (Jordal and Cognato 2012, Gohli et al. 2017). Ecology of Xyleborini The tribe Xyleborini contains ~1260 species (Smith, unpublished), all of which are ambrosia beetles. They are found in forested habitats worldwide, experiencing rapid speciation events on more than one continent (Jordal et al. 2000, Jordal and Cognato 2012, Cognato et al. 2018, Eliassen and Jordal 2021). They also successfully invade new habitats at a higher rate than non- ambrosia scolytines (Rabaglia et al. 2019, Lantschner et al. 2020). Xyleborines possess a suite of biological characteristics that make them uniquely equipped for transport to, and proliferation in, novel environments. Life inside sapwood protects them from the elements and allows them to survive long distance dispersal via ocean currents (Cognato 2013, Jordal 2015, Cognato et al. 2018) and trade (Seebens et al. 2017, Grousset et al. 2020). Once in a new environment, xyleborine beetles are likely to reproduce and successfully establish a population because they carry their food source with them. When the females leave the fungal garden, they carry the nutritional fungi with them inside specialized exoskeletal structures called mycangia (Francke- 12 Grosmann 1956, Schedl 1962). Since ambrosia fungi can grow on many species of trees given the correct environmental and moisture conditions, dispersing females are likely to find suitable hosts in unfamiliar environments if the climate is relatively similar to that in their native range (Rassati et al. 2016). Furthermore, female xyleborines lay unfertilized eggs that mature into flightless haploid males which mate with their mother (if unfertilized) and sisters inside their natal gallery. Thus, dispersing females do not need to find a mate before founding a new ambrosia colony and reproducing. These traits allow one foundress to establish a population, which makes them excellent colonizers and potential pests in non-native ranges. Their subcortical habitat and small size (1–3 mm long) conceal them from customs and border inspectors (Seebens et al. 2017). Approximately 40 xyleborine species have established populations beyond their native ranges due to human accidental introduction (Table 1.1) (Rabaglia et al. 2006, Kirkendall and Faccoli 2010, Haack and Rabaglia 2013, Gomez et al. 2018, Lin et al. 2021). History of research regarding harmful Xyleborini Ambrosia beetles have long caused ecological and financial harm to forests and the lumber industry in Europe and North America (Schmidberger 1836, Hubbard 1897). However, despite knowing that ambrosia beetles bore into the sapwood and often vector undesirable fungi, the earliest researchers did not appreciate the symbiotic nature of the beetle-fungal association. Many noted that ambrosia beetles ate a white powery substance that was never observed to be associated with bark beetles (Schmidberger 1836, Ratzeburg 1839). Schmidberger (1836) established the term “ambrosia” to describe the beetles’ diet because of its fruity odor, he suspected it was made by females out of tree sap and saliva to feed her offspring (Beling 1873). 13 When Hartig (1844) documented the fungal nature of ambrosia he classified the fungus associated with Anisandrus dispar (Fabricius) as Monila candida (Persoon). More than five decades later, Hubbard (1897) recognized that ambrosia fungi provide nutrition to the beetles and are actively cultivated in a symbiotic relationship. Before the middle of the twentieth century, investigations into the ambrosia symbiome were generally flawed with the assumption that each ambrosia beetle species lives with one obligate fungal symbiont (e.g., Neger 1909, Doane and Gilliand 1929, Leach et al. 1940) and common misidentifications of ambrosia fungi (e.g., Neger 1911, Muller 1933, Verall 1943). As ambrosia research continued, scientists discovered complex assemblages of fungi inhabiting ambrosia beetle galleries (Norris 1965, Batra 1966, Norris 1979). Batra (1967) reclassified the known ambrosia fungi at the time into eight genera, of which three were associated with xyleborines: Ascoidea, Ambrosiella, and Monacrosporium. Recent ambrosia research revealed the complex and varied range of relationships shared among xyleborine ambrosia beetles and fungi (Kostovcik et al, 2015, Skelton et al. 2018). This includes the identification of many fungal symbionts (von Arx and Hennebert, 1965, Harrington et al. 2008, Freeman et al. 2013, Harrington et al. 2014, Li et al. 2016, Lynch et al. 2016, Simmons et al. 2016a, 2016b, Aoki et al. 2018, 2019, Lynn et al. 2020, Aoki et al. 2021) and recognition that the same fungal species might grow in the galleries of both non-xyleborines xyleborines (Gebhart et al. 2004). The discovery and identification of the mycangium (Francke- Grosmann 1956, Schedl 1962, Batra 1963) revealed the complex role this organ plays in the development and maintenance of the ambrosia symbiome (Mayers et al. 2022). The evolution of mycangia correlates with the ambrosia lifestyle throughout Scolytinae (Mayers et al. 2015), and mycangia and their corresponding fungal partners from several ambrosia lineages are coadapted 14 (Johnson et al. 2018, Mayers et al. 2020b). The mycangium/fungi interaction also allows beetles to ensure vertical transmission of the most desirable fungal partners (Skelton et al. 2019), which extends their ability to survive variable conditions such as changing moisture levels inside the gallery and defensive compounds from new host plants (Hulcr and Dunn 2011, Nuotclá et al. 2021). AMBROSIA FUNGI OF THE XYLEBORINI Xyleborines have established mutual symbioses with a taxonomically diverse group of fungi within three orders of Ascomycota and one from Basidiomycota (Table 1.2; Figure 1.1) (Bateman 2018). Given the beetles’ unique capacity for range expansion, their fungal partners share similarly large geographical ranges. Molecular dating estimates indicate that the first fungus farming xyleborines appeared ~21 million years ago followed by a rapid evolutionary radiation fueled by global warming during the early Miocene (Jordal and Cognato 2012). This timing is incongruent with the origins of the xyleborine fungal lineages. Phylogenetic analysis of Raffaelea/Harringtonia shows that the three clades of ambrosia fungi in the genus emerged 86 Ma, 67 Ma, and 33 Ma ago, before the current estimate for the origin of Xyleborini (Vanderpool 2017). Similarly, the Ambrosia Fusarium Clade (AFC), which includes both Fusarium and Neocosmospora species, is ~24 million years old (O’Donnell et al. 2015, de Beer et al. 2022). Mayers et al. (2020b) determined the first appearance of the Ambrosiella lineage associated with xyleborines to be ~12 million years ago, ~9 million years after the origin of Xyleborini. The ages of the ambrosia fungi Dryadomyces and Irpex subulatus (Ryvarden) have not been estimated, but further investigation of the timing of their entrance into the ambrosia symbiome could further illuminate the mechanisms of ambrosial evolution. 15 Current knowledge of the dissimilar patterns of emergence between the Xyleborini and their ambrosia fungi in Raffaelea/Harringtonia, Fusarium/Neocosmospora and Ambrosiella suggests that xyleborine beetles obtained their current ambrosia fungi through lateral transmission (partner switching) (O’Donnell et al. 2015, Peris et al. 2021). Raffaelea/Harringtonia is known to have experienced frequent partner switching (Miller et al. 2019) including at least three independent transitions into symbiosis with xyleborine beetles (Vanderpool 2017). Frequent symbiont switching, including a recent shift to Neotropical Xyleborini was found to be the most likely explanation for the confused phylogenetic patterns in Fusarium (AFC) (O’Donnell et al. 2015, Peris 2021, Osborn et al. 2022a). A constrained lineage of Ambrosiella was also found to have transferred from their historical platypodine beetle hosts to Xylosandrus spp. within Xyleborini (Mayers et al. 2020a). After the first appearance of the invasive xyleborine Xyleborus glabratus Eichhoff and the destructive fungus Harringtonia lauricola (T. C. Harr., Fraedrich and Aghayeva) in the United States, surveys detected H. lauricola in the mycangia of Xyleborus spp., Xylosandrus spp. and two Ambrosiodmus spp. that commonly inhabit the same tree hosts as X. glabratus (Carrillo et al. 2014, Ploetz et al. 2017). Xyleborus bispinatus Eichhoff fed and reproduced when experimentally reared in a garden of H. lauricola (Saucedo et al. 2017). The fungus is also known to cohabitate with other Raffaelea species, both inside the mycangium and upon beetle gallery walls (Harrington et al. 2010, 2011, Simmons et al. 2016b). These studies show that the fungal symbiont of one beetle can enter the mycangium of a different species when the galleries exist in the same tree. They also support the hypothesis posited by Hulcr and Dunn (2011) that invasive ambrosia beetles are more likely to switch to a pathogenic fungus when they expand 16 into new geographic regions or exploit novel host species, because new fungi may enhance their ability to survive tree defenses. CURRENT AND EMERGING PESTIFEROUS XYLEBORINE BEETLE/FUNGAL PARTNERSHIPS Xyleborus glabratus and Harringtonia lauricola The invasive X. glabratus was first detected in North America in 2002 in Chatham County, Georgia (Table 1.3) (Haack 2006, Rabaglia et al. 2006). Although its association with host trees throughout its large native range in Southeast Asia is not well understood (Cognato et al. 2019), it attacks living Lauraceae trees throughout the southeastern United States (Haack 2006, Harrington et al. 2011, Ploetz et al. 2017). Xyleborus glabratus carries several ophiostomatoid fungi originally assigned to the genus Raffaelea. De Beer et al. (2022) recently described the new genus Harringtonia to accommodate three former Raffaelea species: R. aguacate Simmons Dreaden and Ploetz, R. brunnea Batra (associated with Corthylini ambrosia beetles), and Raffaelea lauricola Harrington Fraedrich and Aghayeva, the causal agent of laurel wilt (Fraedrich et al. 2008, Harrington et al. 2008). Harringtonia lauricola quickly spreads throughout its tree host after inoculation; it prevents water transport by blocking the xylem, causing wilting symptoms within 14 days (Inch et al. 2012). This beetle-phytopathogen partnership likely originated in Asia and was co-imported to the southeastern United States (Harrington et al. 2011) where laurel wilt disease causes widespread damage to forest ecosystems (Ploetz et al. 2017, Wingfield et al. 2017) and commercial avocado (Persea americana Mill.) plantations (Inch et al. 2012). Phylogenetic analysis of the cytochrome c 17 oxidase I gene analysis of X. glabratus across its US population revealed only one haplotype, suggesting that the beetle-fungal pair was introduced once to southern North America (Hughes et al. 2017) followed by rapid proliferation to Georgia and throughout 10 southern states (Olatinwa et al. 2021). Within 20 years, the beetle and fungus advanced west to Texas and north to Kentucky, killing over 300 million trees, especially redbay and sassafras (Hughes et al. 2017, Olatinwa et al. 2021). The beetle and fungus may eventually extend their range as far north as Michigan, given the trend of global warming (Formby et al. 2017). As discussed above, H. lauricola is commonly found in the mycangia of cohabitating xyleborine species and supports survival and reproduction of a closely related Xyleborus. Thus, the fungus may be likely to expand its range by jumping to other beetle partners, especially those with preoral mycangia (Ploetz et al. 2017). Investigators have developed rapid field detection techniques (Abdulridha et al. 2018, Hamilton et al. 2021) and have begun to increase the understanding of the pathogenicity of H. lauricola with molecular screening (Zhou et al. 2020) and metabolic examination (Joseph et al. 2021). However, effective methods for controlling the movement of X. glabratus and other vectors are very limited (Rivera et al. 2020), and there are few effective treatments for trees infected with H. lauricola (Olatinwa et al. 2021). Thus, laurel wilt continues to cause significant ecological and economic damage in the southeastern United States (Hughes et al. 2017). Xylosandrus spp. and Ambrosiella spp. The genus Ambrosiella is one of three lineages in the Microascales family Ceratocystidaceae that independently evolved ambrosial associations with beetles (Mayers et al. 2015). Each of the 11 Ambrosiella species maintain tight associations with one ambrosia beetle species (Miller et al. 18 2019, Mayers et al. 2020b). Due to their long evolutionary history with ambrosia beetles, the genus is highly phenotypically converged which makes morphology-only based identification and taxonomic assignments unreliable (Cassar and Blackwell 1996, Alamouti et al. 2009, Mayers et al. 2020b). Ambrosiella also appears to be well adapted to beetles possessing large, complex mycangia that are associated with internal glands (Harrington et al. 2014, Johnson et al. 2018, Mayers et al. 2020b). The most recent phylogenetic treatment of the genus (Mayers et al. 2020b) identified two clades, one of which has been associated with Scolyplatypodini (Curculionidae: Platypodinae) ambrosia beetles since the origin of Ambrosiella ~18 million years ago. The more derived clade contains the majority of Ambrosiella species and has been carried by several xyleborine genera since its origin ~12 million years ago. The granulate ambrosia beetle (Xylosandrus crassiusculus (Motschilsky)) has a history of human-aided range expansion that stretches back more than a century to Africa (Hagedorn 1908), North America (Anderson 1974), and most recently to Central/South America (Kirkendall and Ødegaard 2007, Flechtmann and Atkinson 2016) and Europe (Table 1.3) (Pennacchio et al. 2003, Nageleisen et al. 2015, Gallego et al. 2017, Kavčič 2018). The ability of X. crassiusculus to establish and spread beyond its native Asia is fueled by two factors that are difficult to disentangle because of the complex genetic diversity this species possesses in both its native and introduced ranges (Ito and Kajimura 2009, Flechtmann and Atkinson 2016, Storer et al. 2017). First, the beetle quickly proliferates after establishing a new population. This is evidenced by the fact that it was one of the most prevalent ambrosia beetle species in the Costa Rica just ten years after its first detection in the country (Kirkendall and Ødegaard 2007). Second, repeated introductions of this species are common, and this increases the rate of success for nascent populations to expands their geographic range (Rabaglia et al. 2019). Both factors are 19 responsible for the rapid proliferation of the granulated ambrosia beetle throughout the North America after its first appearance in Dorchester County, South Carolina, United States in 1974 (Anderson 1974, LaBonte et al. 2005, Flechtmann and Atkinson 2016, Rabaglia et al. 2019), and its range expansion throughout South America and Africa (Flechtmann and Atkinson 2016, Landi et al. 2017, Nel et al. 2020). Xylosandrus crassiusculus maintains an obligate relationship with its fungal symbiont Ambrosiella roeperi T.C. Harr & McNew throughout its global geographic range (Harrington et al. 2014, Mayers et al. 2020b, Nel et al. 2020, Saragih et al. 2021). This fungus is not known to cause disease in host plants; however infestation with X. crassiusculus can be associated with unidentified fungal infections that may opportunistically exploit the holes created by dispersing foundresses (Atkinson et al. 1988). The granulated ambrosia beetle impacts ornamental plant nurseries and lumber productivity by killing young saplings (Kirkendall Ødegaard 2007) and damaging stored logs (Atkinson et al. 2000). It also attacks and reproduces in a vastly large range of angiosperms including food crop trees like coffee (Coffea canephora Pierre ex A. Froehner), cacao (Theobroma cacao L.), mango (Mangifera indica L.), papaya (Carica papaya L.), rubber (Hevea brasiliensis Willd. Ex A. Juss), camphor (Cinnamomum camphora L.), mahogany (Swietenia Jacq. sp.), teak (Tectona grandis L.), and stone fruits (Prunus L. spp.) (Schedl 1963, Atkinson et al. 1988, Wood and Bright 1992). Cavaletto et al. (2021) discovered that host preference of X. crassiusculus was more influenced by ethanol levels in the wood than plant taxon. Harrington et al. (2014) anticipated that X. crassiusculus was unlikely to efficiently vector plant pathogens from Raffaelea, but studies on the lateral transfer of pathogenic fungi between sympatric ambrosia beetles have call this prediction into question. Carrillo et al. (2014) and Ploetz et al. (2017) discussed the potential for the granulated ambrosia beetle to spread of laurel 20 wilt disease after finding its causal agent, H. lauricola, inside the mycangia of X. crassiusculus specimens. Like X. crassiusculus, the black twig borer (Xylosandrus compactus (Eichhoff)) uses a wide variety of dicotyledonous angiosperm hosts (Ngoan et al. 1976, Hara and Beardsley 1979). This ambrosia beetle attacks small-diameter twigs and healthy trees of diameter 3 mm or larger (Wood 1982), causing wilting and death in young individuals (Oliviera et al. 2008). It is a significant pest of coffee, cacao, avocado, mango, and many forest and nursery trees (Brader 1964, Ngoan et al. 1976, Oliveira et al. 2008). The native range of X. compactus probably extends through Asia, although it has been widely introduced to many regions and lives in a pantropical distribution today (Table 1.3) (Brader 1964, Rabaglia et al. 2006, Haack and Rabaglia 2013, Urvois et al. 2022). Brader (1964) noted that X. compactus was limited to the paleotropics but predicted it would eventually spread to the Neotropics. A phylogeographical analysis of global patterns of genetic diversity of X. compactus confirmed that two Asian linages independently invaded Africa and the Americas, Pacific Islands and Europe through international trade (Urvois et al. 2022). The introduction to Africa probably occurred several hundred years ago, early in the history of the continent’s participation in international trade (Egonyu et al. 2015). The occurrence of X. compactus in the Americas, Pacific Islands and Europe is more recent. In North America, it was first recorded in Ft. Lauderdale, Florida in 1941 before quickly establishing pestiferous populations in avocado plantations in Florida and Georgia (Wood 1982). Estimates place the introduction of the black twig borer to South America in the 1970s. It was first collected in Peru in 1973 (Oliviera et al. 2008) and in Brazil during the 1980s (Delgado and Couturier 2017). Xylosandrus compactus was introduced to Oahu, Hawaii in 1961; it has since spread to all the Hawaiian Islands where it 21 threatens coffee crops and several native trees (Hara and Beardsley 1979, Greco and Wright 2015). Most recently, this ambrosia species was discovered in Italy in 2011 (Garonna et al. 2012) then France, Spain, and Greece by 2019 (Faccoli 2021). The population in central Europe causes commercial and ecological damage by attacking bay (Laurus nobilis L.) and carob Ceratonia siliqua L.) trees, producing five generations per year (Gugliuzzo et al. 2020). Xylosandrus compactus is associated with two fungi which lead to several years of confusion as investigators uncovered inconsistent results when culturing samples from the beetles’ gallery walls (Hara and Beardslet 1979). Species from two genera have the most consistent association with X. compactus: Ambrosiella xylebori Brader ex von Arx & Hennebert (Mayers et al. 2015, Bateman et al. 2016, Gugliuzzo et al. 2020) and Fusarium spp. from the solani species complex (FSSC) (Muller 1933, Brown 1954, Ngoan et al. 1976, Gugliuzzo et al. 2020). Bateman et al. (2016) suggested that A. xylebori is likely the nutritional symbiont because they repeatedly recovered it from beetle mycangia and galleries. However, the role (if any) of Fusarium in the life cycle of the black twig borer, is less clear. A pathogenicity study in Uganda found that Fusarium spp. vectored by X. compactus are the causal agent of wilting in infested cacao plants (Kagezi et al. 2017). Bateman et al. (2016) found FSSC was consistently recovered from the external surface of the beetles, but not found in the mycangium. They suggested that FSSC has conidiophores that are adapted for insect dissemination and the pathogenicity of FSSC allows X. compactus to overcome plant defenses when it attacks healthy hosts. These fungi may be prone to developing associations with arthropods because the FSSC contains several insect-vectored plant pathogens (Aoki et al. 2003, Summerell and Leslie 2011) and is vectored by several insect and mite orders (O’Donnell et al. 2012). However, these possible explanations for the association between X. compactus and FSSC 22 require further investigation (Bateman et al. 2016). The beetle is responsible for vectoring a fungal disease that is capable of causing significant economic and ecological damage (Egonyu et al. 2015). Xylosandrus morigerus (Blandford) was described in England from specimens taken from infested orchids originating in Papua New Guinea (Blandford 1894a). Thus, its potential for introduction to many non-native regions was clearly notable from the beginning (Table 1.3) (Browne 1961, Kalshoven 1961). This ambrosia beetle is probably native to tropical Asia, with introduced populations worldwide (Wood and Bright 1992, Kirkendall and Ødegaard 2007, Cognato and Rubinoff 2008, Kirkendall and Faccoli 2010). A study of the mitochondrial genetic diversity of the invasive population in Costa Rica suggested that despite widely divergent mitochondrial lineages, X. morigerus maintains the same host generalism throughout the contry (Andersen et al. 2012). This ecological consistancy may contribute to the extreme mobility of the species outside of its native range. Like X. compactus, X. morigerus attacks twigs and small diameter samplings from a wide variety of angiosperms (Browne 1961, Kalshoven 1961). This can cause significant damage to the coffee, cacao, avocado, mahogany, teak, legumes, and many forest trees (Kalshoven 1961, Carreras-Villaseñor et al. 2022). The mycangial contents of X. morigerus have not yet been studied, but the nutritional symbiont of this beetle is presumably an unstudied Ambrosiella species (Batemena et al. 2018). Two isolates from the FSSC were recovered from X. morigerus collected in Veracruz, Mexico and found to cause wilting and necrosis in coffee, and several forest tree species (Carreras- Villaseñor et al. 2022). The full nature of the relationship between X. morigerus and this pathogenic FSSC is still uncertain. 