TRUFFLES IN MICHIGAN: IMPACTS OF HERBICIDES ON THEIR GROWTH, EFFECTIVENESS OF IN-FIELD INOCULATIONS, AND THE DISCOVERY OF A LOCAL TRUFFLE (TUBER RUGOSUM) By Bryan Rennick A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Plant Pathology – Master of Science 2023 ABSTRACT With the advent of promising truffle cultivation techniques, there are many new truffle plantations (truffière) being established and managed throughout the world; one of the biggest challenges these plantations face is weed management. There is little known about the impact popular herbicides have on truffle mycelial growth. Here, I discuss how the use of herbicides in management may impact mycelial growth. Pure culture growth assays were performed to assess the impacts glyphosate and glufosinate chemistries on three species of truffle, two species of morels, and two members of the Mortierellaceae. While there were variable responses by each species tested, most fungi experienced growth inhibition near or above the manufacturer's recommended application rates for general weed use. I also assess how introducing exotic truffle species, local ectomycorrhizal species, and commercial mycorrhizal inoculants in post-methyl bromide fumigated bare-root conifer tree nursery impacts seedling growth. To test whether inoculum benefits seedling health and growth, exotic, and commercial ectomycorrhizal inoculua were added at the time of planting for three species of conifer. The results showed a high mycorrhizal diversity on bare-root seedling roots, which generally differed from those of the added inoculum. Additionally, there were no growth or health improvements over uninoculated control blocks compared to treatment blocks. Finally, I will showcase the taxonomic discovery of a new species of truffle native to Michigan, Tuber rugosum. Also, observing slug fungivory of this new truffle led to an improved ascus technique for performing scanning electron microscopy of truffle spores. Collectively, this thesis aims to inform herbicide use in myco-agricultural lands, to inform the use of added mycorrhizal inoculants in bare-root conifer tree nurseries, and to improve our knowledge on Michigan’s native truffle species. ACKNOWLEDGEMENTS I would like to express my sincere gratitude to my mentor Dr. Gregory Bonito who has been an unwavering source of support, patience, encouragement, and friendship throughout my academic journey. In addition to Dr. Bonito, I would also like to acknowledge Dr. Bert Cregg and Dr. Monique Sakalidis whose encouragement and support through this journey was a source of motivation and reassurance. I also want to acknowledge the critical role Dr. Nico Benucci played throughout my time in the Bonito Lab. Dr. Benucci’s guidance and mentorship was crucial in building the mycological skills I now master. Collectively, your wisdom and mentorship were instrumental in my professional and scientific growth, and I am fortunate to have such individuals guide me for these past few years. Additionally, I am grateful to the department of Plant, Soil and Microbial Science at Michigan State University for providing me with an exceptional experience. The faculty and staff have been instrumental in molding me into the young mycologist I have become. Our faculty members’ passion and insight has been a constant source of inspiration and motivate me to pursue my research with equal fervor. Further, to the countless other lab mates, mentors, and mentees, your unending help, wisdom, and companionship are the foundation for my continued development and inspiration. Your insights have undoubtably pushed my professional development to higher plateaus than I initially imagined possible. I am also deeply grateful to the Veterans Administration system, the medical staff at Bronson Hospital, and the medical staff at Sparrow Hospital for their unyielding support, compassion and dedication to my health and wellbeing through my leukemia diagnosis and treatment. To the VA, thank you for your endless support in providing me with the care that I needed when I needed it most; your dedication to the education, health, and well-being of iii veterans like me is truly commendable, and I am grateful for the services that you have provided. To the medical staff at Bronson Hospital, I cannot thank you enough for your quick, compassionate diagnosis and treatment. To the medical staff at Sparrow Hospital, thank you for keeping me alive, motivated, and for always making me feel welcome and important. Finally, I am profoundly grateful to my friends and family for their steadfast support, encouragement, and faith in my abilities throughout my academic journey. Their love and support have been instrumental in helping me to achieve my goals and pursue my passions. To my spouse Kristy and my children Joshua and Nathan, I cannot thank you enough for your endless love and support; your boundless faith in my abilities and your encouragement to pursue my dreams have been the driving force behind my academic success. Your sacrifices, both big and small, have enabled me to pursue my education and I am forever grateful for everything that you have done for me. To my parents, siblings, and extended family, thank you for your support and encouragement throughout my academic journey. To all my friends and family, I am endlessly grateful for everything that you have done for me in ways words can never express. Thank you. iv TABLE OF CONTENTS LIST OF ABBREVIATIONS ..................................................................................................... vi Chapter 1 Introduction................................................................................................................. 1 Problem statement ...................................................................................................................... 2 Background ................................................................................................................................ 3 Research focus ......................................................................................................................... 15 Value of this research .............................................................................................................. 16 Chapter 2 Impacts of Glufonisate-Ammonium and Glyphosate Herbicides on Mycelial Growth of Truffles, Morels, and Plant Growth Promoting Fungi ......................................... 18 Abstract ..................................................................................................................................... 19 Introduction.............................................................................................................................. 20 Methods .................................................................................................................................... 22 Results....................................................................................................................................... 29 Discussion................................................................................................................................. 41 Chapter 3 Does ectomycorrhizal inoculum after methyl bromide field applications in bare- root conifer nursery seedlings establish and persist? .............................................................. 45 Abstract ..................................................................................................................................... 46 Introduction.............................................................................................................................. 46 Methods .................................................................................................................................... 50 Results....................................................................................................................................... 57 Discussion................................................................................................................................. 73 Summary................................................................................................................................... 76 Conclusions .............................................................................................................................. 76 Chapter 4 Tuber rugosum, a new species from Northeastern North America: Slug mycophagy aides in electron microscopy of ascospores .......................................................... 78 Abstract ..................................................................................................................................... 79 Introduction.............................................................................................................................. 79 Materials and Methods ............................................................................................................ 82 Results....................................................................................................................................... 89 Discussion............................................................................................................................... 105 Conclusion.............................................................................................................................. 108 A key to Tuber spp. in the Rufum clade ................................................................................ 108 Chapter 5 Synthesis .................................................................................................................. 111 Synthesis ................................................................................................................................. 112 Conclusion.............................................................................................................................. 115 REFERENCES.......................................................................................................................... 117 APPENDIX CURRENT CURRICULUM VITAE ................................................................ 133 v LIST OF ABBREVIATIONS ANOVA Analysis of variance BI Bayesian inference BLAST Basic Local Alignment Search Tool CIPRES Cyberinfrastructure for Phylogenetic Research DNA Deoxyribonucleic Acid EF1α elongation factor 1α EPSPS 5-enolpyruvyl shikimic acid-3-phosphate synthase ES Extraction Solution FAME Fatty acid methyl esters Fig Figure H3PO4 Phosphoric acid HCl Hydrogen chloride INVGAMMA Inverse gamma distribution ITS Internal Transcribed Spacer KCl Potassium chloride KOH Potassium hydroxide MCMC Markov chain Monte Carlo MCMCMC Metropolis-coupled Markov Chain Monte Carlo MEB Malt Extract Broth ML Maximum likelihood MSU Michigan State University NCBI National Center for Biotechnology Information vi OTU Operational Taxonomic Unit PBS Phosphate-buffered saline PDA Potato dextrose agar pH Potential of Hydrogen p-value Probability value RAxML Randomized Axelerated Maximum Likelihood rDNA Ribosomal Deoxyribonucleic Acid RNA Ribonucleic acid RPB2 RNA polymerase II RPS Rotations per second RTSF Research Technology Support Facility SEM Scanning electron microscopy sp. Species Sp. Nov. Species nova (new species) TM Tuber Media VA Veterans Administration YE Yeast Extract vii Chapter 1 Introduction 1 Problem statement As with any cropping system, conifer trees and nut trees face many challenges. These tree producers share many risks, from failed crops to increasing disease pressure with few pesticide options available (Frampton et al., 2018; Pettersson et al., 2017). Conifer trees, for example, take seven to 15 years to grow before harvest and represent a substantial time investment before any returns are possible (Zinati et al., 2016). When a conifer fails to sell, or a nut tree is inadequately pollinated for example, there are few if any avenues by which the grower may earn income from that tree. Michigan produces over four million conifers each year and annually, over 25 million conifers are purchased in the United States grown on roughly 350,000 acres (National Christmas Tree Association, 2019). Hazelnut production has also been increasing, and as of 2021 there are over 60,000 nut bearing acres in the United States (USDA, 2022). These trees represent important jobs and revenue; maturing into a diverse agroforestry system could strengthen this delicate sector. One potential revenue source for plantations such as conifer tree farms, may be found growing on the roots as edible mushroom or truffle forming ectomycorrhizal fungi. By pairing ectomycorrhizal fungi that form edible mushrooms or truffles with compatible tree species, some of the risk associated with growing conifers or nut trees in Michigan may be mitigated through increased revenue and decreased pathogen presence within their stands. However, there are several challenges such as weed plants and non-target fungi which need to be addressed for growers to have a chance at success. Researching alternative revenue sources and their challenges is critical for developing a more robust agroforestry plantation. 2 Background Co-cropping with Ectomycorrhizal Fungi Co-cropping, a practice where two or more crops are grown together, has garnered much interest as this technique offers to increase yields by adding an additional source of revenue, trapping unwanted pests, or even by improvements to the soil. Unlike plant-based co-cropping systems, fungi do not compete for sunlight. One group of fungi in particular form beneficial symbiotic relationships with particular host trees. These fungi, collectively called ectomycorrhizal fungi, interact with the roots of their host tree. This root-fungal interface serves as the point of nutrient exchange between the two organisms. Economically important trees within the Betulaceae, Fagaceae, Juglandaceae, and Pinaceae families form mutually beneficial relationships with many genera of ectomycorrhizal fungi (Tedersoo et al., 2009). Evidence of these symbiotic relationships have been long established between some plant and fungal species dating back millions of years. Fossil records from 50 million years ago (Eocene Princeton chert) provide evidence of well-developed ectomycorrhiza on the roots of Pinus which show the Hartig net, a pseudoparenchymous mantle, and extramatrical hyphae (Lepage et al., 1997). Ectomycorrhizal fungi are characterized by the formation of these unique morphological structures and the interactions this group of fungi has with compatible plants. A Hartig net is the site of nutrient exchange between fungi and host; it consists of fungal hyphae growing outside of, and between root cortical cells of the host plant. The Hartig net can be seen by placing thin cross sections of colonized root tips on a microscope slide for close inspection. An ectomycorrhizal mantle is a sheath composed of pseudoparenchyma hyphae surrounding host plant root tips. The densely packed puzzle-like pseudoparenchymal hyphae which form the mantle may act as a physical 3 barrier and be one of the means by which