23 Xylosandrus germanus was described by Blandford (1894b) from 16 specimens collected from the Japanese island of Honshu. It was not collected outside of Asia until Felt (1932) reared several hundred from infested grapevines in Nassau County, New York. Initial biological descriptions of this species noted that it occasionally infested healthy trees but had better success attacking freshly fallen or sick hosts (table 1.3) (Hoffmann 1941). Hoffmann (1941) also observed that X. germanus relied on a fungus for food and concluded that this fungus was unlikely to be a plant pathogen. As the beetle spread through the eastern United States (Weber and McPherson 1982, LaBonte et al. 2005), it was linked to a Fusarium fungal disease responsible for killing young walnut trees in plantations in southern Illinois, Indiana, Missouri, and Iowa (Kessler et al. 1974). It was first reported in Europe in Germany in 1951 (Groschke 1953) and established populations thorough central Europe by the 1990s (Wood and Bright 1992). In 2018, the beetle spread to the Mediterranean scrubland (Contarini et al. 2020). In 2007 X. germanus was discovered in Hawaii (Cognato and Rubinoff 2008). Phylogeographic analysis across its native and introduced ranges indicates that X. germanus populations in Europe and North America each descended from independent invasions by Japanese founders, with Europe having been invaded once, and North America several times (Dzurenko et al. 2021). Non-native populations of X. germanus are so abundant they are a common model for studies about xyleborine sex ratios and inbreeding rates (Peer and Taborsky 2005, Keller et al. 2011). Although X. germanus generally attacks trees that are already physiologically compromised (Ranger et al. 2018) it is a significant pest causing death and dieback in nurseries (Reding et al. 2015) and orchards (Agnello et al. 2017) in the United States. In Europe, it causes significant losses in forests (Grégoire et al. 2003), vineyards (Ruzzier et al. 2021), and the lumber industry (Galko et al. 2018). The mycangial symbiont of X. germanus is Ambrosiella 24 grosmanniae McNew, C. Mayers, and T.C. Harr which is closely related to A. roeperi and A. xylobori and has repeatedly been isolated from specimens in the United States, Germany, Netherlands, and Switzerland (Mayers et al. 2015). Molecular data from the fungi from X. germanus Japan (within the native range) indicated that there may be more genetic diversity than proposed by Mayers et al. (2015) (Ito and Kajimura 2017). Additional fungi, including Fusarium spp., have been recovered from X. germanus galleries (Yang et al. 2008) but their relationship to X. germanus is currently unknown. Disease in New York apple orchards associated with Fusarium sp. is thought to be vectored by X. germanus, but this link remans unconfirmed because specific identification and pathogenicity studies have not been performed (Agnello et al. 2017). Similar to other species of Xylosandrus, X. germanus can live inside an exceptionally wide species of plants including conifers (Weber and McPherson 1983). Multi-year monitoring of non-native X germanus in forests in Slovakia showed that populations did not decline significantly after an exceptionally cold winter, suggesting that they may be capable of continuing to expand into colder areas (Dzurenko et al. 2022). In addition, as global climate change continues to bring milder winters and trigger stress in host trees, this beetle’s range will likely continue to expand throughout its non-native range (Henin and Versteirt 2004). Ambrosia Fusarium Clade and Euwallacea spp. + Coptoborus spp. As discussed above, three Xylosandrus spp. vector FSSC pathogens that cause a significant global impact but are not nutritional symbionts for the beetles (Carreras-Villaseñor et al. 2022). The genera Euwallacea and Coptoborus maintain nutritional relationships with an unusual group of Fusarium species collectively referred to as the Ambrosia Fusarium Clade (AFC) (Li et al. 25 2016, Lynch et al. 2016, Osborn et al. 2022a). These fungi form a monophyletic group that diverged from the FSSC ~21–24 million years ago (Kasson et al. 2013, O’Donnell et al. 2015). The AFC consists of 19 species, 11 of which have been formally described (Gadd and Loos 1947, Nirenberg 1990, Freeman et al. 2013, Na et al. 2018, Aoki et al. 2018, 2019, Lynn et al. 2020, Aoki et al. 2021). Euwallacea fornicatus (Eichhoff) has been a well-documented pest of tea since the late 19th century (Table 1.3) (Walgama 2012). However, there was little examination of its gallery fungus until Gadd and Loos (1947) described Monacrosporium ambrosium Gadd and Loos from beetles infesting tea plantations in Sri Lanka. In 1987, Brayford also noticed the fungus vectored by E. fornicatus in tea in the Indian state of Maharashtra and wrote a duplicate description providing the name Fusarium bugnicourtii (Gadd and Loos). These two names were synonymized three years later (Nirenburg 1990) and there was no further discussion of this ambrosia fungus until 2012 when E. fornicatus was discovered vectoring an undescribed Fusarium causing Fusarium Dieback Disease in avocado in California, United States, and Israel (Eskalen et al. 2012, Mendel et al. 2012). An inclusive survey examining the fungi from worldwide Euwallacea showed that several species including Euwallacea interjectus (Blanford), Euwallacea validus (Eichhoff), E. fornicatus, and Euwallacea sp. cultivate AFC (Kasson et al. 2013). Interestingly, this study also recovered the AFC species AF-9 from Xyleborus ferrugineus (Fabricius) from Costa Rica (Kasson et al. 2031). Testing for co-cladogenesis in Euwallacea and AFC revealed that the AFC are vertically transmitted by the beetles and share a deep and dynamic relationship with Euwallacea that includes frequent partner switching (O’Donnell et al. 2015). The beetles and fungi likely became associated close to the origin of Euwallacea (19–24 26 million years ago). However, the nutritional relationship between the beetles and fungi has not been examined and needs investigation (O’Donnell et al. 2015). Despite the long history of research and monitoring of E. fornicatus, identification of this species based on morphology is quite difficult. The taxonomy of the species has historically been unstable with scientists describing several species with nearly identical diagnoses, placing them into synonymy, and resurrecting some (see Stouthammer et al. 2017 and Gomez et al. 2018). The rediscovery of a syntype from the initial type series of Xyleborus fornicatus Eichhoff provided clarity about the identity of the original description (Smith et al. 2019). However, considerable uncertainty around the diagnosis of E. fornicatus and the correct use of its common name, polyphagous shot hole borer remains (Carrillo et al. 2020a, 2020b). Members of the E. fornicatus species complex spread beyond their native Asia and Oceania into Central America in the 1980s (Wood 1982). Between 2006 and 2018, they spread further to North America (California and Florida, United States) (Rabaglia et al. 2006), Israel (Mendel et al. 2012), and South Africa (Paap et al. 2018). Some AFC fungi associated with Euwallacea spp. cause plant disease (Eskalen et al. 2012, Mendel et al. 2012, Paap et al. 2018), yet pathogenicity among all members of the AFC has not been evaluated. Beetles from the Neotropical genus Coptoborus were linked to a dieback disease affecting balsa (Ochroma pyramidale Cavanilles ex Lamark) plantations in Ecuador in the early 1990s (Stilwell et al. 2014). The beetle was described as Coptoborus ochromactonus Smith and Cognato after its association with dying balsa trees (Table 1.3) (Stilwell et al. 2014). Morphological and molecular analysis both identified the fungus associated with this beetle- vectored disease to be a Fusarium sp. (Stilwell et al. 2014, Castro et al. 2019) and a study of infested balsa plantations in western Ecuador showed that tree age and stress levels had the most 27 influence on infection and mortality rates (Martínez et al. 2020). Osborn et al. (2022a) completed a survey of the fungi in the mycangia of several Coptoborus from Ecuador including C. ochromactonus and concluded that they carry the unnamed species AF-9 from the AFC. Close investigation of the AFC associated with C. ochromactonus is still needed to fulfil Koch’s postulates for disease causality (Smith 1905). Ambrosiodmus minor and Irpex subulatus The sister ambrosia beetle genera Ambrosiodmus and Ambrosiophilus are the only xyleborines with documented symbioses with the basidiomycete fungus I. sublatus (= Flavodon subulatus) (You et al. 2015, Simmons et al. 2016a, Li et al. 2017). The presence of this fungus, which was first assigned to Flavodon and recently was moved into Irpex by Tian et al. (2022), is very uncommon among the Xyleborini. Ambrosius minor (Stebbing) is native to Eastern Asia (Wood and Bright 1992, Lin et al. 2019), but it has been introduced to the southeastern United states, spreading from Nassau County, Florida to Georgia, Alabama, and Mississippi by 2017 (Table 1.3) (Schiefer 2018). The ambrosia partnership between A. minor and I. subulatus could result in unique ecological consequences and challenges as the beetle continues to spread. Irpex sublatus, like other white-rot fungi, can efficiently degrade lignin, one of the most recalcitrant structural molecules that is responsible for the rigidity of plant walls (Eriksson et al. 1990). Because this fungus colonizes dead wood efficiently and may outcompete similar native fungi, it may alter the carbon cycle in forests and increase falling limbs and damage to trees in human environments (Gomez and Hulcr 2020, Jusino et al. 2020). 28 AMBROSIA BEETLE/FUNGAL RESEARCH IN REGIONAL CENTERS OF DIVERSITY There is a greater understanding of introduced xyleborine beetles and their fungi compared to the species residing in their native range. Approximately 1300 xyleborine species are distributed in the tropics but adequate knowledge of taxonomy, ecology, and fungal symbioses is lacking (Wood 2007, Smith et al. 2017, Eliassen and Jordal 2021). Similarly, most studies exploring ambrosia fungus diversity and their relationships to ambrosia beetles focus on the minority of fungi that cause economic or ecological harm (Batra 1967, Alamouti et al. 2009, Dreaden et al. 2014, Hughs et al. 2017, Short et al, 2017). In many ways, this bias is necessary because of the urgency required to mitigate the damage caused by a newly discovered pathogen (Leibold and Kean 2018). However, future-oriented studies aimed at gaining a holistic understanding of the diversity of ambrosia symbioses and the biological mechanisms maintaining them may be the best way to fully understand the ambrosia symbiome and prevent the ecological consequences of waiting until after a new pest presents itself to begin research (e.g., Hulcr and Dunn 2011, Skelton et al. 2018, Miller et al. 2019, Mayers et al. 2020b). Xyleborine diversity is thoroughly documented in North America (Bright 1968, Wood 1982), Europe (Pfeffer 1995), and Asia (Smith et al. 2020). Detailed catalogues also exist for the Xyleborini of Africa (Schedl 1963) and South America (Wood 2007). However, these regions have not been fully explored and probably contain undescribed species and genera (Wood 2007, Smith et al. 2017, Eliassen and Jordal 2021, Osborn et al. 2022b). Despite considerable historical discourse regarding the classification of ambrosia fungi and their connection to the Xyleborini (see Leach et al. 1940, Francke-Grosmann 1967), the identities and ecological impacts of most xyleborine ambrosia fungi are poorly understood or unknown. There are several comprehensive 29 discussions of beetle-fungal interactions, but most focus broadly on Scolytinae (Harrington 2005, Six 2012, Hulcr and Stelinski 2017), or contrast ambrosia and bark beetles with other fungus- farming insects (Farrell et al. 2001, Biedermann and Vega 2020). Norris (1979) reviewed the fungi associated with xyleborines more than four decades ago, but there have been taxonomic changes and new discoveries since then (Figure 1.1). During the last two and a half decades, several phylogenetic studies of the ambrosia fungi from Hypocreales, Ophiostomatales, and Microascales created robust classifications of these groups based on isolates obtained from fungal repositories and molecular sequences from GenBank (Cassar and Blackwell 1996, Jones and Blackwell 1998, Alamouti et al. 2009, Dreaden et al. 2014, Vanderpool 2017 Mayers et al. 2020b, de Beer et al. 2022). The global scope of these is useful for understanding broad evolutionary patterns, but finer understanding of the evolutionary interactions between ambrosia fungi and beetles requires the collection of beetles and isolation of the fungi for comparison across geographic space. Studies accomplishing this have focused on beetles from Asia (Gadd and Loos 1947, Brayford 1987, Gebhardt 2005, Li et al. 2016, Lin et al. 2017, Carrillo et al. 2019, Lynn et al. 2020), North America (Six et al. 2009, Harrington et al. 2008, Eskalen et al. 2013, Harrington et al. 2014, Lynch et al. 2016, Simmons et al. 2016a, Mayers et al. 2017, Aoki et al. 2018), or both (Harrington et al. 2010, Freeman et al. 2013, Simmons et al. 2016b, Na et al. 2018, Aoki et al. 2021). A few included specimens from Australia (von Arx and Hennebert 1965, Kasson et al. 2013, Aoki et al. 2019), Europe (Nirenberg 1990, Mayers et al. 2015), and Africa (von Arx and Hennebert 1965, Scott and du Toit 1970), but the xyleborine ambrosia fungi from these areas are sparsely studied. The fungi living with xyleborines native to South America are virtually unknown despite a few studies 30 exploring one endemic genus from Ecuador (Stilwell et al. 2014, Castro et al. 2019, Osborn et al. 2022a). ENHANCING GLOBAL RESEARCH OF AMBROSIA SYMBIOSIS. The nature of the xyleborine ambrosia beetle-fungus relationship remains poorly understood because of varying levels of fidelity/promiscuity, evolutionary histories, and taxonomic diversity of the fungi. Yet xyleborines are found globally and are common invaders in every biogeographic region. They cause harm through mass-aggregation on weakened trees and structural damage to wood, but they are the most ecologically and economically impactful when they inoculate host trees with pathogenic fungi (Hulcr and Stelinski 2017). The recent formation of the Bark Beetle Mycobiome collaborative group (Hulcr et al. 2020) and its framework for standardizing methods and communication is a crucial first step towards studying and combating harmful ambrosia beetles as a global community rather than as individual countries or regions. Invasive ambrosia beetles are monitored and studied by well- established research programs in several regions, but there are parts the world that currently lack such sustained efforts. Many of these are developing nations located around the equator where there is the richest diversity of xyleborine beetles. These countries may be the source of future invasive populations as well as suffer from the consequences of new introductions, yet they rarely have robust and enduring programs for studying the ambrosia symbiosis and monitoring for non-native species. To better strengthen all regions against harmful new invasive xyleborines and their possibly destructive fungi, research needs to be supported equitably around the world to enhance understudied areas. The founding consortium of researchers that crafted the Bark Beetle 31 Mycobiome includes an impressive group from institutions in the United States and South Africa. To maximize its potential, the collaborative relationships codified in its foundation, as cited in this review, should be expanded to benefit research programs in Central and South America, Oceania, Asia, and Sub-Saharan Africa (apart from South Africa) should be developed and supported. Partnerships between traditionally privileged nations and these underrepresented regions would be mutually beneficial because local knowledge, infrastructure and ingenuity can be harnessed to gain knowledge that benefits the global community. 32 APPENDIX 33 Genus species Common Name, Notes Origin Region(s) Ambrosiodmus lewisi Asia punky wood ambrosia Ambrosiodmus minor beetle Asia Ambrosiodmus rubricollis Asia/East Asia Ambrosiophilus aratus Asia/East Asia Ambrosiophilus nodulosus Asia Anisandrus dispar Europe Anisandrus maiche Asia Cnestus mutilatus camphor shot borer Asia Coptoborus coartatus South America Coptoborus crinitulus South America Coptoborus ricini South America Coptoborus villosulus (= theobromae) South America Cyclorhipidion bodoanum (= californicus) North Asia Cyclorhipidion fukiense Asia Cyclorhipidion pelliculosum Asia Dryocoetoides cristatus South America Dryoxylon onoharaense Asia polyphagous shothole Euwallacea fornicatus borer Asia Euwallacea interjectus Asia Euwallacea similis Africa, Asia Euwallacea validus Asia Xyleborinus alni Asia, Europe Xyleborinus andrewesi Asia Xyleborinus attenuatus East Asia Xyleborinus exiguus Asia Xyleborinus octiesdentatus Asia Xyleborinus saxesenii Asia, Europe Table 1.1: Introduced Xyleborini, their native region, and region(s) of introduction. Data inferred from Wood and Bright 1992, Pennacchio et al. 2003, Rabaglia et al. 2006, Kirkendall and Ødegaard 2007, Wood 2007, Cognato and Rubinoff 2008, Kirkendall and Faccoli 2010, Haack and Rabaglia 2013, Gomez et al. 2018, Schiefer 2018 Lin et al. 2021, and Urvois et al. 2022. 34 Table 1.1 (cont’d) Genus species Common Name, Notes Origin Region(s) Ambrosiodmus lewisi Asia punky wood ambrosia Ambrosiodmus minor beetle Asia Ambrosiodmus rubricollis Asia/East Asia Ambrosiophilus aratus Asia/East Asia Ambrosiophilus nodulosus Asia Anisandrus dispar Europe Anisandrus maiche Asia Cnestus mutilatus camphor shot borer Asia Coptoborus coartatus South America Coptoborus crinitulus South America Coptoborus ricini South America Coptoborus villosulus (= theobromae) South America Cyclorhipidion bodoanum (= californicus) North Asia Cyclorhipidion fukiense Asia Cyclorhipidion pelliculosum Asia Dryocoetoides cristatus South America Dryoxylon onoharaense Asia polyphagous shothole Euwallacea fornicatus borer Asia Euwallacea interjectus Asia Euwallacea similis Africa, Asia Euwallacea validus Asia Xyleborinus alni Asia, Europe Xyleborinus andrewesi Asia Xyleborinus attenuatus East Asia Xyleborinus exiguus Asia Xyleborinus octiesdentatus Asia Xyleborinus saxesenii Asia, Europe 35 Table 1.1 (cont’d) Genus species Introduced Region(s) Ambrosiodmus lewisi North America (USA) Ambrosiodmus minor North America (USA) Ambrosiodmus rubricollis Australia, Europe (Italy), North America (Mexico, USA) Ambrosiophilu s aratus Europe (Italy), North America (USA) Ambrosiophilu s nodulosus North America (USA) Anisandrus dispar North America (USA) Anisandrus maiche North America (USA) Cnestus mutilatus North America (USA) Coptoborus coartatus Africa Coptoborus crinitulus Africa Coptoborus ricini Africa Coptoborus villosulus Africa Cyclorhipidion bodoanum Europe, North America (USA) Cyclorhipidion fukiense North America (USA) Cyclorhipidion pelliculosum North America (USA) Dryocoetoides cristatus Africa Dryoxylon onoharaense North America (USA) Australia, Central America, Hawaii, North America (USA), Euwallacea fornicatus South America (Brazil) Euwallacea interjectus North America (USA) Euwallacea similis North America (USA), South America (Brazil) Euwallacea validus North America Xyleborinus alni North America Xyleborinus andrewesi Hawaii, North America (USA) Xyleborinus attenuatus Europe (Austria), North America Xyleborinus exiguus Central America Xyleborinus octiesdentatus North America (USA) Africa, Australia, Hawaii, New Zealand, North America Xyleborinus saxesenii (USA), South America 36 Table 1.1 (cont’d) Genus species Introduced Region(s) Xyleborus abberrans South America (Brazil) Xyleborus affinis Africa, Asia, Australia, Europe, Hawaii, North America (USA) Xyleborus ferrugineus Africa, Asia Australia, Hawaii Xyleborus glabratus North America (USA) Xyleborus pfeilii Europe, New Zealand, North Amercia Xyleborus seriatus North America (USA) Xyleborus spinulosus North America (USA), Hawaii Xyleborus xylographus Asia Xylosandru s amputatus North America (USA) Xylosandru Africa, Central America, Europe, Hawaii, New Zealand, North s compactus America (USA), South America Xylosandru crassiusculu Africa, Australia, Central America, Europe (Italy), Hawaii, North s s America, South America Xylosandru s germanus Europe, North America Xylosandru Africa, Central America, Europe, Hawaii, Middle East (Jordon, s morigerus Lebanon), North America, Pacific Islands, South America 37 Genus species Beetle(s) Ambrosiella batrae Anisandrus sayi Ambrosiella beaveri Cnestus mutilatus Ambrosiella catenulata Eccoptopterus spp., Hadrodemius spp. Ambrosiella cleistominuta Anisandrus maiche Ambrosiella grosmanniae Xylosandrus germanus Ambrosiella hartigii Anisandrus dispar Ambrosiella nakashimae Xylosandrus amputatus Ambrosiella roeperi Xylosandrus crassiusculus Ambrosiella xylebori Xylosandrus compactus Dryadomyces amasae Amasa concitatus, Amasa aff. glaber Dryadomyces sulphureus Xyleborinus saxexenii Fusarium AF-6 Euwallacea sp. AF-8 Fusarium duplospermum Euwallacea sp. Fusarium AF-9 Coptoborus spp. Xyleborus ferrugineus AF-10 Fusarium drepaniforme Euwallacea fornicatus Fusarium AF-11 papillatum Euwallacea sp. from Taiwan Fusarium AF-13 Euwallacea sp. from Taiwan Fusarium AF-14 Euwallacea sp. from Taiwan Fusarium AF-15 Euwallacea sp. from Taiwan Fusarium AF-16 Euwallacea sp. from Taiwan Fusarium AF-17 Euwallacea sp. Fusarium AF-18 Euwallacea sp. Table 1.2: Ambrosia fungal species associated with xyleborine beetles and their beetle partners. Inferred from von Arx and Hennebert 1965, Batra 1967, Scott and du Toit 1970, Brayford 1987, Gebhardt 2005, Harrington et al. 2008, Six et al. 2009, Harrington et al. 2010, Kasson et al. 2013, Mayers et al. 2015, O’Donnell et al. 2015, Simmons et al. 2016a, 2016b, Lin et al. 2017, Mayers et al. 2017, Na et al. 2018, Carrillo et al. 2019, Lynn et al. 2020, Nel et al. 2021, and Osborn et al. 2022a. 38 Table 1.2 (cont’d) Genus species Beetle(s) Harringtonia lauricola Xyleborus glabratus Irpex subulatus Ambrosiodmus minor Neocosmospora AF-1 ambrosia Euwallacea fornicatus AF-2 Neocosmospora euwallaceae Euwallacea sp. Neocosmospora AF-3 floridana Euwallacea interjectus Neocosmospora AF-4 oligoseptata Euwallacea validus AF-7 Neocosmospora obliquiseptata Euwallacea sp. Neocosmospora AF-12 kuroshio Euwallacea sp. Neocosmospora AF-19 rekana Euwallacea perbrevis Raffaelea arxii Xyleborus vovulus (= X. torquatus) Raffaelea campbelliorum Xyleborus glabratus Raffaelea cyclorhipidii Cyclorhipidion ohnoi Raffaelea ellipticospora Xyleborus glabratus Raffaelea fusca Xyleborus glabratus Raffaelea promiscua Xyleborinus saxesenii Raffaelea subalba Xyleborus glabratus Raffaelea subfusca Xyleborus glabratus Raffaelea xyleborini Xyleborinus andrewsii 39 Table 1.2 (cont’d) Genus species Notes Ambrosiella batrae Ambrosiella beaveri Ambrosiella catenulata Ambrosiella cleistominuta Ambrosiella grosmanniae Ambrosiella hartigii Ambrosiella nakashimae Ambrosiella roeperi Ambrosiella xylebori Dryadomyces amasae Dryadomyces sulphureus = Raffaelea sulphurea Fusarium AF-6 Fusarium AF-8 duplospermum Fusarium AF-9 AF-10 = Fusarium Fusarium drepaniforme bugnicourtii Fusarium AF-11 papillatum Fusarium AF-13 Fusarium AF-14 Fusarium AF-15 Fusarium AF-16 Fusarium AF-17 Fusarium AF-18 40 Table 1.2 (cont’d) Genus species Notes Harringtonia lauricola = Raffaelea lauricola Irpex subulatus = Flavodon ambrosius Neocosmospora AF-1 ambrosia = Fusarium ambrosium AF-2 Neocosmospora euwallaceae = Fusarium euwallaceae Neocosmospora AF-3 floridana = Fusarium floridanum Neocosmospora AF-4 oligoseptata = Fusarium oligoseptatul AF-7 = Fusarium Neocosmospora obliquiseptata obliquiseptatum Neocosmospora AF-12 kuroshio = Fusarium kuroshium Neocosmospora AF-19 rekana = Fusarium rekanum Raffaelea arxii Raffaelea campbelliorum Raffaelea cyclorhipidii Raffaelea ellipticospora Raffaelea fusca Raffaelea promiscua Raffaelea subalba Raffaelea subfusca Raffaelea xyleborini 41 Species Nutrutional symbiont Pathogenic fungus Asia Harringtonia Xyleborus glabratus Harringtonia lauricola lauricola Native Xylosandrus crassiusculus Ambrosiella roeperi … Native Xylosandrus compactus Ambrosiella xylebori Fusarium solani sp. Native Xylosandrus morigerus Ambrosiella sp. Fusarium solani sp. Native Xylosandrus germanus Ambrosiella grosmanniae Fusarium solani sp. Native Euwallacea fornicatus spp. Fusarium spp. (AFC) Fusarium spp. (AFC) Native Coptoborus ochromactonus Fusarium sp. AF-9 Fusarium sp. AF-9 N/A Ambrosiodmus minor Irpex Subulatus … Native Table 1.3: Harmful Xyleborini, their nutritional symbionts, associated pathogenic fungi, native region, year of first report in introduced regions, and significant plant hosts. Inferred from Blanford 1894a, Hagedorn 1908, Hoffmann 1941, Groschke 1953, Anderson 1974, Kessler et al. 1974, Hara and Beardsley 1979, Wood 1982, Weber and McPherson 1983, Nirenburg 1990, Pennacchio et al. 2003, Haack 2006, Rabaglia et al. 2006, Kirkendall and Ødegaard 2007, Cognato and Rubinoff 2008, Olivera et al. 2008, Eskalen et al. 2012, Garonna et al. 2012, Mendel et al. 2012, Stilwell et al. 2014, Egonyu et al. 2015, Mayers et al. 2015, Nageleisen et al. 2015, You et al. 2015, Flechtmann and Atkinson 2016, Simmons et al. 2016a, Gallego et al. 2017, Li et al. 2017, Kavčič 2018, Paap et al. 2018, Schiefer 2018, Carreras-Villaseñor et al. 2022, and Osborn et al. 2022a. 42 Table 1.3 (cont’d) North Species Africa Europe America Xyleborus glabratus 2002 N/A N/A Xylosandrus crassiusculus 1700s? 1700s? 2003 Xylosandrus compactus 1941 1700s? 2011 Xylosandrus morigerus unknown unknown unknown Xylosandrus germanus 1941 N/A 1951 Euwallacea fornicatus spp. 2006 2018 Native Coptoborus ochromactonus N/A N/A N/A Ambrosiodmus minor 2017 N/A N/A 43 Table 1.3 (cont’d) Central/South Species Oceania Australia America Xyleborus glabratus N/A N/A N/A Xylosandrus crassiusculus 1996 N/A N/A 1961 Xylosandrus compactus 1970s N/A (Hawaii) Xylosandrus morigerus unknown Native N/A 2007 Xylosandrus germanus N/A (Hawaii) Euwallacea fornicatus spp. 1980s 1980s Native Coptoborus ochromactonus Native N/A N/A Ambrosiodmus minor N/A N/A N/A 44 Table 1.3 (cont’d) Species Significant hosts Xyleborus glabratus Lauraceae, Persea Xylosandrus Carica, Cinnamomum, Coffea, Hevea, Magnifera, Prunus, crassiusculus Swietenia, Tectona, Theobroma Xylosandrus compactus Ceratonia, Coffea, Laurus, Magnifera, Persea, Theobroma, Xylosandrus morigerus Coffea, Cacao, Fabaceae, Persea, Swietenia, Tectona Xylosandrus germanus Malus, Pinus, Vitis Euwallacea fornicatus spp. Persea Coptoborus ochromactonus Ochroma Ambrosiodmus minor … 45 Figure 1.1: Schematic representation of ambrosia associations between xyleborine and fungal genera. 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Applied Microbiology and Biotechnology. 104: 7331–7343. 