ectomycorrhizal fungi impart protection against invading pathogens. Variations in the formation and structure of the mantle such as the shape and density of the hyphae can be a tool by which the absence or presence of target ectomycorrhizal fungi can be ascertained. Extending beyond the mantle out into the surrounding soil environment are determinant hyphae called cystidia. The morphology of cystidia can also aid in the identification of the fungi present, for example, Tuber borchii Vittad. short, awl-shaped hyaline cystidia, but Tuber aestivum Vittad. produces curled, interwoven hyaline cystidia (Giomaro et al., 2000; Molinier et al., 2016). These host interactions cannot occur with any host however, compatibility between ectomycorrhizal fungi and host plant can be highly specific. Ectomycorrhizal Host Specificity Though many ectomycorrhizal fungi are generalists in that they can form these symbiotic relationships with multiple different host species; it is wise for any targeted ectomycorrhizal tree producer to ensure the targeted fungi are compatible (G. M. Bonito et al., 2010). Suillus for example, are well known for showing high levels of host specificity amongst Pinus, Pseudotuga, and Larix (Pérez-Pazos et al., 2021). The edible North American truffle T. lyonii Butters and recently described T. brennemanii A. Grupe, Healy & M.E. Sm. and T. floridanum A. Grupe, Sulzbacher & M.E. Sm. dominate the roots of pecan (Carya illinoinensis (Wangenh.) K.Koch) trees (G. Bonito et al., 2011; Grupe et al., 2018) but can be found under oak trees as well. Many tree species depend on these symbionts to survive. Some of these symbioses have been shown to promote growth and health benefits. Picea A. Dietr. trees, which form symbiotic relationships with the ectomycorrhizal fungus Laccaria bicolor (Maire) P.D. Orton, are known to obtain significant increases in biomass after ectomycorrhizal colonization compared to those grown in the same soils without 4 fungal colonization (Quoreshi & Timmer, 2000). The most commonly grown tree in Northern nurseries, Picea abies (L.) H.Karst., has been shown to be resistant to one of the most damaging root rot causing pathogens, Heterobasidion annosum P.K. Buchanan, when its roots are colonized by various ectomycorrhizal forming fungi (Velmala et al., 2017). It is also clear that the presence of ectomycorrhizal fungi on the roots of trees has a positive influence on the growth and health of trees even when soil conditions are less than favorable. Pinus L. trees inoculated with the ectomycorrhizal fungi Pisolithus Alb. & Schwein., Cenococcum Moug. & Fr., and Laccaria Berk. & Broome have been shown to grow significantly better in poor soils which contain high concentrations of heavy metals (Zong et al., 2015). Additionally, chestnut trees have been shown to greatly benefit from ectomycorrhizal colonization when growing on retired and contaminated coal mined landscapes (Bauman et al., 2018). Where there are contaminated soils, the pairing of edible ectomycorrhizal fungi with host trees should be avoided as it has been noted that the fruiting body of certain ectomycorrhizal fungi may act as a toxin sink for the fungi. (Cocchi et al., 2006) found that in many mushroom species such as the ectomycorrhizal fungi Boletus edulis Bull., there were arsenic and cadmium levels equal to or exceeding the maximum weekly dosage recommended by the World Health Organization (Cocchi et al., 2006). Economically and Culinarily Valuable Ectomycorrhizal Fungi Beyond some of the direct benefits to the tree crop through growth promotion, pathogen resistance, or resilience against heavy metal toxicity, there are many ectomycorrhizal fungi which form edible mushrooms or truffles which often fetch very high market prices. Matsutake (Tricholoma matsutake (S. Ito & S. Imai) Singer), porcini (Boletus edulis Bull.), chanterelle (Cantharellus sp. Adans. ex Fr.), saffron milk cap (Lactarius deliciosus (L.) Gray), and truffles 5 (Tuber sp.) are among those ectomycorrhizal fungi possessing high economic and culinary value. Though there have been recent advances, many of the highly prized edible mushroom producing ectomycorrhizal fungi such as Tricholoma matsutake (Yamanaka et al., 2020), Lactarius deliciosus (Guerin-Laguette et al., 2014; Wang et al., 2021), and Cantharellus anzutake W. Ogawa, N. Endo, M. Fukuda & A. Yamada (Ogawa et al., 2019) have little commercial success in cultivation and their entire presence in the market is based on wild foraging efforts (Yun & Hall, 2004). Other edible ectomycorrhizal fungi of interest such as Lactarius deliciosus (L.) Gray, Suillus luteus (L.) Roussel and Suillus variegatus (Sw.) Richon & Roze have been successfully cultivated (González-Ochoa et al., 2003). The apparent inability to cultivate economically important fungi highlights how little is understood about these organisms and how much there is remaining to be discovered in regard to their cryptic lifecycles. The most economically valuable ectomycorrhizal fungi are truffles (Ascomycota, Pezizales, Tuberaceae) in the genus Tuber. What are Truffles? Tuber species, commonly called truffles, are ectomycorrhizal forming fungi which are best known for their aroma with many species fetching high prices in international markets. Though some European truffles command high prices and garner much media attention, there is a rich diversity of truffle species in North America with many species yet to be described (Bonito et al., 2010; Healy et al., 2016). There are a few famous North American truffles, Tuber canaliculatum Gilkey, Tuber gibbosum Harkn., Tuber lyonii, and Tuber oregonense Trappe, Bonito & P. Rawl. contributing greatly toward building the culinary reputation of North American truffles. Many of these truffle species are known as a prized delicacy due to the wide 6 range of complex aromatic compounds they release and can be differentiated based on these compounds alone (Culleré et al., 2013). Unlike many sought after fungal fruiting bodies such as morels (Morchella sp. Dill. ex Pers.), truffles are a sequestrate hypogeous fungi which would ideally be located by a trained animal with a good sense of smell to ensure that the maturity of the mushroom is such that the aromatic qualities the market demands are present (Grupe et al., 2018; Pieroni, 2016). Immature truffles will not produce the desired aroma and are not considered to have much culinary value. Using a meta-analysis of global ITS rDNA, it has been predicted that there are over 180 Tuber species (Bonito et al., 2010). Despite the abundance of truffle species, most are not economically important and only a dozen or so species are harvested in economic quantities. Some of the most sought-after truffle species including Tuber aestivum, T. melanosporum Vittad., and T. borchii, have long culinary histories in Europe where there have been documented efforts to cultivate truffles dating back to the 18th century (Reyna & Garcia- Barreda, 2014; Yun & Hall, 2004). Tuber melanosporum alone accounts for tens of millions of dollars of direct revenue for producers within Europe every year (Reyna & Garcia-Barreda, 2014). The cultivation of these truffles is no simple task as the life-cycle of these fungi are complex and partially unknown. As an obligate symbiont, germination of truffles spores occurs after the presence of root exudates are intercepted (Ali & Jackson, 1988; Yun & Hall, 2004). The germinated spores will then grow and a mycorrhizal connection with the host tree will take place. Truffles are heterothallic organisms and compatible mating types must interact before completing their reproductive cycle (Rubini et al., 2011), and as such, most truffle tree production is spore-based. 