65 CHAPTER 2: Ecuadorian Coptoborus beetles harbor Fusarium and Graphium fungi previously associated with Euwallacea ambrosia beetles ABSTRACT Ambrosia beetles from the scolytine tribe Xyleborini (Curculionidae) are important to the decomposition of woody plant material on every continent except Antarctica. These insects farm fungi on the walls of tunnels they build inside recently dead trees and rely on the fungi for nutrition during all stages of their lives. Such ambrosia fungi rely on the beetles to provide appropriate substrates and environmental conditions for growth. A small minority of xyleborine ambrosia beetle/fungi partnerships cause significant damage to healthy trees. The xyleborine beetle Coptoborus ochromactonus vectors a Fusarium (Hypocreales) fungus that is lethal to balsa (Ochroma pyramidale (Malvaceae)) trees in Ecuador. Although this pathogenic fungus and its associated beetle are not known to be established in the United States, several other non- native ambrosia beetle species are vectors of destructive plant diseases in this country. This fact and the acceleration of trade between South America and the United States demonstrate the importance of understanding fungal plant pathogens before they escape their native ranges. Here we identify the fungi accompanying Coptoborus ambrosia beetles collected in Ecuador. Classification based ribosomal internal transcribed spacer 1 (ITS) sequences revealed the most prevalent fungi associated with Coptoborus are Fusarium spp. and Graphium spp. (Microascales: Microascaceae), which have been confirmed as ambrosia fungi for xyleborine ambrosia beetles, and Clonostsachys sp. (Hypocreales), which is a diverse genus found abundantly in soils and associated with plants. Phylogenetic analyses of the Fusarium strains 66 based on ITS, translation elongation factor (EF1-a), and two subunits of the DNA-directed RNA polymerase II (RPB1 and RPB2) identified them as Fusarium. sp. AF-9 in the Ambrosia Fusarium Clade (AFC). This Fusarium species was previously associated with a few xyleborine ambrosia beetles, most notably the species complex Euwallacea fornicatus (Eichhoff 1868) (Curculionidae: Scolytinae: Xyleborini). Examination of ITS and EF1-a sequences showed a close affinity between the Graphium isolated from Coptoborus spp. and other xyleborine- associated Graphium as well as the soil fungus Graphium basitruncatum. This characterization of ambrosia fungi through DNA sequencing confirms the identity of a putative plant pathogen spread by Coptoborus beetles and expands the documented range of Fusarium and Graphium ambrosia fungi. INTRODUCTION The weevil subfamily Scolytinae includes bark beetles and ambrosia beetles that are ecologically important in decomposing dead and dying trees and shrubs worldwide (Hofstetter et al. 2015). Bark and ambrosia beetles share an evolutionary history, although they inhabit different ecological niches (Jordal and Cognato 2012, Johnson et al. 2018). Bark beetles excavate tunnels (galleries) underneath and parallel to the bark of their host, feed on phloem, and facultatively interact with fungi that already exist in their environment (Wood 2007, Hulcr et al. 2015). Ambrosia beetles carry symbiotic fungi in specialized cuticular organs known as mycangia. They bore into the xylem of their host, then inoculate the gallery walls with symbiotic fungi that provide nutrition throughout the entire life cycle of the beetles (Hubbard 1897, Hofstetter et al. 2015). The bark beetle lifestyle is ancestral within Scolytinae, but ambrosia feeding evolved 12– 16 independent times (Johnson et al. 2018) with several different fungal lineages. 67 Ambrosia fungi provide three essential services to their beetle partners. They extract carbon and nutrients from the xylem substrate and pass them to the beetles (Bleiker and Six 2007, Six 2012, Hofstetter et al. 2015). As they spread and decompose the wood, the fungi probably protect the beetles from plant defensive responses (Klepzig et al. 2001, Six 2003, Hofstetter et al. 2015), and the invasion of less desirable fungi in the gallery (Klepzig 2006). The saprotrophic action of ambrosia fungi may also soften the wood, allowing the beetles to expand their galleries more easily (Barta 1967). In return, ambrosia beetles disperse the fungi and provide a protected habitat inside the xylem with optimal humidity (Batra 1967). The nature and diversity of the ambrosia symbiosis has always been enigmatic, because ambrosia beetles are small and live most of their lives within their galleries where they cannot be observed easily. Ambrosia fungi are often highly adapted to life inside beetle tunnels (Francke- Grosmann 1967, Malloch and Blackwell 1993, Cassar and Blackwell 1996, Jones and Blackwell 1998), and convergence challenges their morphological identification and description (Francke- Grossman 1967, Alamouti et al. 2009). The relationships they share inside the beetle gallery are complex, often involving nematodes, protozoa, mites, other filamentous fungi, yeasts and bacteria (Hofstetter et al. 2015). Ambrosia beetles and their fungi generally inhabit dead or dying plants and thus are considered harmless. However, a handful of ambrosia beetles vector destructive fungi capable of killing otherwise healthy trees. These partnerships cause serious ecological and economic damage to forest, urban and ornamental trees (Hubbard 1897, Smith and Hulcr 2015). Some of the most impactful vectors are exotic xyleborine ambrosia beetles. The invasive beetle species complex Euwallacea fornicatus (Eichhoff 1868), which originated in Asia, was first documented in the southeastern United States in 2002 (Haack 2006). 68 This group, which contains seven morphologically cryptic beetle species (Smith et al. 2019) was introduced multiple times in North and Central America (Wood and Bright 1992, Haack 2006, Rabaglia et al. 2006, Kirkendall and Ødegaard 2007). Like fungi associated with other invasive xyleborines, the mycangial partners of the Euwallacea fornicatus species group are diverse and include fungi from the nectrioid genera Paracremonium and Fusarium (Hypocreales), and Graphium (Microascales: Microascaceae) (Freeman et al. 2013, 2016, Li et al. 2016, Lynch et al. 2016, O’Donnell et al. 2016, Carrillo et al. 2020a). Many of the species in the genus Graphium are plant pathogens, including some that exploit physically damaged trees (Musvuugwa et al. 2020). Several Graphium species disperse with scolytine beetles, including many bark beetles (Wingfield and Gibbs 1991, Jacobs et al. 2003, Okada et al. 2003, Malacrinó et al. 2017). Baker and Norris (1968) recovered Graphium from the mycangium of the xyleborine beetle Xyleborus ferrugineus (F.), although they never confirmed that it acted as a nutritional symbiont (Kok 1979). Recent work established two Graphium species as ambrosia fungi associated with the E. fornicatus species complex: Graphium euwallaceae Twizeyimana, Lynch & Eskalen and Graphium kuroshium Na, Carrillo & Eskalen (Freeman et al. 2016, Lynch et al. 2016, Na et al. 2018, Carrillo et al. 2020a, 2020b). Interestingly, both Graphium ambrosia fungi (as well as the undescribed species associated with X. ferrugineus) coexist with Fusarium (Baker and Norris 1968, Freeman et al. 2016, Lynch et al. 2016). The genus Fusarium is diverse in species and ecology. Although the taxonomic status of many Fusarium species is still poorly understood (Summerell et al. 2010, Crous et al. 2021), several species complexes cause disease including in numerous important grain and food crops (Aoki et al. 2003, Ploetz 2006, Saremi et al. 2008, Arie 2019), trees (Gardner 1980, Stilwell et al. 69 2014), humans and other animals (O’Donnell et al. 2012, Smyth et al. 2019). Some species in the Fusarium solani (Mart.) Sacc. species complex are vectored by Euwallacea ambrosia beetles (see Aoki et al. 2021 and references therein). These fungi form a monophyletic lineage and are collectively referred to as the Ambrosia Fusarium Clade (AFC) (Kasson et al. 2013, O’Donnell et al. 2015). As with other Fusarium species, some AFC isolates are phytopathogens, causing damage to avocado and forest trees in California and Israel (Eskalen et al. 2013, Kasson et al. 2013, Cognato et al. 2015, Lynch et al. 2016, Short et al. 2017). Researchers examining the AFC uncovered evidence of a longstanding cooperative relationship with Euwallacea beetles beginning 19–24 MA (O’Donnell et al. 2015). The origin of the AFC (~21–24 MA) roughly coincides with the origin of their xyleborine beetle associates (~19 MA) (Jordal et al. 2000, Kasson et al. 2013, O’Donnell et al. 2015). Several AFC species also produce large, club-shaped conidia, presumably an adaptation to their nutritional partnership with beetles (O’Donnell et al. 2015). However, the relationship between the fungi and their beetle hosts appears to be promiscuous, with at least five host-switching events over 19–24 MA (O’Donnell et al. 2015). Furthermore, there is evidence that species from other xyleborine genera such as Xyleborus ferrugineus (Baker and Norris 1968, Norris and Baker 1968) and Xyleborinus saxesenii (Ratzeburg 1837) (Malacrinó et al. 2017) associate with Fusarium. The most destructive xyleborine-fungal partnerships originated in Asia and were imported to the Americas (Haack 2016). Nevertheless, potentially threatening beetle-ambrosia pairings from Central and South America deserve attention (Liebhold and Kean 2018, Stillwell et al. 2014). In the early 1990’s the xyleborine Coptoborus ochromactonus Smith & Cognato (Stilwell et al. 2014) was discovered to vector an undescribed Fusarium pathogen that causes a wilting disease that can kill up to 90% of infected balsa trees (Ochroma pyramidale (Cav. Ex 70 Lam.) Urb.) in Ecuador (Stilwell et al. 2014, Castro et al. 2019). This newly described beetle is a member of a recently revised genus native to the Neotropics comprising 77 species (Smith and Cognato 2021). The identities of Coptoborus-associated fungi are mostly unknown (Wood 2007, Hulcr et al. 2015). The location of the mycangium of Coptoborus spp. has not been confirmed (Castro et al. 2019), although examination of the mycangial diversity within Xyleborini suggests that mandibular mycangia (hereafter referred to as preoral mycangia after Li et al. 2015) are the ancestral state for the tribe (Schedl, 1962, Francke-Grosmann 1967, Cognato et al. 2011). Additionally, phylogenetic analysis indicates that the genus shares a close evolutionary relationship with Xyleborus, which have preoral mycangia (Cognato et al. 2011). Given the damage caused by C. ochromactonus to the Ecuadorian balsa industry and the increasing ecological homogenization (Fisher et al. 2012) from mounting international trade and global climate change (Liebhold and Kean 2018, Aukema et al. 2010, Marini et al. 2011, Rabaglia et al. 2019), further investigation of fungi associated with South American xyleborines has become increasingly relevant. To complement the increasing knowledge of Neotropical xyleborine diversity (Wood 2007, Smith et al. 2017, Bright 2019, Smith and Cognato 2021), this paper identifies and phylogenetically resolves the fungi associated with six Ecuadorian Coptoborus species. METHODS Material collection Live ambrosia beetles were collected from dead and dying trees and woody plant material at the Yasuní Research Station (Orellana Province, Ecuador, –0.661111, –76.400389) between January 71 and March 2018. Specimens from each wood sample were sorted to morphospecies, based on characters of the antennae, protibiae, and declivital sculpturing. They were kept alive on damp Kimwipe tissues in 50 ml vials and transported to Pontifica Universidad Católica del Ecuador in Quito. Depending on availability, up to five specimens per morphospecies were selected for fungal isolation, for a total of 24 beetle specimens. Each individual was surface-sanitized by vortexing for 15 s in sterile phosphate-buffered saline (PBS). To isolate the preoral mycangia, a sterile scalpel was used to separate the head of each beetle from the pronotum. Each head was pulverized with a sterile micropestle and combined with 500 µL of sterile PBS. Fifty µL of this solution was transferred into 500 µL of sterile PBS to create a 9x dilution. The remainder of the body of each specimen was preserved in 100% ethanol and vouchered in The A.J. Cook Arthropod Research Collection at Michigan State University. Fifty µL each from the 1x solution and the 9x dilution were transferred to Petri dishes filled with potato dextrose agar (PDA) fortified with 1.6% agar, 0.2% yeast extract, and 1% Penicillin-Streptomycin antibiotics (Life Technologies; Waltham, Massachusetts). The plated mycangial samples were incubated at 23–25 ºC in the dark and checked for growth as over the course of ten days. Colonies on the plates were visually examined to characterize them into morphotypes and the growing edge of one representative from each morphotype was transferred to PDA in a Petri dish to obtain pure cultures. After incubating the pure cultures for up to five days, plugs with fungal growth were preserved in 100% ethanol for subsequent use. 72 Molecular data DNA was extracted from the fungal samples with a Qiagen plant Mini Kit (Hilden, Germany) according to the manufacturer’s instructions. The ribosomal internal transcribed spacer 1 (ITS) (Schoch et al. 2012) from each isolate was amplified and sequenced for preliminary identification to genus with the nucleotide BLAST search function of the National Center for Biotechnology Information (NCBI) GenBank database (Altschul et al. 1990). Twenty-six fungal samples were identified as the ambrosia lineages Fusarium or Graphium (Table 2.1). We amplified and sequenced two additional loci from these isolates based on their known phylogenetic utility: translation elongation factor (EF1-a), and the second largest subunit of DNA-directed RNA polymerase II (RPB2) (Mendel et al. 2012, O’Donnell et al. 2012, Eskalen et al. 2013, Freeman et al. 2013, Kasson et al. 2013, O’Donnell et al. 2015). PCR primers and protocols followed Jacobs et al. (2004), Lynch et al. (2016), O’Donnell et al. (1998, 2007, 2010), and White et al. (1990) (Tables 2.2 and 2.3). Previous work has also demonstrated the phylogenetic utility of DNA-directed RNA polymerase I (RPB1) (O’Donnell et al. 2012, Freeman et al. 2013, Kasson et al. 2013, O’Donnell et al. 2015), thus we included this locus in our data set when data were available from GenBank (Table 2.4). Repeated attempts to amplify this locus from our Ecuadorian fungal isolates were not successful, however, inclusion of RPB1 data from additional Fusarium strains from GenBank added resolution and branch support to our analyses. Thus, we incorporated RPB1 data into the phylogenetic analysis for Fusarium, but not for Graphium. The total volume of each PCR cocktail for all loci was 25 µL per 50–5 ng DNA template and included 1x buffer, 1.75 mM MgCl2, 200 µM dNTPs, 0.3 µM each forward and reverse primers, and 1.25 units HotStar Taq (Qiagen) (See Table 2.3 for PCR conditions). PCR products were readied for cycle sequencing 73 with Exo-SAP-IT (Applied Biosystems; Waltham, Massachusetts) and sequenced with BIGDYE TERMINATOR 1.1 (Applied Biosystems) at the Michigan State University Research Technology Support Facility. Opposing strands were compiled, trimmed, and edited to correct ambiguous or incorrect base judgements with SEQUENCHER 5.0 BUILD 7082 (Gene Codes Corporation; Ann Arbor, Michigan). Completed sequences were submitted to GenBank (see Table 2.1 for accession numbers). Sequence alignments for both Fusarium spp. and Graphium spp. were created using the default settings of MUSCLE (Madeira et al. 2019). Phylogenetic analyses To identify the putative ambrosia fungi to species, the data from Fusarium spp. and Graphium spp. were each subjected to independent phylogenetic analyses on each locus and on combined datasets. Loci used for Fusarium were ITS, EF1-a, RPB1, and RPB2. Those used for Graphium were ITS, and EF1-a. Both datasets were also supplemented with corresponding data from GenBank (Table 2.4). Data from Fusarium isolates included 17 AFC fungal samples from Yasuní and molecular data from similar ambrosia fungi obtained from GenBank. The additional sequences are GenBank data from 95 known ambrosia strains isolated from xyleborines or wood infested with xyleborines (Mendel et al. 2012, Kasson et al. 2013, O’Donnell et al. 2015, Na et al. 2018, Carrillo et al. 2020b). We also included three representative strains from non-ambrosia Fusarium species (Zhang et al. 2006, O’Donnell et al. 2007, O’Donnell et al. 2010, Kasson et al. 2013). Fusarium neocosmosporiellum O’Donnell and Geiser was used to root the trees based on previous phylogenies of the AFC (see Kasson et al. 2013, Lynn et al. 2020). 74 We evaluated the combined Fusarium dataset using PARTITIONFINDER 2.1.1 to establish appropriate partitions within the data and to choose the best-fit model of nucleotide substitution for each (Guindon et al. 2010, Lanfear et al. 2016). An exhaustive search (using the ‘all’ search algorithm) of seven data blocks (a single data block for the non-coding ITS gene, and the 1st, 2nd and 3rd codon positions of EF1-a, RPB1, and RPB2) established seven partitions selected based on the corrected Akaike information criterion (AICc). The ITS, EF1-a genes, and a subset that combines the third codon positions from RPB1 and RPB2 were each assigned to a separate general time-reversible model with variable nucleotide frequencies, and gamma distributed rate of variation (GTR+I+G) (Lanave et al. 1984, Rodríguez et al. 1990). A general time-reversible model with variable nucleotide frequencies (GTR+I) (Lanave et al. 1984, Rodríguez et al. 1990) fit a combined subset of the first codon positions of RPB1 and from RPB2. The second codon position of RPB1 was fit to the general time-reversible model with a gamma distribution (GTR+G) (Lanave et al. 1984, Rodríguez et al. 1990). The second codon position of RPB2 fit the Hasegawa-Kishino model with variable nucleotide frequencies and a gamma distribution (HKY+I+G) (Hasegawa et al. 1985). Sequence data of ITS, and EF1-a, from the nine Graphium isolates were concatenated with data from the same two loci belonging to 65 Graphium ambrosia strains found in GenBank (Table 2.4). Thirty-nine of these were isolated from wood infested by scolytine beetles (Cruywagen et al. 2010, Lynch et al. 2016, Na et al. 2018), 16 were isolated from xyleborine beetles (Na et al. 2018, Carrillo et al. 2020b), seven were isolated from non-xyleborine Scolytinae (Jacobs et al. 2003, Hulcr et al. 2007, Cruywagen et al. 2010), and three were isolated from Pissodes weevils (Molytinae) (Cruywagen et al. 2010, Paciura et al. 2010). We also included a three-member outgroup consisting of the non-ambrosia species Graphium 75 basitruncatum (Matsush.) Seifert & G. Okada, Ophiostoma ulmi (Buisman) Nannf., and Geosmithia putterillii (Thom) Pitt (formerly G. pallida (G. Sm.) M. Kolarík, Kubátová & Pazoutová) after previous work concerning ambrosia-beetle-associated Graphium (Kolarík and Hulcr 2008, Cruywagen et al. 2010, Lynch et al. 2016, Na et al. 2018, Carrillo et al. 2020b). The combined Graphium dataset was divided into seven data blocks using PARTITIONFINDER 2.1.1 (Guindon et al. 2010, Lanfear et al. 2016). The non-coding gene ITS was assigned to a single block, whereas the coding genes EF1-a was divided into three blocks containing the first, second and third codon positions, respectively. A complete search of these data blocks identified five subsets for partitioning. ITS and the second codon position of EF1-a each fit a separate general time-reversible model with a gamma distribution (GTR+G) (Lanave et al. 1984, Rodríguez et al. 1990). The first and third codon positions from EF1-a best fit was Hasegawa-Kishino with gamma distributed rate of variation (HKY+G) (Hasegawa et al. 1985). We imported the model schemes for Fusarium and Graphium into MRBAYES 3.2.5 (Ronquist and Huelsenbeck 2003) and used them to search the treespace of each combined dataset with Monte Carlo Markov chains. Both analyses used identical parameters including two independent runs, each consisting of three heated chains and one cold chain (metropolis coupling), searching across 40 million generations using Monte Carlo rules for tree acceptance or rejection at each generation. We sampled the tree at every 100th generation and discarded the first 25% (100,000 trees) as burn-in before using the remaining 300,000 trees to build a majority- rule consensus tree. The data from both genera were also subjected to independent heuristic searches using PAUP* 4.0A BUILD 167 (Swofford 2002) to search for most parsimonious trees. Gaps were treated as missing. Characters were unordered and equally weighted during the heuristic search with 76 2000 repetitions of random stepwise addition of taxa using tree bisection/reconnection. Bootstrap support was calculated using the same character parameters and 2000 full heuristic searches with simple stepwise addition. We created gene trees for Fusarium ITS, EF1-a, RPB1, and RPB2, and Graphium ITS and EF1-a with the same parameters used on the combined parsimony and Bayesian analyses. Alignments from each locus in the combined data sets (ITS, EF1-a, RPB1, and RPB2 for Fusarium, and ITS, and EF1-a, for Graphium) were separated to create seven individual gene alignments. The PARTITIONFINDER 2.1.1 partition schemes from the combined data sets were applied to each of the genes for both genera and each was subjected to Bayesian analysis using to settings specified above for the combined data. We also used PAUP* 4.0A BUILD 167 to conduct combined parsimony searches on each gene alignment. RESULTS Fungal identification via ITS We recovered 50 fungal cultures from 24 Coptoborus specimens. Each beetle yielded up to three morphologically distinct fungal cultures from their mycangial region. Searched with BLAST based on ITS sequences preliminarily identified these fungi as belonging to nine taxa that included one basidiomycete order (Exobasidiales) and seven genera in the ascomycete class Pezizomycotina (Table 2.5). Four isolates were only recovered from single beetles, significantly aligning to Penicillium griseofulvum Dierckx, Hypocrea virens P. Chaverri, Samuels & E.L. Stewart, Chaetomium globosum Kunze, and Exobasidiales. The remaining four fungal taxa were recovered with greater frequency. Among them, two strains tentatively identified as 77 Phialemoniopsis sp. and Clonostachys sp. (Hypocreales) were isolated from only Coptoborus tolimanus (Eggers). One known and two putative ambrosia fungi were recovered from the examined specimens. Isolates with top BLAST matches to Graphium euwallaceae Twizeyimana, Lynch and Eskalen was cultured from the mycangia of one individual each of Coptoborus coartatus (Sampson 1921) and Coptoborus osbornae Smith & Cognato 2021. The BLAST search also identified two Fusarium species previously isolated from dead and diseased wood. Fusarium sp. 1 RJ2014 was isolated from two individuals of C. coartatus, four Coptoborus cracens Wood 2007, five C. osbornae, and one Coptoborus pseudotenuis (Schedl 1936). Fusarium solani was isolated from one individual each of C. pseudotenuis and C. osbornae. Given the close association both Fusarium share with ambrosia beetle habitat and plant disease (Mohali and Stewart 2017, Jankowiak et al. 2019), it is possible that further analysis will place them within the AFC. However, additional molecular data and phylogenetic analysis are required to confirm this. Specific identification of ambrosia fungi via phylogenetic analyses Comparison of the Yasuní fungal isolates with GenBank sequences preliminarily identified 26 isolates as possible ambrosia fungi. Of these, 17 were tentatively placed into Fusarium and nine were assigned to Graphium euwallaceae. We used combined data of four and two genes to provide specific identification of the Fusarium and Graphium sequences, respectively. Length-variable regions were not found in the alignment of the four Fusarium loci. We subjected the concatenated alignment to Bayesian and parsimony analyses. Two Bayesian runs converged within 40 million generations with a split distribution between analyses that reached a 78 mean standard deviation of 0.015. Most of the clades of the Bayesian consensus tree were well supported with greater than 0.9 posterior probabilities (PP). The Fusarium isolates from Ecuadorian beetles were placed in a clade with Fusarium. sp. AF-9 with very strong support (PP = 1.00) (Figures 2.1 and 2.2). The parsimony analysis of the Fusarium data (293 informative characters) evaluated 1.0 x 1011 rearrangements and yielded 909 000 trees with a best score of 641 steps. A strict consensus of these shared general agreement with the consensus tree generated from the Bayesian analysis. Together, the parsimony and Bayesian analyses provided strong support placing the Ecuadorian Fusarium within the AFC AF-9 lineage (Kasson et al. 2013, O’Donnell et al. 2015). Gene trees created from single-gene alignments and subjected to identical analyses as the combined Fusarium data were largely congruent with the combined tree. Most differences stemmed from lack of resolution in the gene trees. The gene trees for ITS, EF1-a and RPB2 placed the Coptoborus fungal isolates inside the Euwallacea AFC lineage and support the identification of them as F. sp. AF-9 (Figures 2.3–2.6). Several length-variable regions were found in the alignment of the ITS and EF1-a data from Graphium. These length-variable regions were most evident in the sequences other than those from the Ecuadorian fungal cultures. After 40 million generations, the two Bayesian runs had converged. The mean standard deviation of the split distribution between them was 0.002. The consensus tree resulting from Bayesian analysis was poorly resolved (Figures 2.7 and 2.8). Some lineages associated with non-xyleborine beetles (Graphium pseudormiticum M. Mouton & M.J. Wingf., Graphium fimbriasporum (M. Morelet) K. Jacobs, Kristis & M.J. Wingf., and Graphium scolytodis M. Kolařík & J. Hulcr) and several Graphium species associated with storm-injured trees (Graphium adansoniae Cruwy., Z.W. de Beer & Jol. Roux, Graphium 79 madagascariense Cruwy., Z.W. de Beer & Jol. Roux, Graphium fabiforme Cruwy., Z.W. de Beer & Jol. Roux, and