7 Mycorrhizal Tree Production One of the most widely used methods for inoculating trees in a nursery with ectomycorrhizal species is through the use of spores (Reyna & Garcia-Barreda, 2014), though other ectomycorrhizal species seem to perform better with mycelial inoculation. Suillus mycorrhizas can be synthesized by spore-based inoculation, but some ectomycorrhizal fungi such as Lactarius Pers. show greater success colonizing nursery trees from vegetative, mycelium-based inoculation techniques (González-Ochoa et al., 2003). Lactarius deliciosus, for example, has been successfully cultivated by layering mycelium into the pots of nursery trees in Pinus species (Guerin-Laguette et al., 2014). Still, spore-based inoculation is the current standard with Tuber species. The precise methodology of spore inoculation will vary based on the target fungal species. Truffle trees start their life as seedlings potted with Tuber spores in sterile or pasteurized soil in a greenhouse until they are mature enough to plant in the field (Iotti et al., 2012). Growing truffle trees by the pot can be expensive due to environmental, labor, and material costs. Roughly 500,000 truffle trees are produced every year in nurseries (Murat, 2014). Ensuring the spores being applied come from the correct target fungus is of critical importance. Best practices require nurseries to purchase fresh mushrooms or truffles for positive identification and quality checks to ensure that no time is wasted inoculating with a non-target species. Buying pieces and scraps of poor quality truffles may increase the chances of less desirable truffle species becoming part of the inoculum. After positive identification of the truffle species, the ascoma containing spores are often placed in a blender with an abrasive such as ice or sand to help break the asci and release the spores from the asci (Iotti et al., 2012). These spores are then homogenized into the substrate the 8 trees are grown in. One key to successfully cultivating the correct target fungi is cleanliness in both the soil, the roots of the tree, the growing environment, and even the water source. The time needed between inoculation and good colonization may extend through an entire growing season before mycorrhization checks can be made and identification of the ectomycorrhizal fungus on the roots can be verified (Benucci et al., 2012). Non-target ectomycorrhizal fungi such as Sphaerosporella (Svrček) Svrček & Kubička may dominate the pot and make inoculation efforts ineffective (Iotti et al., 2012). In truffle tree production, it is standard to wait six months after inoculation before attempts are made to verify the presence of truffle on the roots (Benucci et al., 2012; Iotti et al., 2012). Verifying that the desired truffle species is on the roots of the host tree is important before out-planting since it may be five years or longer before truffle production begins and this would constitute a considerable loss of time for the grower anticipating truffle production. Pairing the right truffle with the right tree is also of considerable importance. Even though truffles are often thought of as generalists, there is some host specificity which needs to be considered. What Tree Plantations May Benefit Most? A pecan grower would be interesting in knowing that the European truffles T. aestivum and T. borchii form good ectomycorrhizal relationships with pecan roots in a nursery setting and should therefore consider these as viable options (Benucci et al., 2012). Additionally, the chestnut (Castanea sp. Mill.) industry is young in North America and in a prime position to begin looking at the possibility of adding value through the use of valuable, edible ectomycorrhizal fungi. Chestnut trees are in the Fagaceae family along with an important truffle producing trees in the Quercus genus. Quercus trees are known to be natural hosts to many ectomycorrhizal truffles such as T. borchii, T. aestivum, and T. melanosporum (García-Montero 9 et al., 2014; Reyna & Garcia-Barreda, 2014). It therefore appears that since Castanea spp. have been shown to host ectomycorrhizal relationships with many fungi, and that they are closely related to genera known to strongly associate with truffle species, that these trees may be a good candidate determining the potential of co-cropping with these high-value fungi. Additionally Corylus L. nut producers may also consider ways to increase the value of their farm since these trees are also ectomycorrhizal forming. Both T. magnatum Picco, the Piedmont white truffle, T. aestivum, and T. borchii are known to grow on the roots of trees in the Corylus genus (Bonito et al., 2010; Wang & Marcone, 2011). These trees have been widely used in establishing truffle orchards in Australia where the goal is to produce T. melanosporum, the European black truffle (Bradshaw, 2005). As this tree is susceptible to a myriad of disease pressures such as the devastating Eastern Filbert Blight which as caused by Anisogramma anomala (Peck) E. Müll., efforts to increase the value of these trees should be of great interest to growers looking to mitigate the risks associated with dependence on the full production of this nut (Pscheidt et al., 2018). For Corylus and all of the other trees mentioned here, the time to consider inoculations should begin while at the nursery stage since the best time to sway the community of the roots on a plant is when they are uncolonized and before they enter the diverse environment of the tree orchard. In the tree cropping systems utilizing compatible trees, there is an opportunity to greatly improve the value of these trees by forming a system whereby targeted, high-value fungi are cultivated on their roots. Understanding the need to verify the target ectomycorrhizal fungi prior to planting by looking at the colonization percentage and morphology based on some of the characteristics described in this thesis may prove to be important as North America matures its ectomycorrhizal cultivation. In conifer and nut tree nurseries, the benefits ectomycorrhizal fungi 10 impart on their host in terms of pathogen defense and nutrient and water uptake has generated much interest. The potential for increasing the value of these tree crops through the use of edible ectomycorrhizal fungi is too great to ignore and throughout Europe a great deal of work has been done to produce edible ectomycorrhizal producing trees. By adapting European methods for producing truffle trees in Michigan and for its tree crops, farms may benefit through the added revenue these high-value fungi will provide. In addition to providing a second crop, particularly culinarily valuable truffle species may also impart other benefits to the plantation in which they are grown. Weed Management The summer truffle, T. aestivum, can be found growing on the roots of Pinus and Quercus L. species. Growth of this truffle on both tree species results in the formation of a brûlé. A brûlé is used to describe the burned look seen in the vegetation growing in competition to the host tree and is associated with certain truffles species such as T. aestivum (García-Montero et al., 2014). It has been noted that the brûlé associated with T. aestivum is different in both size and intensity based on the associated tree species, for example, on Pinus, the brûlé is known to be over double the diameter that what has been seen under Quercus