Graphium penicillioides Corda) formed well supported clades (Figures 2.7 and 2.8). However, the Graphium isolated from Coptoborus and those associated with wood- infesting insects, including Euwallacea, formed a weakly supported clade (PP = 0.64) that also included the soil fungus G. basitruncatum. Notably, the ambrosia species Graphium euwallaceae remained unresolved within this group. Parsimony analysis of the combined Graphium dataset included 280 informative characters and evaluated 6.7 x 1010 combinations. The best tree score, 1115, was shared by 1 989 000 trees. Gene trees from EF1-a were consistent with the combined tree while the ITS tree showed low branch support and poor resolution for the ambrosia species. (Figures 2.9–2.10). DISCUSSION This study is the first to survey Neotropical xyleborine beetles for symbiotic fungi within their native range. We obtained 50 fungal cultures representing at least eight taxa from the mycangia of 24 beetles collected in Yasuní, Orellana Province, Ecuador. Fungi putatively identified as representing the order Exobasidiales and the species Penicillium griseofulvum, Hypocrea virens, and Chaetomium globosum were each recovered from only a single beetle. However, several taxa were repeatedly isolated from more than one beetle: Phialemoniopsis sp., a poorly understood genus, and Clonostachys sp., commonly associated with both plants and soil where it functions as an endophyte, saprotroph, and parasite to nematodes and Neonectria fungi (Yu and Sutton 1997, Schroers et al. 1999, Zhang et al. 2008, Stauder et al. 2020). Notably, Clonostachys rosea (Link: Fries) Schroers, Samuels, Seifert, and Gams is a known biological control agent 80 against several plant parasites, including Fusarium graminearum Schwabe, which causes Fusarium head blight on wheat and barley (Xue et al. 2014, Nygren et al. 2018, Demissie et al. 2018). Clonostachys spp. have also been recovered from the wood of trees affected by the ambrosia beetle-vectored disease Fusarium Dieback (Carrillo et al. 2020b). Notably in this study, Clonostachys spp. was cultured from C. tolimanus from the same infested wood sample as C. coartatus and its cultured Fusarium and Graphium fungi. Given that contamination of ambrosia beetle galleries with competing saproxylic fungi is common (Franke-Grosmann 1967, Malacrinó et al. 2017), and that neither Clonostachys nor Phialemoniopsis have been reported in association with ambrosia beetles, it is unlikely that they participate in the ambrosia symbiome. However, if these fungi are found in the mycangia of other xyleborines, they may deserve further investigation to rule out a symbiotic association. More broadly, fungi identified as Fusarium sp AF-9. and Graphium euwallaceae were associated with Coptoborus collected from Yasuní. These fungi were previously shown to function as nutritional partners with other ambrosia beetles (e.g., Norris and Baker 1968, Freeman et al. 2016, Carrillo et al. 2020a). We propose that they may be symbionts of xyleborines of the area. Our recovery of Fusarium sp. AF-9 in Ecuador is a novel discovery from neotropical xyleborines. While the Ambrosia Fusarium Clade has been well documented from beetles in the related genera Euwallacea and Xyleborus, this is the first time the fungus has been isolated from mycangia of Coptoborus spp. with confirmation from DNA data. With the exception of a single Costa Rican specimen of Xyleborus ferrugineus whose identification is questionable and unconfirmed (Kasson et al. 2013, O’Donnell et al. 2015), genetic data have only documented Fusarium to associate with Euwallacea spp. in the Middle East (Israel), Australia (Queensland), 81 Asia (Taiwan, India, Sri Lanka, Malaysia, Singapore), and North America (Florida, Pennsylvania, and California, USA) (Mendel et al. 2012, Kasson et al. 2013, O’Donnell et al. 2015, Na et al. 2018). Most of these records are from invasive beetle species living outside their native ranges. Phylogenetic analyses that included wide representation of the previously collected AFC placed our Fusarium isolates from Coptoborus with Ambrosia Fusarium sp. AF-9 supported by strong posterior probability and bootstrap values (Figure 2.1). We isolated this fungus from three Coptoborus species living in two different plant hosts. As previously reported, this group is sister to Fusarium pseudensiforme Samuels, Nalim & Geiser and closely related to F. duplospermum (sp. AF-8, Figure 2.1) (Kasson et al. 2013, Aoki et al. 2021).Together with previous work on the AFC, we show that Fusarium lives with their beetle partners in a variety of plant hosts and associates with several Euwallacea and Coptoborus species, including some living within their native ranges and some exotic invasive species (Mendel et al. 2012, Eskalen et al. 2013, Freeman et al. 2013, Stilwell et al. 2014, O’Donnell et al. 2015, Li et al. 2016, Lynch et al. 2016, Short et al. 2017, Na et al. 2018, Castro et al. 2019). The fidelity between ambrosia beetles and their associated fungi spans a spectrum where some partnerships are constant, and others are promiscuous. Even within Xyleborini, some lineages share the same fungal genus as a symbiont. For instance, Ambrosiodmus and Ambrosiophilus maintain a symbiotic relationship with the basidiomycete fungus Flavodon (Li et al. 2015, Kasson et al. 2016). The clade including Cnestus, Hadrodemius, Diuncus, Anisandrus, Eccoptopterus, and Xylosandrus associates with various fungi from a single lineage in the genus Ambrosiella (Mayers et al. 2015, Skelton et al. 2019). The relationships among other ambrosia beetle genera and their fungi are looser and characterized by multiple partner-switching events. 82 Various Xyleborus species carry multiple fungi in their mycangia (Norris 1965, Baker and Norris 1968). Although many rely on Raffaelea spp. for nutrition (Gebhardt et al. 2004, Harrington et al. 2010, Saucedo et al. 2017), lateral transfer of the laurel wilt fungus Harringtonia lauricola (T.C. Harr., Fraedrich & Aghayeva) Z.W. de Beer & M. Procter (= Raffaelea lauricola) appears to be common among Xyleborus species (Carrillo et al. 2014, Ploetz et al. 2017). Euwallacea spp. rely on two nutritional symbionts from the fungal genera Fusarium and Graphium (Freeman et al. 2013, 2016, Li et al. 2016, Lynch et al. 2016, Carrillo et al. 2020a) and likely underwent several host-switching events in their evolutionary histories (O’Donnell et al. 2015). Factors mediating the specificity of scolytine-fungal relationships are poorly understood. It is likely that some characteristics of the beetle, such as internal mycangium structure and the associated glands help regulate which fungi can successfully survive inside the mycangium (Schedl 1962, Skelton et al. 2019, Mayers et al. 2020a, 2020b). However, there is also evidence that fungal preference for specific environmental conditions, and other taxonomically constrained characteristics play a role in determining whether ambrosia fungi are associated with a limited number of ambrosia beetle species, or if they are likely to switch partners frequently (Francke-Grosmann 1967, Rassati et al. 2016 Miller et al. 2019). Considering previous work on relationships of xyleborine beetles and fungal associates, the isolation of Fusarium sp. AF-9 from Coptoborus is not surprising. Coptoborus and the other Neotropical xyleborine genera are sister to Xyleborus (Cognato et al. 2011, Smith and Cognato 2021), which use Raffaelea spp. as symbionts but appear to switch fungal partners readily (Freeman et al. 2013, 2016, Li et al. 2016, Lynch et al. 2016, Carrillo et al. 2020a). Fusarium- associating Euwallacea beetles are not monophyletic with Coptoborus but more distantly related (Cognato et al. 2011, Gohli et al. 2017). This indicates a potential lateral transfer of Fusarium sp. 83 AF-9 among non-related beetle taxa given the close phylogenetic distance between the Fusarium sp. AF-9 we recovered, and those previously isolated from Euwallacea (Kasson et al. 2013, O’Donnell et al. 2015). Graphium isolates from Coptoborus spp. were weakly associated with other Graphium from Euwallacea spp. and other wood-infesting beetles and G. basitruncatum. More informative sequence data are required to fully reveal the relationships within the genus and between various ambrosia Graphium spp. Similarly, more work is needed to illuminate the nature of the relationship between Coptoborus and Fusarium sp. AF-9 and Graphium. While association with multiple specimens and species is compelling evidence, repeated discoveries of the association from multiple locations and a more in-depth nutritional study is needed before their association can be categorized with confidence as symbiotic or mutualistic. More inclusive sampling, including a wider sampling of infested tree hosts, more representative species, and from throughout their neotropical range, is necessary to fully document fungal associations of Coptoborus and other Neotropical xyleborine genera. This should include morphological study of the fungi to characterize the reproductive structures. Our confirmation that Coptoborus spp. in Ecuador associate with AFC fungi could be important for the management of balsa and other crop trees in South America. The new discovery of Graphium spp. living with these beetles may also be important for silviculture and ecological conservation. Additionally, knowledge about these potentially pathogenic fungi vectored by Coptoborus beetles will enhance protective measures if they establish themselves beyond their native ranges. 84 ACKNOWLEDGMENTS This paper and the associated research would not have been possible without generous help from Clifford Keil, who helped plan logistics and facilitated collection, transport, and export permits. The authors also thank Violet Butterwort and Malena Martínez Chévez who provided valuable help collecting specimens and culturing ambrosia fungi. We thank two anonymous reviews for their helpful suggestions that improved this study. Finally, thanks are due to Sarah M. Smith for assisting with the identification of Coptoborus specimens and to Jiri Hulcr and his team for their advice on fungal culturing. Specimens were collected and exported under the following agreements, MAE-DNB-CM-2015-M-0001, ATM-CM-2015-0039-005–2019-M-0001, 003-17- IC-FAU-DNB/MA, QCAZ-2018-004Y. This study was funded by the Committee for Research and Exploration of the National Geographic Society to Anthony I Cognato (9975-16); The Coleopterists Society Graduate Student Research Enhancement Award, Society of Systematic Biologists Award for Graduate Student Research, Department of Entomology and The Graduate School, MSU to Rachel K Osborn. 85 APPENDIX 86 Species Strain Xyleborine Associate Fusarium sp. AF-9* ROF4 Coptoborus coartatus Fusarium sp. AF-9* ROF7 Coptoborus coartatus Fusarium sp. AF-9* ROF8 Coptoborus coartatus Fusarium sp. AF-9* ROF40 Coptoborus cracens Fusarium sp. AF-9* ROF49 Coptoborus pseudotenuis Fusarium sp. AF-9* ROF50 Coptoborus pseudotenuis Fusarium sp. AF-9* ROF52 Coptoborus cracens Fusarium sp. AF-9* ROF53 Coptoborus cracens Fusarium sp. AF-9* ROF55 Coptoborus cracens Fusarium sp. AF-9* ROF56 Coptoborus cracens Fusarium sp. AF-9* ROF57 Coptoborus osbornae Fusarium sp. AF-9* ROF59 Coptoborus osbornae Fusarium sp. AF-9* ROF60 Coptoborus osbornae Fusarium sp. AF-9* ROF62 Coptoborus osbornae Fusarium sp. AF-9* ROF64 Coptoborus osbornae Fusarium sp. AF-9* ROF67 Coptoborus osbornae Fusarium sp. AF-9* ROF70 Coptoborus osbornae Graphium sp. ROF5 Coptoborus coartatus Graphium sp. ROF6 Coptoborus coartatus Graphium sp. ROF58 Coptoborus osbornae Graphium sp. ROF61 Coptoborus osbornae Graphium sp. ROF63 Coptoborus osbornae Graphium sp. ROF65 Coptoborus osbornae Graphium sp. ROF66 Coptoborus osbornae Graphium sp. ROF69 Coptoborus osbornae Graphium sp. ROF71 Coptoborus osbornae Table 2.1: Fusarium and Graphium fungal isolates cultured from Ecuadorian Coptoborus beetles collected in Yasuní, Orellana, Ecuador and used in phylogenetic analyses. 87 Table 2.1 (cont’d) GenBank accession no. Strain ITS EF1a RPB2 ROF4 OL711912 OM416955 OM304845 ROF7 OL711913 OM416956 … ROF8 OL711914 OM416957 … ROF40 OL711915 OM416958 OM304846 ROF49 OL711916 OM416959 OM315195 ROF50 OL711917 OM416960 OM315196 ROF52 OL711918 OM416961 OM315197 ROF53 OL711919 OM416962 OM304847 ROF55 OL711920 OM416963 OM315198 ROF56 OL711921 OM416964 OM304848 ROF57 OL711922 OM416965 OM315199 ROF59 OL711923 OM416966 OM315200 ROF60 OL711924 OM416967 OM315201 ROF62 OL711925 OM416968 OM297005 ROF64 OL711926 OM416969 OM304849 ROF67 OL711927 OM416970 … ROF70 OL711928 OM416971 … ROF5 OL963597 … … ROF6 OL963598 … … ROF58 OL963599 … … ROF61 OL963600 … … ROF63 OL963601 … … ROF65 OL963602 … … ROF66 OL963603 … … ROF69 OL963604 … … ROF71 OL963605 … … 88 Locus Primer Sequence 5'-3' Source ITS4 TCCTCCGCTTATTGATATGC ITS ITS5 GGAAGTAAAAGTCGTAACAAGG White et al., 1990 EF1F TGCGGTGGTATCGACAAGCGT EF1-𝛂 EF1R AGCATGTTGTCGCCGTTGAAG Lynch et al., 2016 Fa CAYAARGARTCYATGATGGGWC O'Donnell et al., RPB1 G2R GTCATYTGDGTDGCDGGYTCDCC 2010 52f GGGGWGAYCAGAAGAAGGC RPB2 7cr CCCATRGCTTGYTTRCCCAT Lynch et al., 2016 Table 2.2: Amplification primers used for gene sequencing. 89 # Locus Description Temperature Time Cycles 15 Hot Start 95.0°C 1 min 30 Denature 94.0°C sec ITS, 30 Annealing 55.0°C 40 RPB1 sec Extension 72.0°C 1 min Final 10 72.0°C 1 Extension min 15 Hot Start 95.0°C 1 min 30 Denature 94.0°C sec EF1-𝛂 30 40 Annealing 57.0°C sec Extension 72.0°C 1 min Final 10 72.0°C 1 Extension min 15 Hot Start 95.0°C 1 min 30 Denature 94.0°C sec RPB2 30 40 Annealing 59.0°C sec Extension 72.0°C 1 min Final 10 72.0°C 1 Extension min Table 2.3: PCR conditions used for gene sequencing. 90 Species Strain Insect Associate Fusarium ambrosium (AF-1) NRRL 62942 Euwallacea sp. NRRL Fusarium ambrosium (AF-1) 22345** Euwallacea fornicatus NRRL Fusarium ambrosium (AF-1) 36510** Euwallacea fornicatus NRRL Fusarium ambrosium (AF-1) 46583** Euwallacea fornicatus NRRL Fusarium ambrosium (AF-1) 62605** Euwallacea fornicatus Fusarium euwallaceae (AF-2) NRRL 62626 Euwallacea sp. NRRL Fusarium floridanum (AF-3) 62606** Euwallacea sp. Fusarium floridanum (AF-3) NRRL 62608 NA Fusarium floridanum (AF-3) NRRL 62628 Euwallacea interjectus Fusarium floridanum (AF-3) NRRL 62629 Euwallacea interjectus Fusarium oligoseptatum (AF-4) NRRL 62578 Euwallacea validus Fusarium oligoseptatum (AF-4) NRRL 62579 Euwallacea validus Fusarium oligoseptatum (AF-4) NRRL 62580 Euwallacea validus Fusarium oligoseptatum (AF-4) NRRL 62581 Euwallacea validus Fusarium oligoseptatum (AF-4) NRRL 62582 Euwallacea validus Fusarium tuaranense (AF-5) NRRL 22231 NA Fusarium tuaranense (AF-5) NRRL 46518 NA Fusarium tuaranense (AF-5) NRRL 46519 NA Table 2.4: Fusarium and Graphium fungal sequences obtained from GenBank and used in phylogenetic analysis. Sources: aO'Donnell et al. 2015, bKasson et al. 2013 cCarrillo et al. 2019, d Mendel et al. 2012, eNa et al. 2018, fZhang et al. 2006, gO'Donnell direct submission, hO'Donnll et al. 2007, iO'Donnell et al. 2010, jLynch et al. 2016, kCruywagen et al. 2010, lPaciura et al. 2008, mJacobs et al. 2003, nHulcr et al. 2007, oOkada et al. 2000, pTwizeyimana et at Direct Submission, qKolarik et al. unknown date, and rHameline et al. Direct Submission. *Outgroup. ** Fungus was isolated from xyleborine gallery. TEx-type strain. ?Identification of this beetle partner is questionable (O'Donnell et al., 2015). 91 Table 2.4 (cont’d) Species Strain Insect Associate NRRL Fusarium sp. AF-6 62590** Euwallacea sp. NRRL Fusarium sp. AF-6 62591** Euwallacea sp. Fusarium obliquiseptatum (AF-7) NRRL 62610 Euwallacea sp. NRRL Fusarium obliquiseptatum (AF-7) 62611** Euwallacea sp. Fusarium duplospermum (AF-8) NRRL 62583 Euwallacea sp. Fusarium duplospermum (AF-8) NRRL 62584 Euwallacea sp. Fusarium duplospermum (AF-8) NRRL 62585 Euwallacea sp. Fusarium duplospermum (AF-8) NRRL 62586 Euwallacea sp. Fusarium duplospermum (AF-8) NRRL 62587 Euwallacea sp. Fusarium duplospermum (AF-8) NRRL 62589 Euwallacea sp. Fusarium sp. AF-9 NRRL 22643 Xyleborus ferrigineus? Fusarium sp. AF-9 NRRL 66088 unknown Fusarium pseudensiforme NRRL 46517 NA Fusarium drepaniforme (AF-10) NRRL 62941 unknown Fusarium papillatum (AF-11) NRRL 62943 Euwallacea sp. Fusarium papillatum (AF-11) NRRL 62944 Euwallacea sp. Fusarium kuroshium (AF-12) NRRL 62945 Euwallacea sp. Fusarium kuroshium (AF-12) NRRL 62946 Euwallacea sp. Fusarium sp. AF-13 UCR 5584 Euwallacea sp. Fusarium sp. AF-13 UCR 6394 Euwallacea sp. Fusarium sp. AF-13 UCR 6403 Euwallacea sp. Fusarium sp. AF-13 UCR 6409 Euwallacea sp. Fusarium sp. AF-13 UCR 6432 Euwallacea sp. Fusarium sp. AF-14 TW 2 NA Fusarium sp. AF-14 TW 56 NA Fusarium sp. AF-14 UCR 5499 Euwallacea sp. Fusarium sp. AF-14 UCR 5509 Euwallacea sp. Fusarium sp. AF-14 UCR 5546 Euwallacea sp. Fusarium sp. AF-14 UCR 6436 Euwallacea sp. Fusarium sp. AF-15 UCR 6395 Euwallacea sp. 92 Table 2.4 (cont’d) Species Strain Insect Associate Fusarium sp. AF-15 TW 15 NA Fusarium sp. AF-15 TW 45 NA Fusarium sp. AF-16 TW 4 NA Fusarium sp. AF-16 TW 25 NA Fusarium sp. AF-16 TW 34 NA Fusarium sp. AF-16 TW 37 NA Fusarium sp. AF-16 UCR 5508 Euwallacea sp. Fusarium sp. AF-16 UCR 5513 Euwallacea sp. Fusarium sp. AF-16 UCR 6405 Euwallacea sp. Fusarium sp. AF-17 TW 40 NA Fusarium sp. AF-17 UCR 5545 Euwallacea sp. Fusarium sp. AF-17 UCR 6414 Euwallacea sp. Fusarium sp. AF-18 TW 1 NA Fusarium sp. AF-18 TW 44 NA Fusarium sp. AF-18 TW 55 NA Fusarium sp. AF-18 UCR 5557 Euwallacea sp. Fusarium sp. AF-18 UCR 6411 Euwallacea sp. Fusarium sp. AF-18 UCR 6417 Euwallacea sp. Fusarium euwallaceae (AF-2) NRRL 54722 Euwallacea sp. Fusarium euwallaceae (AF-2) NRRL 54723 Euwallacea sp. Fusarium euwallaceae (AF-2) NRRL 54724 Euwallacea sp. Fusarium euwallaceae (AF-2) NRRL 54725 Euwallacea sp. Fusarium euwallaceae (AF-2) NRRL 54726 Euwallacea sp. Fusarium sp. NRRL 54727 Euwallacea sp. Fusarium sp. NRRL 54728 Euwallacea sp. Fusarium sp. JM-2017a UCR 3641 NA Fusarium sp. JM-2017a UCR 3644 NA Fusarium sp. JM-2017a UCR 3651 Euwallacea sp. Fusarium sp. JM-2017a UCR 3652 NA Fusarium sp. JM-2017a UCR 3653 NA 93 Table 2.4 (cont’d) Species Strain Insect Associate Fusarium sp. JM-2017a UCR 3654 NA Fusarium sp. JM-2017a UCR 3657 Euwallacea sp. Fusarium sp. JM-2017a UCR 3659 Euwallacea sp. Fusarium sp. JM-2017a UCR 3660 Euwallacea sp. Fusarium sp. JM-2017a UCR 3661 NA Fusarium sp. JM-2017a UCR 4672 Euwallacea sp. Fusarium sp. JM-2017a UCR 4673 Euwallacea sp. Fusarium sp. JM-2017a UCR 4674 Euwallacea sp. Fusarium sp. JM-2017a UCR 4675 Euwallacea sp. Fusarium sp. JM-2017a UCR 4676 Euwallacea sp. Fusarium sp. JM-2017a UCR 4677 Euwallacea sp. Fusarium sp. JM-2017a UCR 4678 Euwallacea sp. Fusarium sp. JM-2017a UCR 4679 Euwallacea sp. Fusarium sp. JM-2017a UCR 4680 Euwallacea sp. Fusarium sp. JM-2017a UCR 4681 Euwallacea sp. Fusarium kuroshium TW 43 NA Fusarium kuroshium UCR 6408 Euwallacea sp. Fusarium neocosmosporiellum NRRL 22468* NA Fusarium neocosmosporiellum NRRL 43467* NA Fusarium lichenicola NRRL 32434* NA Graphium kuroshium UCR 4593 NA Graphium kuroshium UCR 4594T NA Graphium kuroshium UCR 4606 NA Graphium kuroshium UCR 4607 NA Graphium kuroshium UCR 4608 Euwallacea sp. Graphium kuroshium UCR 4609 Euwallacea sp. Graphium kuroshium UCR 4616 NA Graphium kuroshium UCR 4617 NA Graphium kuroshium UCR 4618 Euwallacea sp. Graphium kuroshium UCR 4622 Euwallacea sp. 94 Table 2.4 (cont’d) Species Strain Insect Associate Graphium sp. UCR 5497 Euwallacea sp. Graphium sp. UCR 5501 Euwallacea sp. Graphium sp. UCR 5506 Euwallacea sp. Graphium sp. UCR 5512 Euwallacea sp. Graphium sp. UCR 5517 Euwallacea sp. Graphium kuroshium UCR 5519 Euwallacea sp. Graphium sp. UCR 5528 Euwallacea sp. Graphium sp. UCR 5531 Euwallacea sp. Graphium sp. UCR 5548 Euwallacea sp. Graphium kuroshium UCR 5549 Euwallacea sp. Graphium sp. UCR 6662 Euwallacea sp. Graphium sp. UCR 6667 Euwallacea sp. Graphium sp. II UCR 2132 NA Graphium sp. II UCR 2137 NA Graphium sp. II UCR 2140 NA Graphium sp. I UCR 2159 NA Graphium sp. I UCR 2160 NA Graphium sp. I UCR 2162 NA Graphium sp. I UCR 2163 NA Graphium sp. I UCR 2164 NA Graphium sp. I UCR 2165 NA Graphium sp. I UCR 2166 NA Graphium sp. III UCR 2289 NA Graphium sp. III UCR 2291 NA Graphium carbonarium UCR 2300 NA Graphium euwallaceae UCR 2308 NA Graphium carbonarium UCR 2325 NA Graphium carbonarium UCR 2329 NA Graphium euwallaceae UCR 2974 NA Graphium euwallaceae UCR 2975 NA Graphium euwallaceae UCR 2976 NA 95 Table 2.4 (cont’d) Species Strain Insect Associate Graphium euwallaceae UCR 2977 NA Graphium euwallaceae UCR 2978 NA Graphium euwallaceae UCR 2979 NA Graphium euwallaceae UCR 2980T NA Graphium euwallaceae UCR 2981 NA Graphium pseudormiticum CMW 12285 Pissodes sp. (Molytinae) Graphium adansoniae CMW 30617 NA Graphium adansoniae CMW 30618T NA Graphium adansoniae CMW 30620 NA Graphium fabiforme CMW 30626T NA Graphium fabiforme CMW 30627 NA Graphium madagascariense CMW 30628T NA Graphium madagascariense CMW 30629 NA Graphium penicillioides CMW 5292 NA Graphium penicillioides CMW 5295 NA Graphium carbonarium CMW 12418 Pissodes sp. (Molytinae) Graphium carbonarium CMW 12420T Pissodes sp. (Molytinae) Graphium pseudormiticum CMW 503T Orthotomicus erosus Graphium laricis CMW 5601T Ips cembrae Graphium laricis CMW 5603 Ips cembrae Graphium fimbriasporum CMW 5605T Ips typographus Graphium fimbriasporum CMW 5606 Ips typographus Graphium scolytodis CCF 3566 Scolytodes unipunctatus Graphium scolytodis CCF 3570 Scolytodes unipunctatus Graphium basitruncatum JCM 9300* NA Geosmithia putterillii U 160* NA Ophiostoma ulmi Q 412T-0* NA 96 Table 2.4 (cont’d) Strain Host Plant/Source Origin NRRL 62942 Camellia sinensis (tea) Sri Lanka NRRL 22345** Camellia sinensis (tea) India NRRL 36510** Camellia sinensis (tea) India NRRL 46583** Camellia sinensis (tea) India NRRL 62605** Camellia sinensis (tea) India NRRL 62626 Persea americana (avocado) California, USA NRRL 62606** Acer negundo (box elder) Florida, USA NRRL 62608 Acer negundo (box elder) Florida, USA NRRL 62628 Acer negundo (box elder) Florida, USA NRRL 62629 Acer negundo (box elder) Florida, USA NRRL 62578 Ailanthus altissima (tree of heaven) Pennsylvania, USA NRRL 62579 Ailanthus altissima (tree of heaven) Pennsylvania, USA NRRL 62580 Ailanthus altissima (tree of heaven) Pennsylvania, USA NRRL 62581 Ailanthus altissima (tree of heaven) Pennsylvania, USA NRRL 62582 Ailanthus altissima (tree of heaven) Pennsylvania, USA NRRL 22231 Hevea brasiliensis (Pará rubber) Malasia NRRL 46518 Hevea brasiliensis (Pará rubber) Malasia NRRL 46519 Hevea brasiliensis (Pará rubber) Malasia NRRL 62590** Persea americana (avocado) Florida, USA NRRL 62591** Persea americana (avocado) Florida, USA Queensland, NRRL 62610 Unknown Australia NRRL Queensland, 62611** Persea americana (avocado) Australia NRRL 62583 Persea americana (avocado) Florida, USA NRRL 62584 Persea americana (avocado) Florida, USA NRRL 62585 Persea americana (avocado) Florida, USA NRRL 62586 Persea americana (avocado) Florida, USA NRRL 62587 Persea americana (avocado) Florida, USA 97 Table 2.4 (cont’d) Strain Host Plant/Source Origin NRRL 62589 Persea americana (avocado) Florida, USA NRRL 22643 Unknown Costa Rica NRRL 66088 Delonix regia (royal poinciana) Florida, USA NRRL 46517 Unknow recently dead tree Sri Lanka NRRL 62941 Unknown Singapore, Malaysia NRRL 62943 Camellia sinensis (tea) Sri Lanka NRRL 62944 Camellia sinensis (tea) Sri Lanka NRRL 62945 Platanus racemosa (sycamore) California, USA NRRL 62946 Platanus racemosa (sycamore) California, USA UCR 5584 Unknown Taiwan UCR 6394 Unknown Taiwan UCR 6403 Unknown Taiwan UCR 6409 Unknown Taiwan UCR 6432 Unknown Taiwan TW 2 Persea americana (avocado) Taiwan TW 56 Persea americana (avocado) Taiwan UCR 5499 Unknown Taiwan UCR 5509 Unknown Taiwan UCR 5546 Unknown Taiwan UCR 6436 Unknown Taiwan UCR 6395 Unknown Taiwan TW 15 Persea americana (avocado) Taiwan TW 45 Persea americana (avocado) Taiwan TW 4 Persea americana (avocado) Taiwan TW 25 Persea americana (avocado) Taiwan TW 34 Persea americana (avocado) Taiwan TW 37 Persea americana (avocado) Taiwan 98 Table 2.4 (cont’d) Strain Host Plant/Source Origin UCR 5508 Unknown Taiwan UCR 5513 Unknown Taiwan UCR 6405 Unknown Taiwan TW 40 Persea americana (avocado) Taiwan UCR 5545 Unknown Taiwan UCR 6414 Unknown Taiwan TW 1 Persea americana (avocado) Taiwan TW 44 Persea americana (avocado) Taiwan TW 55 Persea americana (avocado) Taiwan UCR 5557 Unknown Taiwan UCR 6411 Unknown Taiwan UCR 6417 Unknown Taiwan NRRL 54722 Persea americana (avocado) Israel NRRL 54723 Persea americana (avocado) Israel NRRL 54724 Persea americana (avocado) Israel NRRL 54725 Persea americana (avocado) Israel NRRL 54726 Persea americana (avocado) Israel NRRL 54727 Persea americana (avocado) Israel NRRL 54728 Persea americana (avocado) Israel UCR 3641 Platanus racemosa (sycamore) California, USA UCR 3644 Platanus racemosa (sycamore) California, USA UCR 3651 Unknown California, USA UCR 3652 Persea americana (avocado) California, USA UCR 3653 Persea americana (avocado) California, USA UCR 3654 Persea americana (avocado) California, USA UCR 3657 Unknown California, USA UCR 3659 Unknown California, USA 99 Table 2.4 (cont’d) Strain Host Plant/Source Origin UCR 3660 Unknown California, USA UCR 3661 Persea americana (avocado) California, USA UCR 4672 Unknown Taiwan UCR 4673 Unknown Taiwan UCR 4674 Unknown Taiwan UCR 4675 Unknown Taiwan UCR 4676 Unknown Taiwan UCR 4677 Unknown