trees (García-Montero et al., 2014). This effect on the vegetation surrounding the host plant is the result of phytotoxic compounds produced by the truffle mycelium and the reduction in plant competition serves as one of the mutualistic returns this ectomycorrhizal fungus provides to the host tree (García- Montero et al., 2014). Beyond the cost of specialized equipment and herbicide chemicals, there is the ever-climbing cost of fuel and labor producers must consider, thus, a high-value edible mushroom producing ectomycorrhizal fungus that has an affinity for killing weeds probably sounds too good to be true. In practice, though reduced, weed management practices are still 11 needed since these brûlé formations are not impervious to all weed growth, nor are these formations uniform in dispersal beneath the trees since the colonization by T. aestivum may not be uniform. Weed plants reduce colonization and truffle production (Mamoun & Oliver, 1997; Olivera et al., 2011) and therefore many orchard operators manage weeds either mechanically (Olivera et al., 2014) or chemically (Gómez-Molina et al., 2020). Offering significant savings in time and labor over manually removing weeds, many cropping systems utilize herbicides to increase crop productivity. Herbicides play important roles in agriculture by reducing competition for nutrients, water, and sunlight. This type of pesticide can also prove critical for use in controlling invasive plant species which in turn aids in conservation efforts to preserve native ecosystems. Though there has been work to develop fungal based bioherbicides (Julia et al., 2022), most herbicides are produced with synthetic chemicals. Despite these benefits, there are also many potential side effects related to the use of herbicides and many unknown outcomes especially in relation to fungal cultivation. Herbicide Sensitivity in Fungi There have been numerous studies showing that fungi can be sensitive to chemical exposure. For example, glyphosate inhibits 5-enolpyruvyl shikimic acid-3-phosphate synthase (EPSPS) which is one reason it is such an effective herbicide (Steinrücken & Amrhein, 1980). However, in addition to plants, both bacteria and fungi have the shikimic acid pathway (Gupta & Crissman, 2013). As such, it stands to reason that herbicide overspray containing glyphosate may impact organisms other than the targeted weed plant. Currently there have been no studies directly exploring fungal EPSPS interactions with glyphosate (Hammerschmidt, 2018). Paraquat (1,10-dimethyl-4,4-bipyridinium dichloride), has been shown to interfere with biofilm and 12 melanin production in Cryptococcus Vuill. (Castelo-Branco et al., 2022). Flurochloridone and prosulfocarb inhibit conidial germination of Beauveria bassiana (Bals.-Criv.) Vuill. (Celar & Kos, 2016). Pleurotus ostreatus (Jacq.) P. Kumm. experiences mycelial growth reduction with glyphosate (Connelly et al., 2019), and glufosinate ammonium has been shown to reduce mycelial growth of Guignardia bidwellii (Ellis) Petr. by nearly 80% (Albrecht & Kortekamp, 2009) to name a few examples. There has been sparse work done investigating conventional herbicide-fungal interactions, and even less investigating organic herbicide-fungal interactions. These sorts of interactions should be further investigated as many agroforestry mushroom growers may consider organic herbicides as a safer alternative, but it is unclear how these herbicides impact growth or consumer safety. With any pesticide, there is also the possibility residuals may be found on or in mushrooms which were previously exposed. This could have great health implications as many pesticides are not intended for human consumption. Recent publications outline reliable methods to identify over 100 pesticides in mushroom tissue (Cao et al., 2016; Le et al., 2021). One alarming finding Le et al. (2021) found was that various pesticides were detected in every Lentinula edodes (Berk.) Pegler, Agaricus bisporus (J.E. Lange) Imbach, and Pleurotus ostreatus mushrooms purchased for this study (Le et al., 2021). This includes the herbicides Atrazine, MCPA, Dicamba, Diuron, 2,4-D, and Cyhalofop Butyl (Le et al., 2021). Le et al. (2021) postulate that these pesticides enter the mushroom via residuals on the substrate from which these fungi are cultivated. Pesticide chemistries, their interactions with edible fungi and human health implications will vary product to product suggesting an area potentially rich for further study. Despite its use, there are still many questions about how the use of herbicides may impact fungi including those beneficial to plant growth, ectomycorrhizal fungi, and even saprotrophic fungi. 13 Herbicide-fungal interactions are discussed more in Chapter 2, “Impacts of Glufonisate- Ammonium and Glyphosate Herbicides on Mycelial Growth of Truffles, Morels, and Plant Growth Promoting Fungi.” Fungal Priority Effects Additionally, growers must also contend with non-target weed fungi if they are trying to cultivate a specific edible mycorrhizal species in their agroforestry system. As with weed plants, weed fungi compete for space, nutrients, and water. Priority effects also play an important role in the establishment of target species and subsequent development. Fumigated bare-root outdoor tree nurseries offer an opportunity to investigate priority effects. However, as this is an outdoor cropping system, there are many opportunities for non-target fungal species to take advantage of priority effects. There is a need to characterize and investigate priority effects in bare-root tree nurseries as well as a need to learn which native ectomycorrhizal species may be present and able to disrupt target-species colonization. These are further discussed in Chapter 3, “Does Ectomycorrhizal Inoculum After Methyl Bromide Field Applications in Bare-Root Conifer Nursery Seedlings Establish and Persist?” The need to better understand non-target ectomycorrhizal fungi does not stop at the plantation though, there are a wealth of new species to be discovered in native tree stands as well. New Ectomycorrhizal Species Characterizing fungal species include examining spore size, spore shape, spore ornamentation, and determining that purported new species yields a unique genetic sequence. Many truffle species are yet to be described and a global meta-analysis was conducted by Bonito et al. (2010) of over 2000 Tuber ITS rDNA sequences to offer a framework by which future studies may be built. This 2010 study predicted between 180 and 230 Tuber species globally and 14 used numerical designations for species not yet described (Bonito et al., 2010). Truffles, as with many fungi, often use the internal transcribed spacer (ITS) as the primary region used to differentiate species. There are limitations to building Tuber phylogenies with the ITS region and therefore primer sets have been developed for assist with multigene phylogenies including elongation factor 1a (EF1α_Tuber_f, EF1α_Tuber_r), and 2nd subunit of RNA polymerase (RPB2_Tuber_f, RPB2_Tuber_r)(Bonito et al., 2013; Bonito et al., 2010). Together with morphological characteristics, phylogenetics offer a powerful tool for researchers to better understand how taxa relate to each other. There are numerous reasons why it is important to describe novel taxa. Firstly, we know the importance fungi play in nearly every ecosystem. By expanding our knowledge of fungal diversity we better understand their ecological roles and functions in sustaining the health of ecosystems. Secondly, fungi provide countless industrial