Taiwan UCR 4678 Unknown Taiwan UCR 4679 Unknown Taiwan UCR 4680 Unknown Taiwan UCR 4681 Unknown Taiwan TW 43 Persea americana (avocado) Taiwan UCR 6408 Unknown Taiwan NRRL 22468* Stored peanuts Guinea NRRL 43467* Human eye Louisiana, USA NRRL 32434* Human Germany UCR 4593 Persea americana (avocado) California, USA UCR 4594T Persea americana (avocado) California, USA UCR 4606 Persea americana (avocado) California, USA UCR 4607 Persea americana (avocado) California, USA UCR 4608 unknown California, USA UCR 4609 unknown California, USA UCR 4616 Persea americana (avocado) California, USA UCR 4617 Persea americana (avocado) California, USA UCR 4618 unknown California, USA UCR 4622 unknown California, USA 100 Table 2.4 (cont’d) Strain Host Plant/Source Origin UCR 5497 unknown Taiwan UCR 5501 unknown Taiwan UCR 5506 unknown Taiwan UCR 5512 unknown Taiwan UCR 5517 unknown Taiwan UCR 5519 unknown Taiwan UCR 5528 unknown Taiwan UCR 5531 unknown Taiwan UCR 5548 unknown Taiwan UCR 5549 unknown Taiwan UCR 6662 unknown Florida, USA UCR 6667 unknown Florida, USA UCR 2132 Durio sp. (Durian) Thailand UCR 2137 Durio sp. (Durian) Thailand UCR 2140 Durio sp. (Durian) Thailand UCR 2159 Ailanthus altissima (tree of heaven) Pennsylvania, USA UCR 2160 Ailanthus altissima (tree of heaven) Pennsylvania, USA UCR 2162 Ailanthus altissima (tree of heaven) Pennsylvania, USA UCR 2163 Ailanthus altissima (tree of heaven) Pennsylvania, USA UCR 2164 Ailanthus altissima (tree of heaven) Pennsylvania, USA UCR 2165 Ailanthus altissima (tree of heaven) Pennsylvania, USA UCR 2166 Ailanthus altissima (tree of heaven) Pennsylvania, USA Acacia auriculiformis (earleaf UCR 2289 acacia) Veitnam Acacia auriculiformis (earleaf UCR 2291 acacia) Veitnam Acacia auriculiformis (earleaf UCR 2300 acacia) Veitnam Acacia auriculiformis (earleaf UCR 2308 acacia) Veitnam UCR 2325 Ricinus communis (castor bean) Veitnam 101 Table 2.4 (cont’d) Strain Host Plant/Source Origin UCR 2329 Ricinus communis (castor bean) Veitnam UCR 2974 Ricinus communis (castor bean) California, USA UCR 2975 Acer negundo (box elder) California, USA UCR 2976 Ricinus communis (castor bean) California, USA UCR 2977 Acacia floribunda (weeping acacia) California, USA UCR 2978 Erythrina atitlanensis California, USA UCR 2979 Quercus agrifolia (coast live oak) California, USA UCR 2980T Persea americana (avocado) California, USA UCR 2981 Persea americana (avocado) California, USA CMW 12285 Tsuga dumosa Himalayan hemlock China CMW 30617 Adansonia digitata (African baobab) South Africa CMW 30618T Adansonia digitata (African baobab) South Africa CMW 30620 Adansonia digitata (African baobab) South Africa CMW 30626T Adansonia rubrostipa (fony boabab) Madagascar CMW 30627 Adansonia rubrostipa (fony boabab) Madagascar CMW 30628T Adansonia rubrostipa (fony boabab) Madagascar CMW 30629 Adansonia rubrostipa (fony boabab) Madagascar CMW 5292 Populus nigra (black poplar) Czech Republic CMW 5295 Populus nigra (black poplar) Czech Republic CMW 12418 Salix babylonica (weeping willow) China CMW 12420T Salix babylonica (weeping willow) China CMW 503T Pinus sp. (pine) South Africa CMW 5601T Larix decidua (European larch) Austria CMW 5603 Larix decidua (European larch) Austria CMW 5605T Picea abies (Norway spruce) France CMW 5606 Picea abies (Norway spruce) Austria 102 Table 2.4 (cont’d) Strain Host Plant/Source Origin CCF 3566 Cercropia sp. Costa Rica CCF 3570 Cercropia sp. Costa Rica * JCM 9300 Soil Solomon Islands U 160* Ulmus pumila (Siberian elm) Colorado, USA Q 412T-0* Ulmus americana (American elm) … 103 Table 2.4 (cont’d) GenBank accession no. Strain ITS EF1a RPB1 RPB2 NRRL 62942 KM406631a KM406624 KM406638 KM406645a a a NRRL 22345** KC691557b KC691529b KC691586b KC691618b NRRL 36510** KC691558b KC691530b KC691588b KC691619b NRRL 46583** KC691556b KC691528b KC691585b KC691617b NRRL 62605** KC691559b KC691531b KC691589b KC691620b NRRL 62626 KC691560b KC691532b KC691590b KC691621b NRRL 62606** KC691561b KC691533b KC691591b KC691622b NRRL 62608 KC691562b KC691534b KC691592b KC691623b NRRL 62628 KC691563b KC691535b KC691593b KC691624b NRRL 62629 KC691564b KC691536b KC691594b KC691625b NRRL 62578 KC691565b KC691537b KC691595b KC691626b NRRL 62579 KC691566b KC691538b KC691596b KC691627b NRRL 62580 KC691567b KC691539b KC691597b KC691628b NRRL 62581 KC691568b KC691540b KC691598b KC691629b NRRL 62582 KC691569b KC691541b KC691599b KC691630b NRRL 22231 KC691570b KC691542b KC691600b KC691631b NRRL 46518 KC691571b KC691543b KC691601b KC691632b NRRL 46519 KC691572b KC691544b KC691602b KC691633b NRRL 62590** KC691574b KC691546b KC691604b KC691635b NRRL 62591** KC691573b KC691545b KC691603b KC691634b NRRL 62610 KC691575b KC691547b KC691605b KC691636b NRRL 62611** KC691576b KC691548b KC691606b KC691637b NRRL 62583 KC691581b KC691553b KC691611b KC691642b NRRL 62584 KC691582b KC691554b KC691612b KC691643b NRRL 62585 KC691577b KC691549b KC691607b KC691638b NRRL 62586 KC691578b KC691550b KC691608b KC691639b NRRL 62587 KC691579b KC691551b KC691609b KC691640b 104 Table 2.4 (cont’d) GenBank accession no. Strain ITS EF1a RPB1 RPB2 b b b NRRL 62589 KC691580 KC691552 KC691610 KC691641b NRRL 22643 KC691583b … … KC691644b NRRL 66088 KM406632a KM406625a KM406639a KM406646a NRRL 46517 KC691584b KC691555b KC691615b KC691645b NRRL 62941 KM406633a KM406626a KM406640a KM406647a NRRL 62943 KM406635a KM406628a KM406642a … NRRL 62944 KM406634a KM406627a KM406641a KM406648a NRRL 62945 KM406636a KM406629a KM406643a KM406649a NRRL 62946 KM406637a KM406630a KM406644a KM406650a UCR 5584 MK432880c MK435457c MK435509c MK435541c UCR 6394 MK432881c MK435458c MK435510c MK435542c UCR 6403 MK432883c MK435460c MK435512c MK435544c UCR 6409 MK432886c MK435463c MK435515c MK435547c UCR 6432 MK432890c MK435467c MK435519c MK435551c TW 2 MK432862c MK435439c MK435491c MK435523c TW 56 MK432872c MK435449c MK435501c MK435533c UCR 5499 MK432873c MK435450c MK435502c MK435534c UCR 5509 MK432875c MK435452c MK435504c MK435536c UCR 5546 MK432878c MK435455c MK435507c MK435539c UCR 6436 MK432891c MK435468c MK435520c MK435552c UCR 6395 MK432882c MK435459c MK435511c MK435543c TW 15 MK432861c MK435438c MK435490c MK435522c TW 45 MK432870c MK435447c MK435499c MK435531c TW 4 MK432866c MK435443c MK435495c MK435527c TW 25 MK432863c MK435440c MK435492c MK435524c TW 34 MK432864c MK435441c MK435493c MK435525c TW 37 MK432865c MK435442c MK435494c MK435526c 105 Table 2.4 (cont’d) GenBank accession no. Strain ITS EF1a RPB1 RPB2 UCR 5508 MK432874 MK435451 MK435503 MK435535c c c c UCR 5513 MK432876c MK435453c MK435505c MK435537c UCR 6405 MK432884c MK435461c MK435513c MK435545c TW 40 MK432867c MK435444c MK435496c MK435528c UCR 5545 MK432877c MK435454c MK435506c MK435538c UCR 6414 MK432888c MK435465c MK435517c MK435549c TW 1 MK432860c MK435437c MK435489c MK435521c TW 44 MK432869c MK435446c MK435498c MK435530c TW 55 MK432871c MK435448c MK435500c MK435532c UCR 5557 MK432879c MK435456c MK435508c MK435540c UCR 6411 MK432887c MK435464c MK435516c MK435548c UCR 6417 MK432889c MK435466c MK435518c MK435550c NRRL 54722 JQ038014d JQ038007d JQ038021d JQ038028d NRRL 54723 JQ038015d JQ038008d JQ038022d JQ038029d NRRL 54724 JQ038016d JQ038009d JQ038023d JQ038030d NRRL 54725 JQ038017d JQ038010d JQ038024d JQ038031d NRRL 54726 JQ038018d JQ038011d JQ038025d JQ038032d NRRL 54727 JQ038019d JQ038012d JQ038026d JQ038033d NRRL 54728 JQ038020d JQ038013d JQ038027d JQ038034d UCR 3641 KX262196e KX262216e KX262236e KX262256e UCR 3644 KX262197e KX262217e KX262237e KX262257e UCR 3651 KX262198e KX262218e KX262238e KX262258e UCR 3652 KX262199e KX262219e KX262239e KX262259e UCR 3653 KX262200e KX262220e KX262240e KX262260e UCR 3654 KX262201e KX262221e KX262241e KX262261e UCR 3657 KX262202e KX262222e KX262242e KX262262e UCR 3659 KX262203e KX262223e KX262243e KX262263e 106 Table 2.4 (cont’d) GenBank accession no. Strain ITS EF1a RPB1 RPB2 e e e UCR 3660 KX262204 KX262224 KX262244 KX262264e UCR 3661 KX262205e KX262225e KX262245e KX262265e UCR 4672 KX262206e KX262226e KX262246e KX262266e UCR 4673 KX262207e KX262227e KX262247e KX262267e UCR 4674 KX262208e KX262228e KX262248e KX262268e UCR 4675 KX262209e KX262229e KX262249e KX262269e UCR 4676 KX262210e KX262230e KX262250e KX262270e UCR 4677 KX262211e KX262231e KX262251e KX262271e UCR 4678 KX262212e KX262232e KX262252e KX262272e UCR 4679 KX262213e KX262233e KX262253e KX262273e UCR 4680 KX262214e KX262234e KX262254e KX262274e UCR 4681 KX262215e KX262235e KX262255e KX262275e TW 43 MK432868c MK435445c MK435497c MK435529c UCR 6408 MK432885c MK435462c MK435514c MK435546c NRRL 22468* DQ094318f AF178349g KC691616e … NRRL 43467* EF453092h EF452940h HM347178i … NRRL 32434* DQ094444f DQ246977f HM347156i … UCR 4593 KX262276e KX262286e … … UCR 4594T KX262277e KX262287e … … UCR 4606 KX262278e KX262288e … … UCR 4607 KX262279e KX262289e … … UCR 4608 KX262280e KX262290e … … UCR 4609 KX262281e KX262291e … … UCR 4616 KX262282e KX262292e … … UCR 4617 KX262283e KX262293e … … UCR 4618 KX262284e KX262294e … … UCR 4622 KX262285e KX262295e … … 107 Table 2.4 (cont’d) GenBank accession no. Strain ITS EF1a RPB1 RPB2 c c UCR 5497 MK432903 MK435469 … … UCR 5501 MK432902c MK435470c … … UCR 5506 MK432901c MK435471c … … UCR 5512 MK432900c MK435472c … … UCR 5517 MK432899c MK435473c … … UCR 5519 MK432898c MK435474c … … UCR 5528 MK432897c MK435475c … … UCR 5531 MK432896c MK435476c … … UCR 5548 MK432895c MK435477c … … UCR 5549 MK432894c MK435478c … … UCR 6662 MK432893c MK435479c … … UCR 6667 MK432892c MK435480c … … UCR 2132 KM592367j KM363259j … … UCR 2137 KJ131236j KJ131246j … … UCR 2140 KJ131237j KJ131247j … … UCR 2159 KJ131228j KJ131238j … … UCR 2160 KJ131229j KJ131239j … … UCR 2162 KJ131231j KJ131241j … … UCR 2163 KJ131232j KJ131242j … … UCR 2164 KJ131233j KJ131243j … … UCR 2165 KJ131234j KJ131244j … … UCR 2166 KJ131235j KJ131245j … … UCR 2289 KM592368j KM592360j … … UCR 2291 KM592369j KM592361j … … UCR 2300 KM592370j KM592362j … … UCR 2308 KM592371j KM592363j … … UCR 2325 KM592372j KM592364j … … 108 Table 2.4 (cont’d) GenBank accession no. Strain ITS EF1a RPB1 RPB2 j j UCR 2329 KM592373 KM592365 … … UCR 2974 KF540218j KF534799j … … UCR 2975 KF540219j KF534800j … … UCR 2976 KF540220j KF534801j … … UCR 2977 KF540221j KF534802j … … UCR 2978 KF540222j KF534803j … … UCR 2979 KF540223j KF534804j … … UCR 2980T KF540224j KF534805j … … UCR 2981 KF540225j KF534806j … … CMW 12285 HM630608k HM630587k … … CMW 30617 GQ200610k HM630596k … … CMW 30618T GQ200611k HM630598k … … CMW 30620 GQ200613k HM630597k … … CMW 30626T GQ200616k HM630592k … … CMW 30627 GQ200617k HM630593k … … CMW 30628T GQ200619k HM630595k … … CMW 30629 GQ200620k HM630594k … … CMW 5292 HQ335310k HM630600k … … CMW 5295 HQ335311k HM630601k … … CMW 12418 FJ434980l HM630602k … … CMW 12420T FJ434979l HM630603k … … CMW 503T AY148186m HM630586k … … CMW 5601T AY148183m HM630588k … … CMW 5603 AY148182m HM630589k … … CMW 5605T AY148177m HM630590k … … CMW 5606 AY148180m HM630591k … … 109 Table 2.4 (cont’d) GenBank accession no. Strain ITS EF1a RPB1 RPB2 CCF 3566 AM267264n … … … CCF 3570 AM267265n … … … JCM 9300* AB038427o KJ131248p … … U 160* HF546278q HG799859q … HG799911q Q 412T-0* KF854010r KF899888r … … 110 Beetle Fungal Beetle Identity Fungal Identification voucher Strain Coptoborus catulus B88 ROF3 Exobasidiales ROF9 Phialemoniopsis sp. B13 ROF10 Clonostachys sp. ROF11 Phialemoniopsis sp. B14 ROF12 Clonostachys sp. ROF13 Clonostachys sp. B15 ROF72 Phialemoniopsis sp. ROF14 Phialemoniopsis sp. B28 Penicillium ROF15 griseofulvum ROF16 Clonostachys sp. B29 ROF17 Phialemoniopsis sp. Coptoborus tolimanus ROF18 Clonostachys sp. B30 ROF19 Clonostachys sp. ROF20 Clonostachys sp. B31 ROF21 Clonostachys sp. ROF22 Clonostachys sp. ROF23 Clonostachys sp. B32 ROF24 Clonostachys sp. ROF25 Phialemoniopsis sp. ROF26 Clonostachys sp. B33 ROF27 Clonostachys sp. ROF28 Clonostachys sp. Table 2.5: BLAST identities of fungi isolated from Coptoborus spp. 111 Table 2.5 (cont’d) Beetle Fungal Beetle Identity Fungal Identification voucher Strain Coptoborus pseudotenuis B88 ROF49 Fusarium solani ROF50 Fusarium sp. 1 RJ2014 Coptoborus cracens B83 ROF40 Fusarium sp. 1 RJ2014 B89 ROF52 Fusarium sp. 1 RJ2014 B90 ROF53 Fusarium sp. 1 RJ2014 ROF54 Chaetomium globosum B91 ROF55 Fusarium sp. 1 RJ2014 ROF56 Fusarium sp. 1 RJ2014 B9 ROF4 Fusarium sp. 1 RJ2014 ROF5 Graphium euwallaceae B10 Coptoborus coartatus ROF6 Graphium euwallaceae ROF7 Fusarium sp. 1 RJ2014 B11 ROF8 Fusarium sp. 1 RJ2014 ROF58 Graphium euwallaceae B92 ROF59 Fusarium sp. 1 RJ2014 ROF57 Fusarium sp. 1 RJ2014 ROF60 Fusarium solani B93 ROF61 Graphium euwallaceae ROF62 Fusarium sp. 1 RJ2014 B94 ROF63 Graphium euwallaceae Coptoborus osbornae ROF64 Fusarium sp. 1 RJ2014 B95 ROF65 Graphium euwallaceae ROF66 Graphium euwallaceae ROF67 Fusarium sp. 1 RJ2014 B96 ROF68 Hypocrea virens ROF69 Graphium euwallaceae ROF70 Fusarium sp. 1 RJ2014 B97 ROF71 Graphium euwallaceae 112 Table 2.5 (cont’d) Fungal % Seq % Top GenBank Match Strain Similarity Coverage ROF3 KP229361 98.67 85 ROF9 MT887369 99.57 93 ROF10 KP006352 98.53 98 ROF11 MT887369 99.17 93 ROF12 KP006352 98.53 98 ROF13 KP006352 99.17 99 ROF72 MT887369 99.17 93 ROF14 MT887369 99.17 92 ROF15 KT898767 99.61 99 ROF16 KP006352 98.9 98 ROF17 MT887369 99.59 93 ROF18 KP006352 98.53 98 ROF19 KP006352 99.27 98 ROF20 KP006352 99.63 98 ROF21 KP006352 98.9 98 ROF22 KP006352 98.9 98 ROF23 KP006352 99.63 98 ROF24 KP006352 98.9 98 ROF25 MT887369 99.17 94 ROF26 KP006352 99.27 98 ROF27 KP006352 98.9 98 ROF28 KP006352 98.17 98 113 Table 2.5 (cont’d) Fungal % Seq % Top GenBank Match Strain Similarity Coverage ROF49 KR350652 97.94 99 ROF50 MF782769 97.75 100 ROF40 MF782769 97.75 99 ROF52 MF782769 98.15 99 ROF53 MF782769 97.78 99 ROF54 MF476064 99.66 99 ROF55 MF782769 97.78 99 ROF56 MF782769 97.78 99 ROF4 MF782769 97.41 98 ROF5 EF165016 98.01 98 ROF6 EF165016 98.41 80 ROF7 MF782769 97.78 99 ROF8 MF782769 97.78 99 ROF58 EF165016 98.01 99 ROF59 MF782769 97.76 99 ROF57 MF782769 97.41 99 ROF60 KR350652 98.08 98 ROF61 EF165016 98.41 80 ROF62 MF782769 98.14 99 ROF63 EF165016 98.41 99 ROF64 MF782769 97.05 99 ROF65 EF165016 98.41 99 ROF66 EF165016 98.41 99 ROF67 MF782769 97.05 99 ROF68 GU046491 99.37 99 ROF69 EF165016 98.41 99 ROF70 MF782769 98.47 99 ROF71 EF165016 97.61 99 114 Figure 2.1: Condensed Bayesian consensus tree of Fusarium strains reconstructed from ITS, EF1-a, RPB1, and RPB2 sequence data. Nodes are labeled with bootstrap values/posterior probabilities. Clades containing AFC strains associated with Euwallaceae spp. beetles have been collapsed. Fusarium sp. isolated from Coptoborus sp. for this study are written in bold. See Figure 2.2 for the complete tree. 115 Figure 2.2: Complete Bayesian consensus tree of Fusarium strains reconstructed from ITS, EF1- a, RPB1, and RPB2 sequence data. Nodes are labeled with bootstrap values/posterior probabilities. Fusarium sp. isolated from Coptoborus sp. for this study are written in bold. 116 Figure 2.3: Bayesian consensus tree of Fusarium strains built from gene sequence data of EF1- a. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold. 117 Figure 2.4: Bayesian consensus tree of Fusarium strains built from gene sequence data of ITS. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold. 118 Figure 2.5: Bayesian consensus tree of Fusarium strains built from gene sequence data of RPB1. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold. 119 Figure 2.6: Bayesian consensus tree of Fusarium strains built from gene sequence data of RPB2. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold. 120 Figure 2.7: Condensed Bayesian consensus tree of Graphium strains reconstructed from ITS, and EF1-a sequence data. Nodes are labeled with bootstrap values/posterior probabilities. Clades containing strains from G. sp. I, G. sp. II, G. sp. III, G. carbonarium, G. pseudormiticum, G. laricis, G. fimbriasporum, G. fabiforme, G. adansoniae, G. madagascariense, G. penicillioides, and G. scolytodis have been collapsed. Graphium sp. isolated from Coptoborus spp. for this study are written in bold. T Denotes ex-type strains. See Figure 2.8 for the complete tree. 121 Figure 2.8: Complete Bayesian consensus tree of Graphium strains reconstructed from ITS, EF1-a, and RPB2 sequence data. Nodes are labeled with bootstrap values/posterior probabilities. Graphium sp. isolated from Coptoborus sp. for this study are written in bold. T Denotes ex-type strains. 122 Figure 2.9: Bayesian consensus tree of Graphium strains built from gene sequence data of EF1- a. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold. T Denotes ex-type strains. 123 Figure 2.10: Bayesian consensus tree of Graphium strains built from gene sequence data of ITS. Nodes are labeled with bootstrap values/posterior probabilities. Strains isolated from Coptoborus sp. for this study are written in bold. 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Xyleborini (Coleoptera: Curculionidae: Scolytinae) ambrosia beetles occur in forests worldwide and are likely capable of long range dispersal. In less than 20 million years, this group dispersed from Asia to tropical regions of Africa and South America. The phylogeny, taxonomy, and biogeography of one Xyleborus species group which occurs on both continents are reviewed for this study. Based on a well-resolved molecular phylogeny resulting from parsimony, likelihood, and Bayesian analyses of four gene loci, we describe a new monophyletic genus, Xenoxylebora Osborn, Smith & Cognato, gen. nov., for this bicontinental Xyleborus species group with seven Afrotropical and six Neotropical species. Six new species are described: Xenoxylebora pilosa Osborn, Smith & Cognato, sp. nov. from Africa, and Xenoxylebora addenda Osborn, Smith & Cognato, sp. nov., Xenoxylebora calculosa Osborn, Smith & Cognato, sp. nov., Xenoxylebora hystricosa Osborn, Smith & Cognato, sp. nov., Xenoxylebora serrata Osborn, Smith & Cognato, sp. nov., and Xenoxylebora sulcata Osborn, Smith & Cognato, sp. nov., from South America. Seven new combinations from Xyleborus are proposed: Xenoxylebora caudata (Schedl, 1957) comb. nov., Xenoxylebora collarti (Eggers, 1932) comb. nov., Xenoxylebora perdiligens (Schedl, 1937) comb. nov., Xenoxylebora sphenos (Sampson, 1912) comb. nov., Xenoxylebora subcrenulata (Eggers, 1932) comb. nov., and Xenoxylebora syzygii (Nunberg, 1959) comb. nov. from Africa, 137 and Xenoxylebora neosphenos (Schedl, 1976) comb. nov. from South America. One new synonym is proposed: Xenoxylebora sphenos (Sampson, 1912) =Xyleborus tenellus Schedl, 1957 syn. nov.. Descriptions, diagnoses, images, and a key to the identification of all 13 species are provided. The sequence of colonization between Africa and South America is uncertain for Xenoxylebora. Prevailing ocean currents and predominant locality patterns observed for other organisms suggests an African Xenoxylebora origin. However, the phylogeny and biogeographical analyses suggest a possible South American origin for African Xenoxylebora, which is supported by the occurrence of ocean counter currents between the continents and evidence of dispersal from South America to Africa among some plant and arthropod taxa. INTRODUCTION The occurrence of related organisms across multiple continents has received much scientific attention because distributional patterns of species contain important clues about the geological history of landmass movements and the mechanics of evolution. Patterns of biodiversity are complex mosaics often resulting from several historical forces (Upchurch 2007). These include vicariance and long-distance dispersal via wind and water as well as geodispersal whereby previously confined species expand their ranges when geographical barriers erode or disappear (Renner 2004a, Couvreur et al. 2011, Condamine et al. 2012, Praz and Packer 2014, Luo et al. 2020). Extinction and taxon sampling biases also affect the documentation of biodiversity and understanding of the factors responsible for observed geographic patterns of species (Upchurch 2007). Despite the challenge of untangling all elements contributing to the arrangement of biodiversity, evidence suggests that some taxa can travel over oceans between continents. This 138 includes many plants (Langenheim and Lee 1974, Dick et al. 2003, Givnish et al. 2004, Renner 2004b, Cook and Crisp 2005, Won and Renner 2006, Couvreur et al. 2011), vertebrates (Raxworthy et al. 2002, Bond et al. 2015), and arthropods (Erwin 1979, Bonte et al. 2003, Jordal 2015, Gohli et al. 2016, Murray and Heraty 2016, Luo et al. 2020, Eliassen and Jordal 2021). Plants travel across oceans relatively easily as seeds, or rafts of tangled plant material (floating islands) (Renner 2004a, Peck and Perez-Gelabert 2012). For instance, the plant genus Hymenaea (Fabaceae) produces tough, buoyant seedpods that are probably capable of delivering a viable seed after floating in the ocean for several weeks (Langenheim and Lee 1974, de Queiroz 2005). Some animals – especially insects – are also likely to disperse by rafting across oceans because of their close relationships with plants. Peck and Perez-Gelabert (2012) discovered evidence of widespread trans-oceanic dispersal among the beetle genera native to the West Indies. Most of these genera belong to three families (Cerambycidae, Chrysomelidae, and Curculionidae) whose larvae or adults and adults live inside wood, under bark, or are associated other plant structures such as leaves, fruits, and roots. For example, beetles from the weevil tribe Tesserocerini (Curculionidae: Platypodinae) probably arrived in the Neotropics after drifting there from Africa inside Hymenaea seeds (Peris et al. 2015). Scolytine bark and ambrosia beetles (Curculionidae) are preadapted for trans-oceanic travel because most of their life cycles occur inside their woody plant hosts thus presumably protecting them from saltwater and UV radiation. Bark beetles eat tree phloem and live inside or just underneath tree bark. Several species are known to have traveled relatively short distances among Caribbean islands and South and Central America (Kirkendall and Jordal 2006). Other scolytines including the crypturgine genus Aphanarthrum and several species from the tribe Hypoborini have dispersed in this manner among the Macaronesian Islands (Jordal and Hewitt 139 2004) and between Madagascar and the African continent (Jordal 2021a). Trans-ocean rafting carried Araptus (Corthylini) from Central America to Cocos Island (Kirkendall and Jordal 2006), and two clades of Scolytoplatypodini from Africa to Madagascar (Jordal 2013). The fossil species †Electroborus brighti Cognato, 2013 (Hylesinini) from Dominican amber is sister to the African genus Strombophorus and thus presumably evolved from a common ancestor that traveled from Africa to the Americas (Cognato 2013). Several more bark beetle taxa have experienced extreme long-distance dispersal between Africa and the Americas (Figure 1) (Jordal 2012, 2017, 2021b). Ambrosia beetles farm symbiotic fungi as food for larvae and adults in burrows made in the sapwood of plant hosts. This lifestyle has independently evolved in at least 12 scolytine lineages and once in the Platypodinae (Jordal and Cognato 2012, Gohli et al. 2017, Johnson et al. 2018, Pistone et al. 2018). Nutritional fungus farming is associated with high species diversity – especially in the tribe Xyleborini (Jordal et al. 2000, Farrell et al. 2001, Cognato et al. 2011, Gohli et al. 2017) and is likely an asset to long-range habitat expansion. Given that ambrosia beetles only require host plant conditions that can support fungal growth, they are less constrained in their choice for plant hosts compared to bark beetles, which may survive on only certain tree species (Farrell et al. 2001, Malacrinó et al. 2017, Johnson et al. 2018). The hyperdiverse monophyletic ambrosia beetle tribe Xyleborini originated ~20 million years ago in Asia (Cognato et al. 2011, Jordal and Cognato 2012, Johnson et al. 2018) and currently contains ~1260 species (Smith, unpublished). Xyleborini species exhibit a suite of traits that make them especially well-prepared to successfully colonize new land masses. Their association with woody plants means that they have the potential to be included in floating islands that may move between continents. Dispersing females inoculate insipient galleries with 140 fungus they carry with them when they leave their natal colonies. This ensures that food is readily available once a subcortical farm is established. The entire tribe also exhibits haplodiploid inbreeding in which haploid flightless males mate with their diploid sisters inside the burrow before the females disperse. Thus, they do not need to find a mate after arriving to a new environment (Jordal et al. 2000, 2001, Gohli et al. 2016, 2017). These traits allow xyleborine ambrosia beetles to experience low Allee effect thresholds because small colonizing populations are able to produce nutritious food in novel habitats and are less susceptible to inbreeding depression and extinction by genetic drift (Jordal et al. 2001, Peer and Taborsky 2005, Kirkendall and Jordal 2006, Lantschner et al. 2020). Several Xyleborini have been discovered to have completed ancient long-distance dispersal (Kirkendall and Jordal 2006, Gohli et al. 2016, Cognato et al. 2018, Eliassen and Jordal 2021). Xyleborine ambrosia beetles are known from Africa (206 species), and South America (236 species) (Figure 3.1) (Gohli et al. 2016, Eliassen and Jordal 2021, Smith unpublished) however, these species estimates are low and much of the fauna awaits discovery (e.g., Smith et al. 2017, Eliassen and Jordal 2021, Smith and Cognato 2021). Their generalist host needs, inbreeding reproductive system, and the similar tropical climate shared between the Afrotropics and Neotropics allow this group to successfully invade novel habitats with greater ease than other insects (Jordal et al. 2001, Rassati et al. 2016, Lantschner et al. 2020). Xyleborus – the largest Xyleborini genus lives in every biogeographic realm except the Antarctic (Hulcr et al. 2015). Several species within the genus have a history of ancient trans-oceanic dispersal (Kirkendall and Jordal 2006, Gohli et al. 2016, Cognato et al. 2018) and many species have efficiently colonized new habitats via anthropogenic transfer (Haack 2006, Smith et al. 2015, Morgan et al. 2017, Rabaglia et al. 2019). 141 Early researchers identified Xyleborus as a large and confusing genus because of the morphological similarity of many of its members (Hubbard 1897). Indeed, molecular phylogenies have confirmed that Xyleborus is not monophyletic (Jordal et al. 2000, Hulcr et al. 2007, Dole et al. 2010, Cognato et al. 2011) and several new genera have recently been described from species previously designated as Xyleborus (e.g. Hulcr and Cognato 2009, Cognato et al. 2020). In this paper we reconstruct a dated DNA-based phylogeny of African and South American xyleborine species, infer dispersal scenarios for the African and South American species via parametric historical biogeography analyses (eg. Sanmartín 2012), and as a result describe a new genus from Xyleborus with an unusual Afro/Neotropical distribution that presumably experienced a historical bifurcation between Africa and the Americas. METHODS Taxon Sampling Molecular data were obtained from beetle specimens collected in Africa (Cameroon, Kenya, and Uganda), South and Central America (Brazil, Costa Rica, Ecuador, Guyana, Panama, Peru, and Suriname), and North America (United States) (Table 3.1). Given that one species of the suspected new genus was previously considered to be Coptoborus (Wood and Bright 1992), we included a dense sampling of species from this genus along with other Neotropical Xyleborini genera, and Xyleborus species. Several other Xyleborini genera including Ambrosiodmus Anisandrus, Cnestus, Euwallacea, Xyleborinus were used as the outgroup based on prior phylogenetic analyses of the tribe (Cognato et al. 2011, Cognato et al. 2019). In total, we sampled 59 xyleborine specimens (belonging to 52 species in 11 genera). The ingroup included 142 six specimens belonging to the newly described genus (five of the 13 known species) and 37 specimens from Neotropical xyleborine genera (31 species in five genera), and the outgroup consisted of 16 specimens (representing 16 species) belonging to the five genera listed above. Most specimens were excised from their galleries inside infested wood; the remaining individuals were trapped with ethanol-baited bottle traps (Reding et al. 2011) or modified flight intercept traps (Nikulina et al. 2015). All specimens were preserved in in 95−100% ethanol before transportation to the Holistic Insect Systematics Laboratory at Michigan State University where they were stored at -80 ºC. Molecular dataset DNA extraction from the head and pronotum was accomplished using Qiagen DNeasy Blood and Tissue Kits (Qiagen; Hilden, Germany) following manufacturer protocols. Each beetle was dissected to separate the meso-, metathorax and abdomen from the head and pronotum to expose the thoracic muscle tissue for more efficient DNA extraction, and to remove potentially contaminating organisms in the digestive tract. After the DNA was extracted, each head/pronotum was reunited with its corresponding meso-, metathorax and abdomen for future examination. These voucher specimens were deposited in the Albert J. Cook Arthropod Research Collection (MSUC). The resulting DNA templates were used for amplification of four loci: COI, CAD, EF1-α and 28S. Each PCR reaction contained 1.25 units hot star Taq (Qiagen), 0.3μM each forward and reverse primers (Table 3.2), 200μM dNTPs, 1x buffer, 1.75 mM magnesium chloride and 50–5 ng of DNA template. Amplification protocols consisted of 15 minutes initial denaturation (95 ºC) and 38 cycles of 30 seconds denaturation (95 ºC), 30 seconds annealing (50 ºC for COI, 55 ºC for 143 CAD, EF1-α and 28S), and 5 minutes extension (72 ºC). To confirm amplification success and rule out contamination, each reaction was subjected to gel electrophoresis run through a 1.5% agarose gel stained with ethidium bromide. PCR products that were consistent with the fragment size predicted for each locus were cleaned using the ExoSAP-IT enzymatic reagent (Thermo Fisher Scientific; Waltham, Massachusetts, USA). Sanger sequencing was performed at the Michigan State University Research Technology Support Facility using Bigdye terminator 1.1 chemistry (Applied Biosystems; Waltham, Massachusetts, USA). Complementary strands from each Sanger reaction were assembled and trimmed with Sequencher 5.0 build 7082 (Gene Codes Corporation; Ann Arbor, Michigan, USA). Adjustments to the sequences were also made at this time to fix mistaken or unclear base assignments. Sequences generated for this study were submitted to the NCBI GenBank database (Table 3.1). Each locus was aligned with MUSCLE using the default settings (Madeira et al. 2019) and merged into a single, interleaved nexus file containing all four loci. Phylogenetic analyses Models of evolution were determined with PartitionFinder 2.1.1 (Lanfear et al. 2016) following the corrected Akaike information criterion (AICc). A single data block was established for the ribosomal LSU gene 28S, whereas the first, second, and third codon positions of the mitochondrial (COI) and protein-coding genes (CAD and EF1-α) were evaluated separately with linked branch lengths via the ‘all’ search algorithm. PartitionFinder 2.1.1 returned nine substitution models for each of the suggested partitions (Table 3.3). This partition assembly was used for Bayesian phylogenetic inference via MrBayes 3.2.5 (Ronquist and Huelsenbeck 2003). We ran two independent Monte Carlo runs with 40 million 144 generations each using metropolis coupling (one cold chain and three heated chains per run). Trees were sampled every 100th generation, and the first 100,000 trees were discarded (25% burn-in). The remainder were used to build a majority-consensus tree and to determine posterior probabilities. The nine-partition structure was also evaluated with ModelFinder with the MF+MERGE option to possibly merge partitions during maximum likelihood search with IQ- TREE 2.1.3 using nearest-neighbor interchange and 1000 bootstrap pseudoreplications (Table 3.3) (Nguyen et al. 2015, Chernomor et al. 2016, Kalyaanamoorthy et al. 2017). We also used PAUP* 4.0a build 167 (Swofford 2002) to search the combined interleaved data for most parsimonious trees. Gaps were treated as missing and each character as unordered and equally weighted. The heuristic search consisted of 2000 repetitions with random stepwise addition via bisection/reconnection. A bootstrap analysis of 500 pseudoreplications was also conducted using identical heuristic conditions and simple stepwise addition. For all phylogenetic analyses, an outgroup species Anisandrus sayi (Hopkins, 1915) was used to root the resulting trees because it represents the most distantly related xyleborine genus to the ingroup (Cognato et al. 2011). Biogeographical analyses To estimate the geographic origin of ancestral nodes within the new genus, we evaluated 50,000 trees resulting from the Bayesian inference with Statistical Dispersal Extinction Cladogenesis (SDEC) and BioGeoBEARS using the default settings implemented in RASP version 4.2 (Beaulieu et al. 2013, Matzke 2013a, 2013b, 2014, Massana et al. 2015, Yu et al. 2015, R Core Team 2017). The geographic areas of the taxa were based on Wood and Bright (1992) (Table 3.4). Both analyses used the default settings and included all possible paired combinations of the 145 five relevant geographical regions for comparison (Table 3.4). To create a binary consensus tree, we resolved the six polytomies in the Bayesian consensus tree by grouping unresolved clades based on morphological similarity. This did not affect the analysis for the new genus because its corresponding clade is fully resolved (Figure 3.2). Taxonomy Type material, when available, and additional non-type specimens were examined from the following institutions: CNCI—Canadian National Collection of Insects, Ottawa ICB—Instituto de Ciencias Biologicas, Escuela Politécnica Nacional, Quito MSUC—Albert J. Cook Arthropod Research Collection, Michigan State University, East Lansing NHMUK—Natural History Museum, London NHMW—Naturhistorisches Museum Wien, Vienna NMNH—National Museum of Natural History, Smithsonian Institution, Washington, D.C. NZCS—National Zoological Collection of Suriname, Paramaribo RMCA—Musée Royal de l’Afrique Centrale, Tervuren SEMC— Biodiversity Institute & Natural History Museum, The University of Kansas, Lawrence UNMSM—Universidad Nacional Mayor de San Marcos, Lima Each species was photographed using a Visionary Digital Passport II system (Dun Inc; Palmyra, Virginia, USA) using a Canon EOS 5D Mark II (Tokyo, Japan), 65.0 mm Canon Macro photo lens, two Dynalite (Union, New Jersey, USA) MH2015 road flash heads, Dynalite RoadMax MP8 power pack and a Stack Shot (Cognisys, Inc; Traverse City, Michigan, USA). 146 Montage images were assembled using Helicon Focus Mac Pro 6.7.1 (Helicon Soft; Kharkov, Ukraine) and improved using Adobe Photoshop 2020 (Adobe Systems; Mountain View, California, USA). Specimens were examined using Leica (Wetzlar, Germany) MZ125 and MZ16 stereomicroscopes and illuminated with a Shott (Southbridge, Massachusetts, USA) 150W halogen light source (model ACE®1). Length and width of the body, pronotum, and elytra was measured on up to seven specimens per putative species using a Leica MZ6 stereomicroscope. Length was measured at the longest point of the body or structure and width was measured at the widest. Protruding tubercles were included in length measurements. We followed the pronotal and antennal club types proposed by Hulcr et al. (2007) and excluded the pedicel from the number funicle segments. Distribution and host records were obtained from the following publications: Schedl 1963, Browne 1965, Medler 1980, Wood 1982, Beaver and Löyttyniemi 1985, Wood and Bright 1992, Wood 2007, Smith et al. 2017. Previous work has established that COI pairwise differences of >10% are useful for xyleborine species demarcation (Dole et al. 2010, Cognato et al. 2020). We used this intraspecific threshold as well as congruence between the molecular phylogeny and morphological divergence to delineate species. Data availability DNA sequences are available at NCBI GenBank (Table 3.1). The following files were deposited in DRYAD (https://doi.org/10.5061/dryad.zs7h44jc9): NEXUS files with aligned DNA sequences, most parsimonious trees, a consensus tree resulting from the Bayesian analysis, a resolved tree used in the RASP analysis and a Phylip file with aligned DNA sequences. 147 RESULTS Phylogenetic analysis The combined data included six Xyleborus specimens from Africa and South America, 30 specimens from the Neotropical genera (Coptoborus, Dryocoetoides, Sampsonius and Taurodemus), and 23 specimens representing several widespread genera (Ambrosiodmus, Anisandrus, Cnestus, Euwallacea, Xyleborinus, and Xyleborus) (Table 3.1). The four-gene multisequence alignment contained 2463 bp (606 COI, 477 CAD, 373 EF1-α, and 1007 28S), 666 bp of which were parsimony-informative. The heuristic search evaluated over 3.4 billion rearrangements and located 36 most parsimonious trees (score = 4176). Bayesian inference of the treespace reached a final average split frequency standard deviation of 0.003 and produced a consensus tree that has moderate posterior probability (PP) support (0.60–0.89 PP) at the basal nodes and strong PP (>0.95) (Erixon et al. 2003) for most derived nodes (Figure 3.2). Likelihood analysis using nearest-neighbor-interchange located a most likely tree (log-likelihood = - 20108.9524) which was identical to the trees found in the parsimony and Bayesian analyses except for two Euwallacea. However, some bootstrap values were lower compared to parsimony bootstrap values and posterior probabilities (Figure 3.3). Potentially this is an artefact of applying poor models of nucleotide substitution to bootstrapped data sets. In agreement with previous phylogenetic work on Xyleborini, the Neotropical genera Coptoborus, Dryocoetoides, and Sampsonius are monophyletic; together forming a well- supported lineage sister to the widely distributed genus Xyleborus. Six specimens from tropical Africa and the Amazon formed a robust clade: Xenoxylebora neosphenos (previously Xyleborus neosphenos Schedl, 1976) (Ecuador), Xenoxylebora addenda sp. nov. (previously Coptoborus 148 sp.) (Suriname), Xenoxylebora calculosa sp. nov. (previously Xyleborus sp.) (Peru), Xenoxylebora collarti (Eggers, 1932) (previously Xyleborus) (Kenya), and two Xenoxylebora sphenos (Sampson, 1912) (previously Xyleborus) (Uganda and Cameroon). This group has strong posterior probability, maximum likelihood, and parsimony bootstrap support (1.00, 100 and 96, respectively), however its placement within Xyleborini as sister to Taurodemus and Xyleborinus exhibits mixed support (0.84 PP, 51 maximum likelihood bootstrap and no parsimony bootstrap support). Thus, its position is uncertain. Nevertheless, this group clearly represents a separate origin from the other Neotropical genera. This well-supported grouping of one putative Coptoborus specimen with five Xyleborus species necessitates taxonomic revision of this clade. Biogeographical analyses The SDEC analysis indicated that the most likely place of origin of the new genus is the Neotropics (0.66 Neotropics/0.33 Afrotropics and Neotropics). The clade including the African species could have originated in either the Neotropics or Africa (0.81 Neotropics/0.19 Afrotropics and Neotropics (Figure 3.2). The BioGeoBears analysis was largely congruent with these results and differ by eight or fewer reported probabilities (0.69 Neotropics/0.28 Afrotropics and Neotropics, and 0.27 Neotropics/0.72 Afrotropics and Neotropics, respectively). The biogeographical results for the other xyleborines included in this study are not reported because Xyleborini is greatly under-sampled (59/1260 species; 11/42 genera) and details analysis and discussion of these taxa are beyond the scope of this study. 149 Taxonomy Xenoxylebora Osborn, Smith & Cognato gen. nov. Type species. Xenoxylebora neosphenos (Schedl, 1976). Diagnosis. 1.70–2.70 mm long; 2.38–4.60 × as long as wide. Xenoxylebora is distinguished from all other xyleborine genera by the following combination of characters: antennal club obliquely truncate, typically type 1, with segment 1 encircling the anterior face (Figure 3.4.3) or type 2, with segment 1 nearly covering the posterior face; antennal club wider than long (Figure 3.4.3); protibia slender, posterior face unarmed (Figure 3.4.4); mycangial tufts absent; and scutellum small, flush with elytra and elytral apex typically armed by large subquadrate or quadrate tubercles (Figure 3.4.5). Xenoxylebora is superficially similar in appearance to Xyleborinus with which it shares an elongate form, declivital sculpturing and obliquely truncate antennal club (types 1 and 2). It can be distinguished by the scutellum small, flush with the elytra and flat, the lack of mycangial tufts. In Xyleborinus the scutellum is minute, conical, disconnected from elytra and surrounded by a dense tuft of mycangial setae. Xenoxylebora is also similar to some elongate Coptoborus and can be distinguished by the antennal club type 1, without sutures on posterior face, club wider than long, and protibia slender. Coptoborus species have antennal flat club types 3, or 4 (rarely type 2), with two or three sutures on posterior face, club round or longer than wide (Smith and Cognato 2021). Female. 1.70–2.70 mm long; 2.38–4.60 × as long as wide. Body light to dark brown, pronotum anterior slope and elytral declivity often darker. Head: epistoma entire, transverse, weakly sinuate or sinuate, bearing hair-like setae. Frons shagreened, rarely shining, glabrous or sparsely 150 setose. Eyes narrowly or broadly and moderately emarginate. Submentum large, triangular, deeply impressed. Antennal scape short and thick, shorter than club, funicle 4-segmented. Pedicel shorter than funicle. Club circular, obliquely truncate, slightly wider than long, type 1 or 2; segment 1 corneous, weakly concave, occupying basal 1/3 – 1/2, nearly covering or covering posterior face (Figure 3.4.3). Pronotum: 1.00–1.80 × as long as wide. In dorsal view rounded anteriorly with sides parallel in basal 2/3 (type 7); base subtransverse, posterior angles narrowly rounded. In lateral view rounded anteriorly, slope occupying anterior ~1/4; disc much longer than basal slope (type 8). Anterior slope densely covered in broad or narrow asperities and erect hair-like setae; disc shiny to subshiny, finely punctate, glabrous or with erect hair-like setae; lateral margins obliquely costate. Elytra: 1.33 – 2.80 × as long as wide, 1.22–1.53 × as long as pronotum. Scutellum small, flat, flush with elytra. Base transverse, humeral angles rounded. Sides parallel for basal ~ 1/3 – 3/4 then weakly to acutely rounded to apex; apex entire, rounded to acuminate, rarely truncate and bearing two round to quadrate tubercles at sutural apex, one on each elytron, tubercles small and inconspicuous to large and prominent or elongate (Figure 3.4.5). Disc subshiny to strongly shining, rarely dull; strial punctures deep, each bearing a seta the height of the diameter of a puncture, rarely longer; interstriae flat, impunctate or finely punctate, glabrous or with hair-like setae. Declivity gradually or steeply rounded, occupying apical ~1/4–3/5 of elytra, shagreened, subshiny, rarely strongly shagreened or dull; declivital face weakly convex to convex, rarely concave. Striae flat to impressed, glabrous or setose; impunctate, or punctures smaller than those on disc. Interstriae flat to impressed, glabrous or with erect or semi-recumbent setae, bearing various tubercles from inconspicuous round granules to large prominent triangular denticles. Interstriae 1–3 often bearing largest tubercles near summit. Interstriae 4–6 often ending short of posterolateral margin. Interstriae 7 usually with 151 prominent series of tubercles along posterolateral margin. Posterolateral margin usually costate along interstriae 7. Legs: protibiae very slender, outer edge weakly rounded bearing 6–8 socketed denticles, posterior face flat and unarmed (Figure 3.4.4). Meso- and metatibiae flat, evenly rounded, bearing 6–11 socketed denticles. Male. Unknown. Distribution. AFROTROPICAL (Cameroon, Democratic Republic of the Congo, Ghana, Kenya, United Republic of Tanzania) and NEOTROPICAL (Brazil, Ecuador, Guyana, Peru, Suriname). Biology. Schedl (1963) described and illustrated the gallery systems of X. collarti, X. sphenos and X. syzygii (Nunberg, 1959) (as Xyleborus submontanus Schedl, 1960). The cave-tunnel gallery system reported for all these species is comprised of a short unbranched entrance tunnel that leads to one or two irregularly shaped brood chambers in the longitudinal plane. Specimens of the genus have been recorded from a wide variety of hardwood hosts ranging in size from 2– 12 cm (Schedl, 1963). Numerous African specimens have been collected from leaf litter (Schedl 1963). Fungal associates are unknown. Etymology. G. Xeno = strange (referring to the unusual Afrotropical-Neotropical distribution), xyle (G) = wood, and bora (G) = gluttonous (traditionally interpreted as “borer” by scolytine taxonomists). Xenoxylebora addenda Osborn, Smith & Cognato sp. nov. (Figures 3.5.6–3.5.8) Type material. Holotype, female: SURINAME, Sipaliwini [District], 2.977312°N, 55.38500°W, 200 m., Camp 4 (low), Kasikasima, 20–25.III.2012, T. Larson, SR12-0320-TN1, 2012 CI-RAP survey, DNA voucher Cop. neo 1 (NZCS). 152 Diagnosis. 2.00 mm long (n = 1); 2.86 × as long as wide. This species is distinguished by declivital interstriae 1 with denticles larger than those of interstriae 2 or 3, declivital interstriae 2 moderately impressed and appearing weakly bisulcate. Xenoxylebora addenda is similar to X. sulcata sp. nov.. Female. 2.00 mm long (n = 1), 2.86 × as long as wide. Body brown, pronotum anterior slope and elytra darker. Head: epistoma entire, weakly sinuate, bearing hair-like setae. Frons shagreened, sparsely setose. Eyes narrowly and moderately emarginate. Club type 2, segment 1 corneous, weakly concave, occupying basal 1/3, nearly covering posterior face. Pronotum: 1.29 × as long as wide. Anterior slope with dense, broad, coarse asperities; disc finely punctate, with abundant erect hair-like setae. Elytra: 1.57 × as long as wide, 1.22 × as long as pronotum. Sides parallel for basal ~1/2 then rounded to apex, apex entire, bearing two small, round sutural tubercles as large as denticles on interstriae 7. Disc strongly shining, strial punctures deep, each with a recumbent seta the height of the diameter of a puncture; interstriae finely punctate, with two confused rows of hair-like setae slightly longer than width of an interstria. Declivity gradually rounded, occupying ~1/2 of elytra, shagreened, subshiny; declivital face weakly convex; striae bearing recumbent hair-like setae as long as those on disc, punctures smaller than those on disc, striae 1 and interstriae 2 moderately impressed; interstriae bearing two confused rows of semi- recumbent to erect hair-like setae slightly longer than width of an interstria; interstriae 1–3 with small uniformly spaced granules, those on interstriae 1 larger than those on 2 and 3; interstriae 7 with larger denticles. Posterolateral margin costate and bearing tubercles along interstriae 7. Legs: protibiae with six socketed denticles. Metatibiae with eight socketed denticles. Male. Unknown. Distribution. NEOTROPICAL: Suriname (Sipaliwini). 153 Biology. The holotype was collected in a flight intercept trap. Etymology. L. addenda = to be added, referring to the late identification of this species, necessitating its addition to the manuscript. An adjective. Xenoxylebora calculosa Osborn, Smith & Cognato sp. nov. (Figures 3.5.9–3.5.11) Type material. Holotype, female: PERU, Madre de Dios, Los Amigos Biological Station, CM2, GPS: S12.4492’ W70.2517’, 17–18.V.2008, [S. M.] Smith, [J.] Hulcr, sample Peru 81, ex 9 cm diameter trunk (UNMSM). Six paratypes, female: PERU, as holotype, except Peru 74, ex 2 cm diameter branch (3, MSUC; 1, NHML; 1, NMNH); as previous except: SMS 111, Coptoborus neosphenos, 16.IX.2011 [DNA voucher] (1, MSUC). Diagnosis. 1.90–2.10 mm long (mean = 1.96 mm, n = 7); 2.71–3.00 × as long as wide. This species is distinguished by the tubercles of interstriae 1–3 subequal, declivital striae 2 and interstriae 2 weakly impressed, elytral disc subequal to length of elytral declivity and declivital face shining. Xenoxylebora calculosa is similar to X. serrata sp. nov. and X. neosphenos. Female. 1.90–2.10 mm long (mean = 1.96 mm, n = 7); 2.71–3.00 × as long as wide (holotype 1.90 mm long; 2.17 × as long as wide). Body light brown, pronotum anterior slope and elytra darker. Head: epistoma entire, sinuate, bearing hair-like setae. Frons shagreened, glabrous. Eyes narrowly and moderately emarginate. Club type 2, segment 1 corneous, weakly concave, occupying basal 1/3, nearly covering posterior face. Pronotum: 1.14–1.29 × as long as wide (holotype 1.14 × as long as wide). Anterior slope with dense, broad, coarse asperities; disc sparely covered in minute setae, almost glabrous. Elytra: 1.43–1.71 × as long as wide (holotype 1.57 × as long as wide), 1.37 × as long as pronotum. Sides parallel for basal ~1/2–2/3 then 154 rounded to apex, apex entire, bearing two small, round sutural tubercles, as large as denticles on interstriae 7. Disc strongly shining; strial punctures deep each with a minute seta the length of a diameter of a puncture; interstriae glabrous, punctures biseriate. Declivity gradually rounded, occupying ~1/2 of elytra, shiny; declivital face weakly convex; striae 2 weakly impressed with recumbent hair-like setae longer than those on disc, punctures smaller than those on the disc; interstriae bearing semi-recumbent bristle-like setae slightly longer than the width of an interstria; interstriae 2 weakly impressed; interstriae 1–3 with small, uniformly spaced granules, each ~1/2 the width of an interstria, and bearing a long, erect seta; interstriae 7 with large denticles near the declivital base, gradually transitioning to large, round tubercles at apex. Posterolateral margin costate and bearing moderate subquadrate tubercles along interstriae 7. Legs: protibiae with six socketed denticles. Meso- and metatibiae with six and eight socketed denticles, respectively. Male. Unknown. Distribution. NEOTROPICAL: Peru (Madre de Dios). Biology. Specimens collected from canopy fogging and extracted from infested wood 2–9 cm in diameter. Etymology. L. calculosa = pebbly, referring to the texture and appearance of small denticles on the declivity. An adjective. Xenoxylebora caudata (Schedl, 1957) comb. nov. (Figures 3.5.12–3.5.14) Xyleborus caudatus Schedl, 1957: 110, orig. spelling. 