and medical applications which improve our daily lives. Through understanding and exploring fungal species diversity, we have the opportunity to discover solutions to problems we face. Third, by describing new species, we better develop fungal taxonomy. This is important because it helps us develop better understanding of evolutionary relationships which in turn helps with the identification of species. Finally, as our climate is changing and ecosystems shrink or disappear, describing novel taxa gives us the chance to conserve threatened or endangered species through outreach and conservation efforts. Research focus The three primary goals pursued in this thesis are: 1. Determine mycelial growth in media containing herbicides 15 Hypothesis: (H1) Herbicides containing either glyphosate or glufosinate will impact the mycelial growth of truffles, morels, and plant growth promoting fungi. (H2) There will be variability of response between saptrotrophic and ectomycorrhizal ecologies. (H3) Both herbicides act fungistatically at concentrations near or above the manufacturers recommended application rate for field use. 2. Investigate ectomycorrhizal priority effects in methyl bromide fumigated bare-root Christmas tree nurseries. Hypothesis: (H4) Pseudotsuga menziesii (Mirb.) Franco, Picea abies (L.) H.Karst., and Pinus sylvestris L. can be colonized by exotic ectomycorrhizal fungal species. (H5) There will be little to no significant increases in above ground biomass, tree height, or root collar diameter based on inoculant. (H6) Either by persisting through fumigation or through blowing in, local ectomycorrhizal species will dominate the root systems of Pseudotsuga menziesii, Picea abies, and Pinus sylvestris. 3. Confirm and describe a new species of truffle in the Tuber genus Hypothesis: (H7) Morphology and molecular analysis will support Tuber rugosum Rennick B, Benucci GMN, Du Z, Healy, & Bonito as a novel species in the Rufum clade. (H8) This new species will have a unique fatty acid profile. (H9) Utilizing observed fungivory in slugs may provide an alternative ascus removal for high quality scanning electron microscopy. (H10) Based on numerous new species within the Rufum clade, an updated taxonomic key can be produced. Value of this research The research outlined within this thesis may serve to inform management of truffières and other fungal cultivation systems that may consider the use of herbicides to combat weed plants. Many truffle plantations are already using herbicides and will be interested to know how 16 over applying may lead to fungistatic situations. Furthermore, there is a growing interest in morel cultivation in fields between agronomic crops, this work begins to inform sensitivities of edible fungi to herbicides of which there may still be residuals. Other research covered within this thesis outlines what outcomes may occur if a post methyl bromide fumigated bare-root Christmas tree nursery is interested in adding mycorrhizal inoculum to their fields. As a potentially cost prohibitive activity, growers will be interested in knowing that there are a strong group of ectomycorrhizal beneficial symbionts either persisting through fumigation or blowing in from nearby tree stands. This research also provides evidence that while in-field inoculation of exotic Tuber species is possible, the rates are far below standards set by the truffle tree industry. Decades of efforts to cultivate truffles in North America have proved highly challenging. Understanding competing ectomycorrhizal fungi requires that little or unknown species are properly described and subsequently identified to further make sense of how their ecologies and ranges may impact a truffières ability to establish and persist. Beyond the value describing new ectomycorrhizal species has to truffières, there are many potential benefits gained by understanding their ecological roles and functions. Developing better fungal taxonomy helps improve our understanding of evolutionary relationships and may inform our ability to identify rare taxa for conservation purposes. Cohesively, research presented in this thesis will impact our understanding on the ecology, taxonomy, scanning electron microscopy preparation, herbicide sensitivity, and priority effects of truffles. 17 Chapter 2 Impacts of Glufonisate-Ammonium and Glyphosate Herbicides on Mycelial Growth of Truffles, Morels, and Plant Growth Promoting Fungi 18 Abstract Herbicides are frequently used to control weeds in agricultural and residential settings, however, herbicides can also impact fungi. Understanding how weed management impacts soil fungi may help reduce unintended consequences of these chemicals in agricultural settings. In this study we assessed the impacts of glyphosate [N-(phosphonomethyl)-glycine] (Roundup) on vegetative growth of soilborne edible and plant growth promoting fungi including Benniella erionia, Linnemannia elongata, two species of morels (Morchella), six species of truffles (Tuber). We also assess the impact of glufosinate-ammonium (Liberty), on vegetative growth of two species of morels (Morchella), three species of truffles (Tuber), and two soil fungal species within Mortierellaceae (Benniella and Linnemannia). Each fungal isolate was grown in a common medium containing an herbicide ranging in concentrations spanning above and below the application rate recommended by the manufacturer. To determine the impact of these chemistries on fungal growth, hyphal growth rates on agar media and fungal biomass in liquid media were collected. The fungi grew unimpeded at low herbicide concentrations but were completely inhibited at higher herbicide concentrations. The recommended working glyphosate concentration was found to be near the point of fungal growth inhibition in-vitro. However, the fungi tested here were far less sensitive to glufosinate. Low amounts of this foliar spray would be expected to drench the soils these fungi inhabit when applied during typical application. As such, the use of these herbicides as a foliar spray at recommended rates during typical application is unlikely to interfere with the mycelial growth of the fungal species assayed in this experiment. Nonetheless, use of these herbicides at concentrations higher than recommended could inhibit mycelial growth and fungal biomass in soil environments. 19 Introduction Glyphosate, a systemic pesticide, is the active compound in popular commercial herbicides, and is intended for application to plant photosynthetic surfaces where it is highly phototoxic (Coupland & Caseley, 1979). Within fungi, bacteria, and plants, the shikimic acid pathway feeds into the production of the amino acids tryptophan, tyrosine, and phenylalanine (Gupta & Crissman, 2013). Glyphosate is an inhibitor of 5-enolpyruvyl shikimic acid-3- phosphate synthase (EPSPS) and as such, has great value as an effective herbicide (Steinrücken & Amrhein, 1980). Glufosinate, a contact pesticide, is used in many popular herbicides. Glufosinate on the other hand, is a contact herbicide, that inhibits the production of glutamine synthetase, which leads to the buildup of ammonia in cells leading to cell death (Gupta & Crissman, 2013; Herrmann & Weaver, 1999). Glufosinate does not appear to inhibit the growth of the oomycete Pythium aphanidermatum (Edson) Fitzp., but has shown fungicidal activity against Rhizoctonia solani J.G. Kühn and Sclerotinia homoeocarpa F.T. Benn. indicating variability in response should be expected (Liu et al., 1998). Many agricultural production systems rely upon beneficial