155 Material Examined. Holotype, female: Congo Belge [= DEMOCRATIC REPUBLIC OF THE CONGO], [Tshopo Prov.], Yangambi, 10.X.1952, Dr. [K.E.] Schedl. Paratype, female: as holotype, except: 14.X.1952 (1). (Holotype and paratype in RMCA). Diagnosis. 2.00 mm long (mean = 2.00 mm, n = 2); 2.86 × as long as wide. This species is distinguished by the tubercles of declivital interstriae 3 larger and much more prominent than those of interstriae 1 or 2, sutural tubercles on elytral apex distinctly elongated, laterally angled, and closely grouped to three large denticles on interstriae 1. Xenoxylebora caudata is similar to X. hystricosa sp. nov., X. perdiligens (Schedl, 1937) and X. sphenos. Distribution. AFROTROPICAL: Democratic Republic of the Congo (South Kivu, Tschopo), Zambia. Biology. Breeds in small twigs and branches (Beaver and Löyttyniemi 1985). Host plants: Anacardiaceae (Antrocaryon), Fabaceae (Pterocarpus), Irvingiaceae (Klainedoxa), Urticaceae (Musanga). Xenoxylebora collarti (Eggers, 1932) comb. nov. (Figures 3.5.15–3.5.17) Xyleborus collarti Eggers, 1932: 300. Xyleborinus collarti (Eggers): Wood and Bright 1992: 806. Xyleborus collarti Eggers: Hulcr et al. 2007: 577. Xyleborus semipilosus Eggers, 1932: 300. Synonymy: Schedl 1963: 490. Material Examined. Holoytpe, female: X. collarti: [DEMOCRATIC REPUBLIC OF THE CONGO], Forêt de Kawa, 22.IV.[19]24, A. Collart, USNMENT 01547104 (NMNH); images examined. Holotype, female: X. semipilosus, [DEMOCRATIC REPUBLIC OF THE CONGO], 156 Katanga [= Tanganyika + Haut-Lomami + Lualabba + Haut-Katanga Prov.], Lufudizi, 22.IX.1924, C. Seydel, Musee du Congo (RMCA). Seven females designated X. collarti: KENYA, Kakamega District [= County], Isecheno, Isecheno Forest Reserve, 13.II.2002, R.R. Snelling, ex sifted litter #02-47, SM0677729 (1); as previous except: Yala River Forest Reserve, 0.204ºN 34.873ºE, 1450–1470 m, 15.II.2002, ex sifted litter between buttresses #02-058, SM0668771, SMS 373 [DNA voucher] (1), SM0698653 (1), SM0698704, SMS 372 [DNA voucher] (1); as previous except: 1450 m, 28.II.2002, #02-096, SM0678399 (1); as previous except: 8.III.2002, ex sifted Ficus litter between buttresses #02-0111, SM0698291 (1), SM0698280 (1) (all in SEMC). Diagnosis. 2.20–2.30 mm long (mean = 2.26 mm; n = 5); 2.81–3.29 × as long as wide. This species is distinguished by the tubercles of interstriae 1–3 subequal and declivity convex with a gradual slope. Xenoxylebora collarti is similar to X. pilosa sp. nov., and X. subcrenulata (Eggers, 1932). Distribution. AFROTROPICAL: Angola, Cameroon, Côte d’Ivoire, Democratic Republic of the Congo (Haut-Katanga, Haut-Lomami, Lualabba), Ghana, Kenya (Kakamega), Nigeria, United Republic of Tanzania. Biology. Xenoxylebora collarti is reported to colonize branches ranging from 2–5 cm in diameter. Females construct a cave-tunnel gallery system comprised of a short unbranched entrance tunnel that leads to one irregularly shaped brood chamber in the longitudinal plane. (Schedl, 1963). Host plants: Apocynaceae (Conopharyngia), Fabaceae (Acacia, Millettia), Hypericaceae (Haronga), Meliaceae (Trichilia, Turraeanthus), Myrtacae (Syzygium), Rubiaceae (Cinchona, Galiniera), Solanaceae (Solanum). Specimens designated X. collarti were sifted from Ficus leaf litter. 157 Comments. Schedl (1963) placed X. semipilosus in synonymy without comment. Strangely both species were described in same publication and page (Eggers, 1932: 300) however, Eggers failed to recognize the species were conspecific and compared the form of X. semipilosus to that of Xyleborinus saxesenii (Ratzeburg, 1837) and that of Xenoxylebora collarti to Fraudatrix melas (Eggers, 1927). Both species are clearly conspecific and we agree with the synonymy of X. semipilosus. Xenoxylebora hystricosa Osborn, Smith & Cognato sp. nov. (Figures 3.5.18–3.5.20) Type material. Holotype, female: BRAZIL, Amazonas, 60 K[ilo]m[eters] N[orth] Manaus, Fazenda Esteio, ZF-3 Km-23, 10.V.1985, B.C. Klein, ex arm Malaise (NMNH). Paratypes, 3 females: BRAZIL, as holotype, except: 8.III.1985 (1); 14.V.1986 (1, NMNH); SURINAME, Mariwijine, Palumeu, 160 m, 7–8.VIII.1999, Z.H. Falin, ex FIT (1, CNCI). Diagnosis. 2.15–2.30 mm long (mean = 2.24 mm, n = 4); 2.44–2.88 × as long as wide. This species is distinguished by declivital interstriae 1 and 2 unarmed, spines on interstriae 3 very prominent, significantly larger than those on interstriae 4–7. Xenoxylebora hystricosa is similar to X. caudata, X. perdiligens and X. sphenos. Female. 2.15–2.30 mm long (mean = 2.24 mm, n = 4); 2.44–2.88 × as long as wide (holotype 2.20 mm long; 2.44 × as long as wide). Body light brown, pronotum anterior slope and elytra slightly darker. Head: epistoma entire, weakly sinuate, with hair-like setae. Frons shining, sparsely setose, finely punctate. Eyes broadly and moderately emarginate. Club type 2, segment 1 corneous, weakly concave, occupying basal 1/3 of segment 1, nearly covering posterior face. Pronotum: 1.13–1.43 × as long as wide (holotype 1.33 × as long as wide). Anterior slope with 158 dense, broad, coarse asperities; disc shining, finely punctate with erect hair-like setae. Elytra: 1.33–1.75 × as long as wide (holotype 1.33 × as long as wide), 1.29 × as long as pronotum. Sides parallel for basal ~ 1/3–1/2 then attenuate to apex, apex entire, bearing two subquadrate sutural tubercles similar in size to denticles on interstriae 3. Disc strongly shining, strial punctures deep each with a minute seta the length of a diameter of a puncture; interstriae finely punctate with single row of minute erect hair-like setae ~1 ½ × as long as the width of an interstria. Declivity gradually rounded, occupying ~1/2 of elytra, shagreened, subshiny; declivital face weakly convex; striae flat, with recumbent hair-like setae, thicker and as long as those on disc; punctures smaller than those on disc; interstriae flat with erect hair-like setae ~2 × as long as the width of an interstria; interstriae 1 and 2 unarmed on declivital face, interstriae 1 with two or three denticles at the summit; interstriae 3 with large, distinct, uniformly spaced denticles ~2 × as tall as the width of an interstria extending from declivital base to apex and bearing two similarly sized subquadrate sutural tubercles at apex; interstriae 4–7 with smaller denticles ~2/3 the width of an interstria. Posterolateral margin costate and bearing denticles along interstriae 7. Legs: protibiae with six socketed denticles. Meso- and metatibiae with nine socketed denticles. Male. Unknown. Distribution. NEOTROPICAL: Brazil (Amazonas), Suriname (Marowijine). Biology. Specimens were passively collected in Malaise and FIT traps. Etymology. L. hystricosa = thorny, referring to the appearance of the distinctive row of large denticles on declivital interstriae 3. An adjective. Comments. Specimens collected in malaise and flight intercept traps. 159 Xenoxylebora neosphenos (Schedl, 1976) comb. nov. (Figures 3.6.21–3.6.23) Xyleborus neosphenos Schedl, 1976: 76. Coptoborus neosphenos (Schedl): Wood and Bright 1992: 663. Xyleborus neosphenos Schedl: Smith and Cognato 2021: 620. Material Examined. Holotype, female: BRAZIL, Rondônia, Vilhena, XI.1973, M. Alvarenga (NHMW). ECUADOR, Napo Prov. [= Orellana Prov.], Res[erva]. Ethnica Waorani, 1 km S. Okone Gare Camp, Trans[ect]. Ent[omology]., 00°39'10"S, 076°26'W, January 1994, T. L. Erwin et al., 220 m, insecticidal fogging, terra firme forest, trans[ect] 1, sta[tion] 5, Erwin-lot # 594 (1, ICB); Orellana Prov., Tiputini Biodiversity Station, S00°38.189' W76°08.965', 223 m, 3– 9.VI.2011, S.M. Smith, SMS 102 Coptoborus neosphenos, 16.IX.2011 [DNA voucher] (1, MSUC). PERU, Madre de Dios, Los Amigos Biological Station, CM2, GPS: S12.4492’ W70.2517’, 17–18.V.2008, [S.M.] Smith, [J.] Hulcr, Peru 74, ex 2 cm diameter (MSUC); as previous except Peru 88c, ex 3 cm diameter twig (MSUC). Diagnosis. 1.90–2.10 mm long (mean = 2.00 mm, n = 2); 2.71–3.00 × as long as wide. This species is distinguished by the tubercles of interstriae 1–3 subequal, acuminate elytral apex, and tubercles on elytral apex subquadrate and very prominent. Xenoxylebora neosphenos is similar to X. calculosa and X. serrata. Distribution. NEOTROPICAL: Brazil (Rondônia, Mato Grosso), Ecuador (Orellana), Peru (Cusco, Madre de Dios). Biology. One specimen (NMNH) from Mato Grosso (E78) was collected from small 3.3 cm diameter uprooted tree of Pouteria sp. (Sapotaceae) in gallery forest (Roger A. Beaver, personal 160 communication). Specimens have also been collected from canopy fogging and extracted from additional infested wood 2–3 cm in diameter. Xenoxylebora perdiligens (Schedl, 1937) comb. nov. (Figures 3.6.24–3.6.26) Xyleborus perdiligens Schedl, 1937: 399. Material Examined. Lectotype, female: [UNITED REPUBLIC OF TANZANIA], Urw.[ald] hint.[er] d[em] Randbg. [= Randbgebirge] N.W. Tanganjikasees, 18[00]–2200 m, S[ammler] Grauer (NHMW). Nine females: Congo [= DEMOCRATOC REPUBLIC OF THE CONGO], [North Kivu Prov.], Dorsale de Lubero, Mt. Muleke, June/July 1963, M. J. Célis, Coll. Mus. Tervuren (3, RMCA); as previous except: P.N.A, Massif Ruwenzori, Kalonge, 2.IX.1952, P. Vanschuytbroeck & J. Kekenbosch, alt. 2210 m, 922–26 (3, RMCA); as previous except 9– 11.VIII.1952, 741–42 (1, RMCA); Lubero, route Kimbulu, June 1954, R. P. M. J. Célis, alt. 18300 m, Tamisage Tamisage de terreau sous fougéres arbor; (1, RMCA); as previous except ruiss prés de Kimbulu, alt. 1750 m, (1, RMCA). Diagnosis. 2.5–2.7 mm long (mean = 2.56 mm, n = 8); 2.78–3.38 × as long a wide. This species is distinguished by the tubercles of declivital interstriae 3 larger and much more prominent than those of interstriae 1 or 2, declivital interstriae 1 and 2 sparsely granulate, distance between granules equal to width of at least three granules, and declivital slope very steep, occupying apical quarter of elytra. Xenoxylebora perdiligens is similar to X. caudata, X. hystricosa and X. sphenos. Distribution. AFROTROPICAL: Democratic Republic of the Congo (North Kivu), United Republic of Tanzania (Taganyika). 161 Biology. Specimens were collected by sieving soil and sifting leaf litter under tree ferns. Comments. A series large of X. perdiligens (RMCA; listed above) was incorrectly identified as X. tenellus by M. Nunberg and F. G. Browne. Browne however referred to the specimens and “tenellus Schedl large form”. It is probable other specimens of X. perdiligens are misidentified as X. tenellus in other collections. Xenoxylebora pilosa Osborn, Smith & Cognato sp. nov. (Figures 3.6.27–3.6.29) Type material. Holotype, female: Tanganyika Terr[itory]. [= United Republic of Tanzania]s, [Morogoro Region], Bunduki, Uluguru M[oun]t[ain]s, 1500 m, gorge Mungula, 1–6.V.1957, Mission Zoolog. I.R.S.A.C. en Afrique orientale, P. Basilewsky et N. LeLeup, ex forêt transition dans l’humus (RMCA). Diagnosis. 2.20 mm long (n = 1); 3.14 × as long as wide. This species is distinguished by declivital interstriae 1 and 3 bearing small, round granules, interstriae 2 with very minute granules, almost indistinct, and declivity convex with a gradual slope. Xenoxylebora pilosa is similar to X. collarti, and X. subcrenulata. Female. 2.20 mm long (n = 1); 3.14 × as long as wide. Body medium brown, pronotum anterior slope and elytra darker. Head: epistoma entire, sinuate, with hair-like setae. Frons shagreened, sparsely setose. Eyes narrowly and moderately emarginate. Club type 2, segment 1 corneous, weakly concave, nearly covering posterior face. Pronotum: 1.29 × as long as wide. Anterior slope with dense broad, coarse asperities; disc subshiny, finely punctate, with erect hair-like setae. Elytra: 1.86 × as long as wide, 1.44 × as long as pronotum. Sides parallel for basal ~2/3 then broadly rounded to apex, apex entire, bearing two round sutural tubercles larger than other 162 declivital tubercles. Disc strongly shining; strial punctures deep, each with a seta the height of the distance between punctures; interstriae finely punctate, with two confused rows of hair-like setae slightly longer than width of interstriae. Declivity gradually rounded, occupying ~1/3 of elytra, shagreened, subshiny; declivital face convex, highly obscured by dense setae; striae flat with semi-erect hair-like setae ~3/4 as long as interstrial setae; punctures slightly smaller than those on disc; interstriae flat with two confused rows of erect bristle-like setae as long as the width of an interstria; interstriae 1 with two rows of hair-like setae, small granules and three denticles as tall as the width of an interstria close to apex; interstriae 2 bearing very minute granules; interstriae 3–6 with small, rounded granules; interstriae 7 with slightly larger rounded granules ~ ½ as tall as the width of an interstria. Posterolateral margin costate and bearing tubercles along interstriae 7. Legs: protibiae with seven socketed denticles. Meso- and metatibiae with eight and seven socketed denticles, respectively. Male. Unknown. Distribution. AFROTROPICAL: Democratic Republic of the Congo (Tanganyika). Biology. Specimens were collected from humus in a forest transition zone. Etymology. L. pilosa = hairy, referring to the abundant hair-like setae covering the elytra. An adjective. Comments. The holotype was previously identified as Xyleborus semipilosus (= Xenoxylebora collarti) by M. Nunberg. It is probable that other specimens of this species have been misidentified as Xyleborus semipilosus in collections. 163 Xenoxylebora serrata Osborn, Smith & Cognato sp. nov. (Figures 3.6.30–3.6.32) Type material. Holotype, female: GUYANA, [Potaro-Siparuni Region = Region 8], Iwokrama Forest, Turtle M[oun]t[ain]n, GPS: N04.44.081’ W058.42.830’, Guy 33, 4–9.III.2007, McCall, [A. I.] Cognato, [J.] Hulcr, [S. M.] Smith, [S.] Dole (MSUC). Diagnosis. 1.70 mm long (n = 1); 2.83 × as long as wide. This species is distinguished by the tubercles of interstriae 1–3 subequal, declivital striae 2 and interstriae 2 weakly impressed, elytral disc shorter than length of elytral declivity and declivital face shagreened. Xenoxylebora serrata is similar to X. calculosa and X. neosphenos. Female. 1.70 mm long (n = 1); 2.83 × as long as wide. Body light brown, pronotum anterior slope and elytra distinctly darker. Head: epistoma entire, weakly sinuate, with hair-like setae. Frons shagreened, sparsely setose. Eyes broadly and moderately emarginate. Club type 1, segment 1 corneous, weakly concave, occupying basal 1/3, encircling anterior face. Pronotum: 1.17 × as long as wide. Anterior slope with dense, broad, coarse asperities; disc shining, finely punctate and glabrous. Elytra: 2.00 × as long as wide, 1.43 × as long as pronotum. Sides parallel for basal ~2/3 then broadly rounded to apex, apex entire, bearing two round sutural tubercles as large as denticles on interstriae 7. Disc strongly shining; strial punctures deep, each with a minute seta the length of a diameter of a puncture; interstriae finely punctate, glabrous. Declivity gradually rounded, occupying ~3/5 of elytra, shagreened, subshiny; declivital face weakly convex; striae bearing with recumbent hair-like setae; punctures smaller than those on disc; striae 2 weakly impressed; interstriae bearing semi-recumbent bristle-like setae slightly longer than the width of an interstria; interstriae 2 weakly impressed; interstriae 1–4 with evenly spaced minute granules; interstriae 5 with four denticles close to summit, ~1/2 as tall as the width of an 164 interstria diminishing to small granules toward apex; interstriae 6 with rounded tubercles smaller than those on interstriae 7, diminishing in size toward apex; interstriae 7 with large, round tubercles about as tall as the width of an interstria. Posterolateral margin costate and bearing denticles along interstriae 7. Legs: protibiae with six socketed denticles. Meso- and metatibiae with nine and eight socketed denticles, respectively. Male. Unknown. Distribution. NEOTROPICAL: Guyana (Potaro-Siparuni). Biology. Unknown. Etymology. L. serrata = toothed like a saw, named in reference to the appearance of the denticles on declivital interstriae 7. An adjective. Xenoxylebora sphenos (Sampson, 1912) comb. nov. (Figures 3.6.33–3.6.35) Xyleborus sphenos Sampson, 1912: 247. Xyleborus perdiligens diligens Schedl, 1954: 79. Synonymy: Wood and Bright 1992: 775. Xyleborus montanus tenellus Schedl, 1957: 107. syn. nov. Material Examined. Lectotype, X. perdiligens diligens, female: Gold Coast [= GHANA], [Eastern Region], Mpraeso, 23.I.1947, G.H. Thompson (NHMW); paralectotype, female, as lectotype (NHMUK). Holotype X. montanus tenellus, female: Congo Belge [= DEMOCRATIC REPUBLIC OF THE CONGO], [Tshopo Prov.], Yangambi, 12.IX.1952, Dr. [K.E.] Schedl, Nr. 852 (RMCA). Six females designated X. sphenos: CAMEROON, Southwest Region, Limbe, 22.IX.2007, B. Jordal, ex EtOH trap, (3, MSUC); as previous except: SMS 368 [DNA voucher]. Congo Belge [= DEMOCRATIC REPUBLIC OF THE CONGO], [South Kivu Prov.], Hembe- 165 Bitale, 8.VIII.1952, Dr. [K.E.] Schedl, ex s. 593 (1, NHMW). KENYA, Kakamega District, Isecheno, Isecheno For.[est] Res.[ersve], 0.24°N, 34.85°E, 1600 m, 13.II.2002, R.R. Snelling, sifted litter #02–044, SM0677385 (1 SEMC). 6 females designated X. tenellus (RMCA): Congo Belge [= DEMOCRATIC REPUBLIC OF THE CONGO], [North Kivu Prov.], P.N.A, Massif Ruwenzori, Kalonge, 2210 m, 2.IX.1952, P. Vanschuytbroeck & J. Kekenbosch, 922–26 (2); as previous except: 31.VII-3–5.VIII.1952, 688–69 (1). Face N. Ruwenzori, Kikura, Vallée de la Kafuko, 2000 m, P. N. Virunga, VII/VIII.[19]74, R.P., M. Lejeune, ex dans bois morts (1); as previous except: sol suspend (1). Dorsale de Lubero, Mt. Muleke, JVI/VII. 1963, M.J. Célis (1). Diagnosis. 1.70–2.40 mm long (mean = 2.09 mm, n = 14); 2.38–4.60 × as long as wide. This species is distinguished by the tubercles of declivital interstriae 3 larger and much more prominent than those of interstriae 1 or 2, declivital interstriae 1 and 2 densely granulate, granules subcontiguous, declivital slope gradual, and occupying apical third of elytra. Xenoxylebora sphenos is similar to X. caudata, X. perdiligens, and X. hystricosa. Distribution. AFROTROPICAL: Cameroon (Southwest), Democratic Republic of the Congo (North Kivu, South Kivu, Tshopo), Ghana (Eastern), Kenya (Kakamega), Uganda. Biology. Xenoxylebora sphenos is reported to colonize branches ranging from 2–7 cm in diameter. Females construct a cave-tunnel gallery system comprised of a short unbranched entrance tunnel that leads to one or two irregularly shaped brood chamber in the longitudinal plane. Females deposit eggs in a single heap and brood size ranges from 27–38 individuals (Schedl, 1963). Specimens were collected from liter sifting and alcohol trapping. Host plants: Clusiaceae (Pentadesma), Fabaceae (Albizzia), Malpighiaceae (Acridocarpus), Meliaceae (Lovoa, Turraeanthus). 166 Comments. The holotype of X. sphenos (NHMUK) is not on its point and is likely lost. Our concept of X. sphenos is based on Sampson’s description and specimens independently identified by R. A. Beaver and B. H. Jordal. Xyleborus montanus tenellus Schedl, 1957 is here synonymized with X. sphenos because the specimens examined are morphologically indistinguishable. An additional investigation utilizing additional specimens as part of a molecular analysis will be required to further assess species limits. Xenoxylebora subcrenulata (Eggers, 1932) comb. nov. (Figures 3.7.36–3.7.38) Xyleborus subcrenulatus Eggers, 1932: 301, orig. spelling. Type material. Holotype, female: Congostaat [= DEMOCRATIC REPUBLIC OF THE CONGO], [Kongo Central Prov.], Madimba, 15.iv.[19]30, A. Collart, USNMENT 01547123 (NMNH); images examined. Diagnosis. 1.8 mm long (n = 1); 2.89 × as long as wide. This species is distinguished by the tubercles of interstriae 1–3 subequal and declivity flat with a steep slope. Xenoxylebora subcrenulata is similar to X. collarti and X. pilosa. Distribution. AFROTROPICAL: Democratic Republic of the Congo (Kongo Central). Biology. Unknown. Xenoxylebora sulcata Osborn, Smith & Cognato sp. nov. (Figures 3.7.39–3.7.41) Type material. Holotype, female: ECUADOR, Napo Prov. [= Orellana Prov.], Res[erva]. Ethnica Waorani, 1 km S. Okone Gare Camp, Trans[ect]. Ent[omology]., 00°39'10"S, 076°26'W, 220 m, 167 January 1994, T.L. Erwin et al., insecticidal fogging, terra firme forest, trans[ect] 1, sta[tion] 6, Erwin-lot # 595 (ICB). Diagnosis. 1.90 mm long (n = 1); 2.71 × as long as wide. This species is distinguished by the tubercles of interstriae 1–3 subequal, declivity sulcate between striae 1 and interstriae 4, and declivity appearing bisulcate. Xenoxylebora sulcata is similar to X. addenda. Female. 1.90 mm long (n = 1); 2.71 × as long as wide. Body light brown, elytra slightly darker. Head: epistoma entire, transverse, with hair-like setae. Frons shagreened, sparsely setose. Eyes broadly and moderately emarginate. Club type 1, segment 1 corneous, weakly concave, occupying basal 1/3, encircling anterior face. Pronotum: 1.14 × as long as wide. Anterior slope with dense, broad, coarse asperities; disc shining, finely punctate with erect hair-like setae. Elytra: 1.57 × as long as wide, 2.88 × as long as pronotum. Sides parallel for basal ~2/3 then rounded to apex, apex entire, bearing two small, round sutural tubercles as large as denticles on interstriae 7. Disc subshiny, strial punctures small and shallow, each with one minute recumbent seta the height of a diameter of a puncture; interstriae with two confused rows of hair-like setae ~1 ½ × as tall as the width of an interstria. Declivity gradually rounded, occupying ~1/2 of elytra, shagreened, subshiny; declivital face bisulcate between striae 1 and interstriae 4; striae with recumbent hair-like setae longer than those on disc; punctures smaller than those on disc; interstriae with semi-recumbent hair-like setae slightly longer than the width of an interstria; interstriae 2 and 3 impressed; interstriae 1–3 with small, uniformly spaced granules, each ~1/2 the width of an interstria, and bearing a long, erect seta; interstriae 7 with large denticles near the declivital base, gradually transitioning to large, round tubercles at apex. Posterolateral margin costate and bearing denticles and tubercles along interstriae 7. Legs: pro-, meso-, and metatibiae with seven socketed denticles each. 168 Male. Unknown. Distribution. NEOTROPICAL: Ecuador (Orellana). Biology. The holotype was collected by canopy fogging. Etymology. L. sulcata = furrowed, referring to the sulcate declivity. An adjective. Xenoxylebora syzygii (Nunberg, 1959) comb. nov. (Figures 3.7.42–3.7.44) Xyleborus montanus Schedl, 1957: 106. Preoccupied by Niisima 1910. Xyleborus syzygii Nunberg, 1959: 167. Xyleborus submontanus Schedl, 1960: 106. Unnecessary replacement name. Material Examined. Holotype, female: Congo Belge [= DEMOCRATIC REPUBLIC OF THE CONGO], [South Kivu Prov.], Hembe-Bitale, 15.VIII.1952, Dr. [K.E.] Schedl. Paratypes, 3 females: as holotype. (Holotype and paratypes in RMCA). Diagnosis. 2.00–2.05 mm long (mean = 2.01, n = 4); 2.86–2.93 × as long as wide. This species is distinguished by the declivity obliquely truncate, declivital face concave between suture and interstriae 4, and declivital interstriae 1–3 bearing two or three confused rows of round equally sized sub-contiguous granules. Distribution. AFROTROPICAL: Democratic Republic of the Congo (South Kivu). Biology. Xenoxylebora syzygii is reported to colonize branches ranging from 3–12 cm in diameter. Females construct a cave-tunnel gallery system comprised of a short unbranched entrance tunnel that leads to one irregularly shaped brood chamber in the longitudinal plane. Females deposit eggs in a single heap and brood size ranges from 27–38 individuals (Schedl, 1963). Host plants: Euphorbiaceae (Alchornea), Moraceae (Ficus), Myrtaceae (Syzygium). 