soil fungi to sustain crop yields, while others may be specifically focused on the production of fungal fruiting bodies. For example, there have been many successes in the cultivation of truffles (Tuber spp.) and other ectomycorrhizal fungi (Lactarius spp.) over the past several decades (Chevalier & Pargney, 2014; Wang et al., 2012). Notable successes in truffle cultivation include productive T. melanosporum Vittad. plantations (truffières), particularly in Italy, France, Spain, Australia and New Zealand (Hall et al., 2017). In 2019 and 2020, T. magnatum Picco was detected and harvested in truffières outside the natural range in France (Bach et al., 2021). Similarly, new outdoor cultivation of edible and medicinal morels (Morchella spp.) have had recent successes 20 (Dissanayake et al., 2021; Xu et al., 2022). As with many plant cropping systems, fungi cultivated in soil must contend with weed plants for water and nutrients (Olivera et al., 2011, 2014a). For instance, truffle cultivation involves planting seedlings inoculated with Tuber into prepared calcareous field soils, which must be specifically managed to improve and maintain the production of truffles (Chevalier & Pargney, 2014). Standard management of truffle orchards requires that weeds be removed around host trees to aid in moisture retention and to reduce other forms of biotic competition within the soil (Chevalier & Pargney, 2014; Olivera et al., 2011). While mechanical weed removal is the standard method in truffle orchards, chemical management of weeds in truffle orchards, such as with glyphosate and other herbicides, is becoming more common due to its low cost and high efficacy (Gómez-Molina et al., 2020). There is still little known about the effects of this herbicide on fungal growth. For example herbicides may enhance or suppress fungal disease development, in part due to the influence that the application of the herbicide may have over all facets of the disease triangle (Hammerschmidt, 2018). In particular, rust fungi are inhibited in crops where glyphosate is used, however, suppression is possibly linked to the obligate biotrophic relationship these fungi have with their host plants as these fungi require living plant cells to live (Hammerschmidt, 2018). However, during herbicide application, herbicides also come into contact with the soil and though it quickly adheres to soil minerals, some herbicide can make its way deeper into the soil (Jonge et al., 2000). Both glyphosate and glufosinate have also been found to be translocated through the plant from the foliage down into the roots and where it is exuded into the surrounding rhizosphere (Feng et al., 1999; Laitinen et al., 2007; Steckel et al., 1997). Soil organisms, especially mycorrhizal fungi, may then be exposed to these xenobiotic compounds. For example, (de Novais et al., 2019) found herbicide exposure reduced extramatrical hyphal 21 density, explored area, length, instances of anastomosis, and hyphal branching among of mycorrhizal networks of Cichorium intybus L. by Funneliformis mosseae (T.H. Nicolson & Gerd.) C. Walker & A. Schüßler. In addition, glyphosate has been shown to decrease arbuscular mycorrhizal fungi colonization among Abelmoschus (Brindhavani et al., 2018), Trifolium (Zaller et al., 2014), Festuca (Helander et al., 2018) and to decrease T. melanosporum spore viability amongst truffle inoculated trees (Gómez-Molina et al., 2020). There is far less information available regarding glufosinate interactions with soil inhabiting fungi. To address this gap of knowledge, this study aimed to further investigate direct impacts of the commercially available products glyphosate (Roundup) and glufosinate (Liberty) herbicides on the vegetative growth of diverse fungi including morels (Morchella spp.), truffles (Tuber spp.) and select soil-inhabiting fungi. We hypothesize that fungi would be sensitive to both chemistries, but that mycorrhizal fungi would be less sensitive than saprotrophic fungi as other studies have indicated established Tuber colonies persist after recommended rates of glyphosate applications (Bonet et al., 2006; Gómez-Molina et al., 2020; Olivera et al., 2011). Methods Direct impacts of commercially available glufosinate and glyphosate herbicides on fungal vegetative growth were tested on pure culture isolates through two experiments. In the first experiment, herbicides were incorporated into an agar-based plate assay to assess fungal growth rates in this semi solid matrix. In the second experiment, a broth-based assay was used to assess fungal biomass at different concentrations of herbicide. Benniella erionia Liber & Bonito, L. elongata (Linnem.) Vandepol & Bonito, M. americana Clowez & Matherly, M. importuna M. Kuo, O'Donnell & T.J. Volk, T. borchii Vittad., T. canaliculatum Gilkey, T. floridanum A. Grupe, Sulzbacher & M.E. Sm., T. gibbosum Harkn., T. lyonii Butters, and T. rugosum Rennick 22 B, Benucci GMN, Du Z, Healy R, & Bonito G. were chosen for this study to represent several trophic ecologies as described in detail below and summarized in Table 1. Additionally, Rhizopus oryzae Went & Prins. Geerl., Serendipita indica (Sav. Verma, Aj. Varma, Rexer, G. Kost & P. Franken) M. Weiss, F. Waller, Zuccaro & Selosse, Stropharia rugosoannulata Farl. ex Murrill, and Umbelopsis sp. were used in a pilot experiment at lower replication, further detailed in FIGURE 2.6. 23 Table 2.1 - Fungal isolates and herbicides used in this study. 24 Experiment 1 Solid Media Solid Media Preparation Assays were carried out on half strength potato dextrose agar (PDA½) and quarter strength potato dextrose agar (PDA¼). The PDA½ recipe per liter is as follows: 12.0g potato dextrose broth (PDB), 1.0g yeast extract (YE), 10.0g agar, double distilled water filled to 1000.0mL, then autoclaved. The PDA¼ recipe per liter is as follows: 6.0g PDB, 1.0g YE, 10.0g agar, double distilled water filled to 1000.0mL, then autoclaved. Glyphosate herbicide (Roundup Ready-to-Use Weed and Grass Killer) was filter sterilized at 0.2 microns, and once media were cooled to 45 o C, was added at the following concentrations 0.001 mL/L, 0.01 mL/L, 0.1 mL/L, 0.0 mL/L, 1.0 mL/L, 10.0 mL/L, and 40.0 mL/L. The herbicide bottle provided an application rate of 30fl oz/70ft 2 (887mL/6.5m2 ). The surface area of a Petri dish is 78.54cm2 thus an application rate of 1.07mL/78.54cm2 was chosen as a central concentration. Glufosinate (Liberty 280 SL) herbicide was too viscous to filter sterilize and was therefore added to the agar mediums at 55 o C in the following concentrations: 0.001 mL/L, 0.01 mL/L, 0.1 mL/L, 0.0 mL/L, 1.0 mL/L, 10.0 mL/L, and 40.0 mL/L. The herbicide bottle provided an application rate of 20fl oz/ac (650.6mL/4046.9m2 ). Disposable 100 mm Petri dishes were filled to 25.0 mL using a serological pipette. No contamination was observed. A 4.0 mm cork borer was used to transfer fungal isolates including M. americana (BR3), M. importuna (GB772G), B. erionia (GB_AUS_27b), and L. elongata (GB_AUS_24) onto the center of PDA½ media containing glufosinate or glyphosate herbicides at the concentrations described above. The agar removed from the mother culture was such that the front of the colony's growth was centered across the diameter of the plug cut by the cork borer. Similarly, 25 with a 4.0 mm cork borer, fungal isolates including Tuber borchii (BR11), Tuber borchii (GMNB230), Tuber gibbosum (GMNB46) were transferred onto the center of PDA¼ media containing glufosinate or glyphosate also at concentrations of 0.001 mL/L, 0.01 mL/L, 0.1 mL/L, 0.0 mL/L, 