169 Key to Xenoxylebora species (females only) 1 Tubercles on declivital interstriae 3 much more prominent than those of interstriae 1 or 2 (if present) (Figures 3.5.18, 3.6.24) … 2 - Tubercles on declivital interstriae 3 of similar size as those of interstriae 1 and 2 or tubercles of interstriae 1 larger (Figures 3.5.9, 3.7.39) … 5 2 Sutural tubercles on elytral apex distinctly elongated, laterally angled, and closely grouped to three large denticles on interstriae 1 (Figures 3.5.12–3.5.14) … caudata - Sutural tubercles on elytral apex large but not elongated or laterally angled, and widely separated from subapical tubercles (Figures 3.5.18, 3.6.24) …3 3 Declivital interstriae 1 and 2 unarmed, spines on interstriae 3 very prominent, significantly larger than those on interstriae 4–7 (Figures 3.5.18–3.5.20); Neotropical…hystricosa sp. nov. - Declivital interstriae 1 and/or 2 granulate, denticles on interstriae 3 moderate or similar size to those on interstriae 4–7 (Figures 3.6.24, 3.6.33); Afrotropical…4 4 Declivital interstriae 1 and 2 sparsely granulate, distance between granules equal to width of at least three granules; declivital slope very steep, occupying apical quarter of elytra (Figures 3.6.24–3.6.26) … perdiligens - Declivital interstriae 1 and 2 densely granulate, granules subcontiguous; declivital slope gradual, occupying apical third of elytra (Figures 3.6.33–3.6.35) …sphenos 170 5 Declivity obliquely truncate; declivital face concave between suture and interstriae 4; declivital interstriae 1–3 bearing two or three confused rows of round, sub-contiguous granules (Figs 3.7.42–3.7.44) … syzygii - Declivity rounded (Figure 3.5.9) or acuminate (Figure 3.6.21); declivital face weakly convex (Figure 3.6.22) to convex (Figure 3.5.7), interstriae flat or impressed; declivital interstriae with uniseriate granules or denticles …6 6 Declivity with one or more striae or interstriae clearly impressed (Figures 3.5.6, 3.7.39); Neotropical …7 - Declivity flat (Figure 3.7.37) to convex (Figure 3.5.15); Afrotropical…11 7 Elytral apex acutely acuminate in dorsal profile; tubercles on elytral apex subquadrate and very prominent (Figures 3.6.21–3.6.23) …neosphenos - Elytral apex rounded in dorsal profile; sutural tubercles on elytral apex round and not significantly enlarged (Figures 3.5.9, 3.6.30) …8 8 Declivital striae 1 and interstriae 2 moderately (Figure 3.5.6) or deeply impressed, appearing bisulcate (Figure 3.7.39) …9 - Declivital striae 2 and interstriae 2 weakly impressed (Figures 3.5.9, 3.6.30) …10 9 Declivity sulcate between striae 1 and interstriae 4, appearing bisulcate; denticles of interstriae 1–3 subequal (Figures 3.7.39–3.7.41) …sulcata sp. nov. 171 - Declivital interstriae 2 weakly impressed; denticles of interstriae 1 larger than those of interstriae 2 and 3 (Figures 3.5.6–3.5.8) …addenda sp. nov. 10 Elytral disc shorter than length of elytral declivity; declivital face shagreened (Figures 3.6.30–3.6.32) …serrata sp. nov. - Elytral disc subequal to length of elytral declivity; declivital face shining (Figures 3.5.9–3.5.11) …calculosa sp. nov. 11 Declivity flat, declivital slope steep (Figures 3.7.36–3.7.38) …subcrenulata - Declivity convex, declivital slope gradual (Figure 3.5.16) …12 12 Declivital interstriae 1–3 bearing denticles of consistent size (Figures 3.5.15–3.5.17) …collarti - Declivital interstriae 1 and 3 bearing small round granules, interstriae 2 with very minute granules, nearly indistinct (Figures 3.6.27–3.6.29) … pilosa DISCUSSION With 315 known species (Smith, unpublished), Xyleborus is the most speciose xyleborine genus and researchers have long recognized the need for its taxonomic revision (Hubbard 1897, Wood 1986, Jordal et al. 2000, Cognato et al. 2011). Modern techniques using molecular data have provided better understanding of taxon limits within this polyphyletic genus (Hulcr and Cognato 2009, Cognato et al. 2020) and this study provides further revision through the designation of the new genus Xenoxylebora from African and South American Xyleborus species. 172 Xenoxylebora was recovered in all phylogenetic analyses and well-supported by the Bayesian, likelihood, and parsimony analyses (Figures 3.2 and 3.3). It is morphologically distinguished from other xyleborine genera by characters of the protibiae, antennal club, and scutellum. Its phylogenetic position within Xyleborini clearly places the genus outside Xyleborus, senso stricto, and the other Neotropical genera, i.e., Coptoborus, Dryocoetoides and Taurodemus (Figure 3.2). These results demonstrate that Xenoxylebora colonized the Neotropics independent of the main radiation of endemic Neotropical Xyleborini including Coptoborus, Dryocoetoides and Sampsonius (Cognato et al. 2011). There are many examples of bark and ambrosia beetles dispersing among and between islands as well as the closest continent (Jordal and Hewitt 2004, Kirkendall and Jordal 2006, Jordal 2013, Cognato et al. 2018, Jordal 2021b). Dispersal across large oceanic distances is less frequent and bicontinental scolytine lineages spread between Africa and South America (Jordal 2012, 2015, Gohli et al. 2016, Jordal 2017, Eliassen and Jordal 2021) are especially notable because this distribution is uncommon in the subfamily (Figure 3.1) (Jordal 2012, Hulcr et al. 2015). The most likely place of origin for Xenoxylebora is the Neotropics followed by a dispersal to Africa. This is contrary to the more common westward dispersal from Africa to the Americas in which the strong South Equatorial Current can transport terrestrial arthropods inside floating plant material (Fratantoni et al. 2000, Renner 2004a, Cognato 2013, Jordal 2015). The African Xenoxylebora clade renders the Neotropical Xenoxylebora species paraphyletic. This, and the biogeography analyses suggest that the ancestor of the African clade originated in either Africa or South America (Figure 3.2). 173 Interaction between storm winds and the North Equatorial Countercurrent is probably responsible for the eastward migration of several plant taxa, arthropods and scolytine clades from the Americas to Africa (Richardson et al. 1992, Fratantoni et al. 2000, Givnish et al. 2004, Renner 2004a, Won and Renner 2006, Murray and Heraty 2016, Jordal 2017). Xenoxylebora could have dispersed to Africa through this migration pathway. However, future phylogenetic analyses should include Xenoxylebora specimens from additional localities to better test the relationship between Neotropical and African Xenoxylebora. Also, a denser sampling of xyleborines would allow for further testing of the current biogeographic observations. Only then will the evolutionary and biogeographical histories of Xenoxylebora be more clearly understood. Species and generic diversity of African and South American ambrosia beetles is under- described and would benefit from increased sampling and taxonomic study (Wood 2007, Smith et al. 2017, Cognato et al. 2020, Dole et al. 2021, Jordal 2021a, 2021b). Collection of more Xenoxylebora throughout its known geographic range for the extraction of additional molecular data would likely reveal additional species and help to better define proposed species limits within the genus. Xenoxylebora sphenos was observed to vary morphologically between collection locations and the phylogenetic analysis provides only moderate support of the monophyly of the species (Figure 3.2). However, this study had insufficient data to justify revision of the species. The preponderance of xyleborine ambrosia beetles distributed between Africa and South America makes it clear that there has been an historically important exchange of scolytine fauna between the two continents that likely profoundly impacted the biodiversity of both regions (Figure 3.1) (Jordal 2012, Cognato 2013, Gohli et al. 2016, Jordal 2017, 2021a). Continued 174 investigation of these shared faunas is likely to provide a better understanding of the evolution and biogeographical history of Xyleborini and in general Scolytinae. ACKNOWLEDGEMENTS We thank M. Lourdes Chamorro, Charyn Micheli, Floyd Shockley, Terry Erwin † (NMNH), Harald Schillhammer (NHMW), Stéphane Hanot (RMCA), Zack Falin (SEMC), Andrew Short (University of Kansas), Thomas Atkinson (University of Texas), and Bjarte Jordal (University of Bergen) for providing access to specimens, Roger Beaver (Chiangmai, Thailand) for assistance with the African fauna and Floyd Shockley (NMNH) for permission to use type images of Xyleborus subcrenulatus in this study. Funding for fieldwork and laboratory costs was provided, in part, by NSF DEB-0328920 (PEET), DEB1256663, the Committee for Research and Exploration of the National Geographic Society (9975-16) and USDA Forest Service Early Detection Rapid Response program cooperative agreement (11-DG-11420004-257) awarded to AIC; The Coleopterists Society Graduate Student Research Enhancement Award, and Society of Systematic Biologists Award for Graduate Student Research awarded to RKO; graduate research grants from the Michigan State University Department of Entomology awarded to RKO and SMS; and Amazon Conservation Association awarded to SMS. Collection of Terry Erwin’s specimens was funded by Ecuambiente Consulting Group, Ecuador; Casey Fund, Department of Entomology, NMNH; and National Museum of Natural History Lowland Amazon Project. 175 APPENDIX 176 Voucher Name Genus species Cop.neo1 Xenoxylebora addenda Xyleb_sp_111 Xenoxylebora calculosa SMS373 Xenoxylebora collarti Copto neo 102 Xenoxylebora neosphenos SMS368 Xenoxylebora sphenos AF187142.1 Xenoxylebora sphenos Ambobl Ambrosiodmus obliquus Theo sp 89 Ambrosiodmus sp. nov. Ambtac Ambrosiodmus tachygraphus Xylsay Anisandrus sayi Xyomul Cnestus mutilatus Theo.sp1 Coptoborus capillisoror Cop.sp2 Coptoborus chica The.coa1 Coptoborus coartatus Copto pseu 108 Coptoborus exilis Cop.tol1 Coptoborus furiosa SMS371 Coptoborus leeloo Cop.pse1 Coptoborus martinezae Cop.och1 Coptoborus ochromactonus Table 3.1: Specimens used for molecular phylogenetic reconstruction including voucher name, specific identification, collection location, publication source, and GenBank numbers. 177 Table 3.1 (cont’d) Voucher Name Genus species Cop.sp1 Coptoborus papillicauda Cop.sp3 Coptoborus pristis Cop.neo2 Coptoborus pristis Copto sp 105 Coptoborus pseudotenuis Copto bellus 94 Coptoborus sagitticauda SMS369 Coptoborus scully Cop.ves1 Coptoborus vespatorius 1 Copto vesp 98 Coptoborus vespatorius 98 Theo sp 110 Coptoborus villosulus Theo theo 101 Coptoborus villosulus 101 Theo.sp2 Coptoborus villosulus 2 Theo sp 87 Coptoborus villosulus 87 Theo sp 88 Coptoborus villosulus 88 Dryoc gran 109 Dryocoetoides granulicauda Dry.cap1 Dryocoetoides nr. capucinus 1 nr. capucinus Dryoc cap 107 Dryocoetoides 107 Dry.sp1 Dryocoetoides sp. 1 Dryoc sp 97 Dryocoetoides sp. 97 Xylpos Euwallacea posticus 178 Table 3.1 (cont’d) Voucher Name Genus species Xylsim Euwallacea similis Euwsp02_273 Euwallacea wallacei Euwxan_283 Euwallacea semirudis Samdam 357 Sampsonius dampfi Sam ens 128 Sampsonius ensifer Tau god 1 Taurodemus godmani Tau_var_1 Taurodemus varians Xyipex Xyleborinus exiguus Xyleb_grac_106 Xyleborinus gracilis Xyiint Xyleborinus intersetosus Xyiqua Xyleborinus quadrispinosus Xyleb_rec_96 Xyleborinus reconditus Xyisax Xyleborinus saxsesenii Xyisig Xyleborinus signatipennis Xylaff Xyleborus sp. F Xylfer_352 Xyleborus bispinatus Xylgla Xyleborus glabratus Xylperf Xyleborus perforans Xylall Xyleborus principalis Xylvol_356 Xyleborus volvulus Xylxyl_355 Xyleborus xylographus 179 Table 3.1 (cont’d) Voucher Name Coll Location Source Cop.neo1 Suriname: Sipaliwini This study Xyleb_sp_111 Peru: Madre de Dios This study SMS373 Kenya: Kakamega District This study Copto neo 102 Ecuador: Napo Province, Parque Nacional Yasuní This study SMS368 Cameroon: Southwest Region, Limbe This study Normark et AF187142.1 Uganda: Kibale National Park al. 1999 Cognato et Ambobl USA: SC al. 2011 Theo sp 89 Ecuador: Manabí Province This study Cognato et Ambtac USA: MD al. 2011 Cognato et Xylsay USA: MD al. 2011 Cognato et Xyomul USA: MS al. 2011 Theo.sp1 Brazil: Bahia, Serra Bonita Reserve This study Cop.sp2 Suriname: Sipaliwini This study The.coa1 Ecuador: Los Ríos Province, Samama Nature Reserve This study Copto pseu 108 Panama: Panamá, Parque Nacional Soberanía This study Cop.tol1 Ecuador: Los Ríos Province, Samama Nature Reserve This study SMS371 Ecuador: Napo Province, Parque Nacional Yasuní This study Cop.pse1 Ecuador: Los Ríos Province, Samama Nature Reserve This study Cop.och1 Ecuador: Cotopaxi Province, Yacusinchi Reserve This study 180 Table 3.1 (cont’d) Voucher Name Coll Location Source Cop.sp1 Suriname: Sipaliwini This study Cop.sp3 Brazil: Bahia, Serra Bonita Reserve This study Cop.neo2 Ecuador: Cotopaxi Province, Yacusinchi Reserve This study Copto sp 105 Ecuador: Napo Province, Parque Nacional Yasuní This study Copto bellus 94 Guyana: Region 8, Iwokrama Forest This study SMS369 Ecuador: Napo Province, Parque Nacional Yasuní This study Cop.ves1 Ecuador: Los Ríos Province, Samama Nature Reserve This study Copto vesp 98 Guyana: Region 8, Iwokrama Forest This study Theo sp 110 Peru: Madre de Dios Dept., Los Amigos Biological Station This study Theo theo 101 Guyana: Region 8, Iwokrama Forest This study Theo.sp2 Brazil: Bahia, Serra Bonita Reserve This study Theo sp 87 Ecuador: Manabí This study Theo sp 88 Ecuador: Napo Province, Parque Nacional Yasuní This study Dryoc gran 109 Peru: Madre de Dios Dept., Los Amigos Biological Station This study Dry.cap1 Brazil: Bahia, Serra Bonita Reserve This study Dryoc cap 107 Panama: Panamá, Cerro Azul This study Dry.sp1 Ecuador: Cotopaxi Province, Yacusinchi Reserve This study Dryoc sp 97 Guyana: Region 8, Iwokrama Forest This study Cognato et Xylpos Costa Rica al. 2011 181 Table 3.1 (cont’d) Voucher Name Coll Location Source Cognato et Xylsim Papua New Guinea al. 2011 Cognato et Euwsp02_273 Papua New Guinea al. 2011 Cognato et Euwxan_283 Papua New Guinea al. 2011 Cognato et Samdam 357 Ecuador al. 2011 Sam ens 128 Ecuador: Napo Province, Parque Nacional Yasuní, Tiputini This study Tau god 1 Panama: Chiriquí Province This study Tau_var_1 Peru: Madre de Dios This study Cognato et Xyipex Papua New Guinea al. 2011 Xyleb_grac_106 Panama: PAN 32 This study Cognato et Xyiint Costa Rica al. 2011 Cognato et Xyiqua Madagascar al. 2011 Xyleb_rec_96 Guyana: Region 8, Iwokrama Forest This study Cognato et Xyisax USA: OH al. 2011 Cognato et Xyisig Madagascar al. 2011 Cognato et Xylaff Costa Rica al. 2011 Cognato et Xylfer_352 Papua New Guinea al. 2011 Cognato et Xylgla USA: GA al. 2011 Cognato et Xylperf Papua New Guinea al. 2011 Cognato et Xylall Ghana al. 2011 Cognato et Xylvol_356 Papua New Guinea al. 2011 Cognato et Xylxyl_355 USA:MI al. 2011 182 Table 3.1 (cont’d) Voucher Name GenBank Numbers COI CAD EF1-𝛼 28s Cop.neo1 TBD TBD TBD TBD Xyleb_sp_111 TBD TBD TBD TBD SMS373 TBD TBD TBD TBD Copto neo 102 TBD TBD TBD TBD SMS368 TBD TBD TBD TBD AF187142.1 AF187142 N/A AH011661 N/A Ambobl HM064048 HM064227 HM064154 HM099667 Theo sp 89 TBD TBD TBD TBD Ambtac HM064053 HM064232 HM064159 HM099671 Xylsay GU808704 GU808626 GU808741 GU808589 Xyomul GU808719 GU808641 GU808755 GU808603 Theo.sp1 TBD TBD TBD TBD Cop.sp2 TBD TBD TBD TBD The.coa1 TBD TBD TBD TBD Copto pseu 108 TBD TBD TBD TBD Cop.tol1 TBD TBD TBD TBD SMS371 TBD TBD TBD TBD Cop.pse1 TBD TBD TBD TBD Cop.och1 TBD TBD TBD TBD 183 Table 3.1 (cont’d) Voucher Name GenBank Numbers COI CAD EF1-𝛼 28s Cop.sp1 TBD TBD TBD TBD Cop.sp3 TBD TBD TBD TBD Cop.neo2 TBD TBD TBD TBD Copto sp 105 TBD TBD TBD TBD Copto bellus 94 TBD TBD TBD TBD SMS369 TBD TBD TBD TBD Cop.ves1 TBD TBD TBD TBD Copto vesp 98 TBD TBD TBD TBD Theo sp 110 TBD TBD TBD TBD Theo theo 101 TBD TBD TBD TBD Theo.sp2 TBD TBD TBD TBD Theo sp 87 TBD TBD TBD TBD Theo sp 88 TBD TBD TBD TBD Dryoc gran 109 TBD TBD TBD TBD Dry.cap1 TBD TBD TBD TBD Dryoc cap 107 TBD TBD TBD TBD Dry.sp1 TBD TBD TBD TBD Dryoc sp 97 TBD TBD TBD TBD Xylpos HM064133 HM064312 N⁄A HM099748 184 Table 3.1 (cont’d) Voucher Name GenBank Numbers COI CAD EF1-𝛼 28s Xylsim HM064139 HM064317 HM064217 HM099754 Euwsp02_273 N⁄A HM064265 HM064183 HM099704 Euwxan_283 HM064086 HM064267 HM064185 HM099706 Samdam 357 HM064095 HM064276 HM064190 HM099713 Sam ens 128 TBD TBD TBD TBD Tau god 1 TBD TBD TBD TBD Tau_var_1 TBD TBD TBD TBD Xyipex HM064109 N ⁄A N⁄ A HM099725 Xyleb_grac_106 TBD TBD TBD TBD Xyiint HM064108 N⁄A HM064200 N⁄A Xyiqua HM064111 HM064289 HM064201 HM099726 Xyleb_rec_96 TBD TBD TBD TBD Xyisax HM064112 HM064290 HM064202 HM099727 Xyisig HM064113 HM064291 N⁄ A HM099728 Xylaff GU808696 GU808621 GU808735 GU808581 Xylfer_352 HM064126 HM064305 HM064211 HM099741 Xylgla HM064127 HM064306 N/A HM099742 Xylperf HM064132 HM064311 N ⁄ A HM099747 Xylall HM064118 HM064296 N ⁄ A HM099733 Xylvol_356 HM064149 HM064327 HM064222 HM099763 Xylxyl_355 HM064150 HM064328 N ⁄ A HM099764 185 Gene Primer Primer sequence (5'-3') 1495b AACAAATCATAAAGATATTGGRAC COI rev750 GAAATTATNCCAATTCCTGG ApCADfor4 TGGAARGARGTBGARTACGARGTGGTYCG CAD ApCADrev1mod GCCATYRCYTCBCCYACRCTYTTCAT eflafor1 TACGTAACCATCATTGATGCTYCC EF1-𝛼 eflarev1 CCTTCTTTACGTTCAATGGACCATCC D2F1 ACTGTTGGCGACGATGTTCT D3R2 TCTTCGCCCCTATACCC 28s 3665 AGACAGAGTTCAAGAGTACGTG 4048 TTGCTCCGTGTTTCAAGACGGG Table 3.2: Primers used for PCR reactions. 186 Partition (Gene-Codon Phylogeny Position) Model COI-1st F81+I+G COI-2nd GTR+I+G COI-3rd GTR+I+G PartitionFinder CAD-1st K80+G CAD-2nd HKY+I CAD-3rd K80+I+G Table 3.3: Models of nucleotide substitution assigned by PartitionFinder 2.1.1 and ModelFinder, respectively, and used for phylogeny reconstruction by likelihood and Bayesian analyses. JC = Jukes Cantor model of evolution with equal base frequencies and equal substitution rates (Jukes and Cantor 1969); F81 = Felsenstein model with unequal base frequencies and equal rates of substitution (Felsenstein 1981); K80 = Kimura model with equal base frequencies and unequal rates of transition and transversion substitutions (Kimura 1980); HKY = Hasegawa model with unequal base frequencies and unequal rates of transition and transversion substitutions (Hasegawa et al. 1985); TN = Tamura Nei model with unequal rates of purine and pyrimidine rates and unequal rates of transition and transversion substitutions (Tamura and Nei 1993); TIM2e =Transition model with equal base frequencies and AC and CG substitution rates equal to AT and GT, respectively; TIM3e = Transition model with equal base frequencies and AC and AT substitution rates equal to CG and GT, respectively; GTR = general time reversible model with unequal base frequencies and unequal rates of substitution (Lanave et al. 1984, Rodríguez et al. 1990); I = variable nucleotide frequencies; F = nucleotide frequencies determined from the data; FQ = nucleotides with equal frequencies; G = gamma distributed rates of variation; R = FreeRate model of distributed rates of variation (Yang 1995, Soubrier et al. 2012). 187 Table 3.3 (cont’d) Partition (Gene-Codon Phylogeny Position) Model EF1-𝛼-1st JC+I EF1-𝛼-2nd HKY+I+G PartitionFinder EF1-𝛼-3rd GTR+I+G 28S GTR+G COI-1st, CAD-2nd, EF1-𝛼-1st HKY+F+I+G COI-2nd TN+F+R3 COI3rd TIM3e +FQ+G ModelFinder CAD-1st, EF1-𝛼-3rd TIM3e+FQ_I+G CAD -3rd, EF1-𝛼-2nd TIM2e+FQ+I+G 28s GTR+F+I+G 188 Taxon Label Genus species Coptoborus_n_sp_Surinam Xenoxylebora addenda Xyleb_sp_111 Xenoxylebora calculosa Xyleborus_collarti_Kenya Xenoxylebora collarti Coptoborus_neospenos_Tiputini Xenoxylebora neosphenos Xyleborus_sphenos Xenoxylebora sphenos Xyleborus_sphenos_Camaroon Xenoxylebora sphenos Ambrosiodmus_obliquus Ambrosiodmus obiquus Ambrosiodmus_sp_Ecuador_Lalo_Loor Ambrosiodmus sp. nov. Ambrosiodmus_tachygraphus Ambrosiodmus tachygraphus Anisandrus_sayi Anisandrus sayi Cnestus_mutilatus Cnestus mutilatus Theoborus_capillisoror_Brazil_Bahia Coptoborus capillisoror Coptoborus_chica_Surinam Coptoborus chica Theoborus_coartatus_Ecuador_Samama Coptoborus coartatus Coptoborus_exilis_Panama Coptoborus exilis Coptoborus_furiosa_Ecuador_Samama Coptoborus furiosa Coptoborus_leeloo Coptoborus leeloo Coptoborus_martinezae_Ecuador_Samama Coptoborus martinezae Coptoborus_ochromactonus_Ecuador_Yacusinchi Coptoborus ochromactonus Coptoborus_papillicauda_Surinam Coptoborus papillicauda Table 3.4: Biogeographical states used for biogeography analyses. 189 Table 3.4 (cont’d) Taxon Label Genus species Coptoborus_pristis_Ecuador_Yacusinchi Coptoborus pristis Coptoborus_pristis_Brazil_Bahia Coptoborus pristis Coptoborus_pseudotenuis_Ecuador Coptoborus pseudotenuis Coptoborus_sagitticauda_Guyana Coptoborus sagitticauda Coptoborus_scully Coptoborus scully Coptoborus_vespatorius_Ecuador_Samama Coptoborus vespatorius 1 Coptoborus_vespatorius_Guyana Coptoborus vespatorius 98 Theoborus_villosulus_Peru Coptoborus villosulus Theoborus_villosulus_Guyana Coptoborus villosulus 101 Theoborus_villosulus_Brazil_Bahia Coptoborus villosulus 2 Theoborus_villosulus_Ecuador Coptoborus villosulus 87 Theoborus_villosulus_Tiputini Coptoborus villosulus 88 Dryocoetoides_grandulicauda_Peru Dryocoetoides granulicauda Dryocoetoides_nr_capicinus_2_Brazil_Bahia Dryocoetoides nr. capusinus 1 Dryocoetoides_nr_capicinus_1_Panama Dryocoetoides nr. capusinus 97 Dryocoetoides_sp_1Ecuador_Yacusinchi Dryocoetoides sp. 1 Dryocoetoides_sp_2_Guyana Dryocoetoides sp. 97 Xylpos Euwallacea posticus Xylsim Euwallacea similis Euwsp02_273 Euwallacea wallacei 190 Table 3.4 (cont’d) Taxon Label Genus species Euwxan_283 Euwallacea semirudis Samdam_357 Sampsonius dampfi Sam_ens_128 Sampsonius ensifer Taurodemus_godmani Taurodemus godmani Tau_var_1 Taurodemus varians Xyipex Xyleborinus exiguus Xyleb_grac_106 Xyleborinus gracilis Xyiint Xyleborinus intersetosus Xyiqua Xyleborinus quadrispinosus Xyleb_rec_96 Xyleborinus reconditus Xyisax Xyleborinus saxesenii Xyisig Xyleborinus signatipennis Xylfer_352 Xyleborus bispinatus Xyleborus_glabratus Xyleborus glabratus Xylperf Xyleborus perforans Xylall Xyleborus principalis Xyleborus_affinis Xyleborus sp. F Xylvol_356 Xyleborus vovulus Xylxyl_355 Xyleborus xylographus 191 Table 3.4 (cont’d) Taxon Label Region of Origin Coptoborus_n_sp_Surinam Neotropical Xyleb_sp_111 Neotropical Xyleborus_collarti_Kenya Afrotropical Coptoborus_neospenos_Tiputini Neotropical Xyleborus_sphenos Afrotropical Xyleborus_sphenos_Camaroon Afrotropical Ambrosiodmus_obliquus Nearctic Ambrosiodmus_sp_Ecuador_Lalo_Loor Neotropical Ambrosiodmus_tachygraphus Nearctic Anisandrus_sayi Australasian & Oceanian Cnestus_mutilatus Australasian & Oceanian Theoborus_capillisoror_Brazil_Bahia Neotropical Coptoborus_chica_Surinam Neotropical Theoborus_coartatus_Ecuador_Samama Neotropical Coptoborus_exilis_Panama Neotropical Coptoborus_furiosa_Ecuador_Samama Neotropical Coptoborus_leeloo Neotropical Coptoborus_martinezae_Ecuador_Samama Neotropical Coptoborus_ochromactonus_Ecuador_Yacusinchi Neotropical Coptoborus_papillicauda_Surinam Neotropical 192 Table 3.4 (cont’d) Taxon Label Region of Origin Coptoborus_pristis_Ecuador_Yacusinchi Neotropical Coptoborus_pristis_Brazil_Bahia Neotropical Coptoborus_pseudotenuis_Ecuador Neotropical Coptoborus_sagitticauda_Guyana Neotropical Coptoborus_scully Neotropical Coptoborus_vespatorius_Ecuador_Samama Neotropical Coptoborus_vespatorius_Guyana Neotropical Theoborus_villosulus_Peru Neotropical Theoborus_villosulus_Guyana Neotropical Theoborus_villosulus_Brazil_Bahia Neotropical Theoborus_villosulus_Ecuador Neotropical Theoborus_villosulus_Tiputini Neotropical Dryocoetoides_grandulicauda_Peru Neotropical Dryocoetoides_nr_capicinus_2_Brazil_Bahia Neotropical Dryocoetoides_nr_capicinus_1_Panama Neotropical Dryocoetoides_sp_1Ecuador_Yacusinchi Neotropical Dryocoetoides_sp_2_Guyana Neotropical Xylpos Nearctic Xylsim Australasian & Oceanian Euwsp02_273 Australasian & Oceanian 193 Table 3.4 (cont’d) Taxon Label Region of Origin Euwxan_283 Australasian & Oceanian Samdam_357 Neotropical Sam_ens_128 Neotropical Taurodemus_godmani Neotropical Tau_var_1 Neotropical Xyipex Australasian & Oceanian Xyleb_grac_106 Neotropical Xyiint Neotropical Xyiqua Afrotropical Xyleb_rec_96 Neotropical Xyisax Worldwide Xyisig Afrotropical Xylfer_352 Worldwide Xyleborus_glabratus Neotropical Xylperf Worldwide Xylall Afrotropical Xyleborus_affinis Worldwide Xylvol_356 Worldwide Xylxyl_355 Nearctic 194 Figure 3.1: Map of Scolytinae with native distributions across Africa and South America (Wood and Bright 1992, Jordal 2012, Hulcr et al. 2015, Gohli et al. 2016, Bright 2010, 2019, Atkinson 2021, Eliassen and Jordal 2021, Jordal 2021b, 2021c, 2021d, 2021e, 2021f). Each circled number represents the number of species from the corresponding group endemic or established in the indicated continent. The expanded side of each line indicates the continent containing a plurality of species. Two Premnobius species have cosmopolitan distributions and are therefore each counted once on each continent: P. cavipennis and P. ambitiosus. Xenoxylebora is depicted in blue; black indicates xyleborine genera, and grey indicated non-xyleborine groups. 195 Figure 3.2: Phylogenetic tree resulting from a Bayesian analysis of CO1, CAD, EF1-α and 28S. Nodes are labeled with posterior probability/bootstrap support. Posterior probabilities > 0.95 are considered strong clade support. Nodes within Xenoxylebora are additionally labeled with pie diagrams indicating the relative probabilities of origin in the Neotropics, Afrotropics, and Neotropics/Afrotropics. An outgroup Anisandrus sayi was used to root the tree. 196 Figure 3.3: Maximum likelihood tree resulting from Nearest Neighbor Interchange search of CO1, CAD, EF1-α and 28S sequences using IQ-TREE version 2.1.3. Nodes are labeled with bootstrap support from 1000 pseudoreplications. Anisandrus sayi was used to root the tree. 197 Figure 3.4: Diagnostic characters for Xenoxylebora. (3.4.3) type 1 antennal club. (3.4.4) slender protibia, outer edge weakly rounded, posterior face flat and unarmed. (3.4.5) sutural tubercles on the declivital apex. 198 Figure 3.5: Dorsal, lateral, and declivital aspects of Xenoxylebora addenda holotype (3.5.6– 3.5.8); Xenoxylebora calculosa holotype (3.5.9–3.5.11); Xenoxylebora caudata paratype (3.5.12– 3.5.14); Xenoxylebora collarti (3.5.15–3.5.17); Xenoxylebora hystricosa holotype (3.5.18– 3.5.20). 199 Figure 3.6: Dorsal, lateral, and declivital aspects of Xenoxylebora neosphenos (3.6.21–3.6.23); Xenoxylebora perdiligens (3.6.24–3.6.26); Xenoxylebora pilosa holotype (3.6.27–3.6.29); Xenoxylebora serrata holotype (3.6.30–3.6.32); Xenoxylebora sphenos (3.6.33–3.6.35). 200 Figure 3.7: Dorsal, lateral, and declivital aspects of Xenoxylebora subcrenulata holotype (3.7.36–3.7.38); Xenoxylebora sulcata holotype (3.7.39–3.7.41); Xenoxylebora syzygii paratype (3.7.42–3.7.44). 201 LITERATURE CITED 202 LITERATURE CITED Atkinson TH. 2021. Bark and ambrosia beetles of the Americas. http://www.barkbeetles.info. (October 22, 2021). 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