1.0 mL/L, 10.0 mL/L, and 40.0 mL/L. Benniella erionia, L. elongata, M. americana, and M. importuna was plated with both herbicide brands (n=2) for each concentration (n=6) with three replicates totaling 144 Petri dishes. Additionally, two isolates of T. borchii and one isolate of T. gibbosum were plated with both herbicide brands with all six concentrations and plated in five replicates totaling 180 Petri dishes. Petri dishes were incubated at room temperature (23 C) in a dark location. Plates were randomly distributed within a dark box to prevent any block effects. To measure fungal growth, four quadrants on each Petri dish were marked and daily measurements of the longest hyphal growth within each quadrant were collected for 14 days amongst the fast growing Morchella, Benniella and Linnemannia isolates. For the slower growing Tuber isolates, hyphal growth within each quadrant was collected weekly for an 8 week duration. Experiment 2 Liquid Media Liquid Media preparation Two liquid media were formulated for the second experiment. The first media, referred to as Tuber media (TM) in this experiment, was chosen because it generally supports growth of ectomycorrhizal Tuber species. Tuber media consisted of 5.0g potato dextrose broth, 5.0mL glycerol, and 0.82g calcium nitrate then filled with double distilled H 2 O to 1000mL. Prior to autoclaving the TM, pH was brought up to 7.5 with 5M sodium hydroxide. Once the temperature of the media after autoclaving fell below 45°C, 1mL/L biotin (0.5g/L stock) was added. The 26 second medium, malt extract broth (MEB), was prepared using 10.0g malt extract and 1.0g yeast extract, and double distilled water filled to 1000.0mL, then autoclaved. Once the TM and MEB temperature reached room temperature (23° C), glyphosate herbicide was filter sterilized at 0.2 microns and added at the following concentrations: 0.0mL/L, 0.0456 mL/L, 0.456 mL/L, and 4.65 mL/L. These concentrations were chosen based upon herbicide label coverage information adjusted to the surface area of the liquid media within the 125.0 mL Erlenmeyer flasks used for this experiment. The label on the herbicide bottle recommends a coverage of 73.32cm 2 /mL. The surface area of the liquid within the flasks is 33.44cm2 . Thus a ratio of 33.44cm2 /73.32cm2 /mL (836 : 1833) gives a working concentration of 0.456mL, which was set at the middle strength concentration for this experiment. At each concentration of herbicide 50mL of TM or MEB was aliquoted into Erlenmeyer flasks. Using a 7.5mm cork borer, a single piece of the following isolates was added to each flask. Five species of truffles were screened in the TM: Tuber floridanum (BR61c), Tuber rugosum (BR64a), Tuber canaliculatum (BR7), T. borchii (BR25), and Tuber lyonii (GB17). Four isolates of Morchella and other soil fungi were screened in MEB including: Morchella americana (BR3), Morchella importuna (GB772G), B. erionia (GB_AUS_27b) and L. elongata (GB_AUS24). Five replicates for each Tuber isolate and ten replicates each of Benniella, Linnemannia, and both Morchella isolates at each herbicide concentration (4) were carried out for a total of 260 flasks in this experiment. As in the aforementioned agar experiment, the agar removed from the mother culture was such that the front of the colony's growth was centered across the diameter of the plug cut by the cork borer. All replications for each isolate came from the same mother culture from the same Petri dish such that the mass of the agar plug would be consistent. Flasks were plugged with sterile cotton that had been wrapped in gauze and covered with aluminum foil to allow gas exchange while 27 preventing contamination. These flasks were stored randomized in a dark room on a shelf at room temperature (23o C) and were gently swirled and re-randomized every two days. After 7 days of growth, each flask was vacuum filtered with a Büchner funnel whereby the medium was poured through pre-massed sections of miracloth leaving the mycelial mass and agar plug behind. Prior to use, miracloth (EMD Millipore Corp., Burlington MA, USA) sections were fully desiccated by running them through a lyophilizer and stored in a bag with silica beads. These desiccated miracloth sections were massed for later use. The pre-massed miracloth containing the fungal tissue was stored in a 15mL centrifuge tube and placed in a lyophilizer at - 40o C and -0.01kPa for 48 hours. After lyophilization, tissue was stored in a bag with silica beads to maintain the low moisture content. Miracloth containing fungal tissue was massed and the mass of the miracloth was subtracted from the total mass such that the only mass used in subsequent calculations was the fungal growth and the initial agar plug. Statistical Analysis Data were analyzed and visualized in RStudio (Version 2021.09.1+372 "Ghost Orchid") running R (version 4.1.2), using ggplot 2 (version 3.3.5), ggpubr (version 0.4.0) plyr (version 1.8.6), and readxl (version 1.3.1). Means of treatment groups were compared using a t-test with the reference set as the negative control group. Means were compared by one-way analysis of variance based on a completely randomized design. 28 Results Experiment 1 Solid Media FIGURE 2.1 - REPRESENTATIVE SELECTIONS OF ISOLATES. Here, Benniella eronia (GB_AUS_27b), Linnemannia elongata (GB_AUS_24), Morchella americana (BR3), and Morchella importuna (GB772G) were grown on solid media at seven different concentrations of 29 glyphosate-based herbicide or glufosinate-based herbicide. Hyphal length measurements were collected at four marked poles of each plate. In the first experiment, impacts of the glufosinate and glyphosate herbicides on fungal growth in solid media were tested for seven isolates from four genera. Between 0.1mL/L and 1.0mL/L concentrations, hyphal extension length was similar but mycelial density was reduced as the herbicide concentration increases. There was no hyphal growth for any isolate at herbicide concentrations of 40.0 ml/L (Fig 2.1). FIGURE 2.1 illustrates fungal growth response along the range of colonization from the lowest concentrations to the highest concentrations of the herbicide. FIGURE 2.1 shows a representative for each treatment, however, FIGURE 2.7 and FIGURE 2.8 show all repetitions. While the data collected highlight growth in terms of length from inoculation point, they do not capture mycelial density, hyphal diameter, or aerial growth. However, the reoccurring measurements shown in FIGURE 2.2 and FIGURE 2.3 illustrate the delay in growth and the threshold concentration of herbicide that suppressed the growth of the truffle fungal isolates. were The highest concentration of glufosinate that supported growth of Tuber borchii (BR11 and GMNB230) and T. gibbosum (GMNB46) was 0.1mL/L , and growth was delayed or reduced as herbicide concentration increased as seen in FIGURE 2.2. Among the concentrations tested, T. borchii (BR11) and T. gibbosum (GMNB46) were unable to grow vegetatively above 0.01mL/L glyphosate. 30 FIGURE 2.2 – DAILY GROWTH OF TUBER BOCHII AND TUBER GIBBOSUM. Average hyphal growth of Tuber borchii (BR11 and GMNB230)) and Tuber gibbosum (GMNB46) grown in solid media at five concentrations of either glufosinate or glyphosate herbicides. At 1.0mL/L concentrations of either herbicide there was no measurable growth for both Tuber isolates. At 0.1mL/L concentrations of glyphosate, there was no measurable growth for either Tuber isolate. 31 FIGURE 2.3 - DAILY GROWTH OF SAPROBIC FUNGI. Hyphal growth of Benniella eronia (GB_AUS_27b), Linnemannia elongata (GB_AUS_24), Morchella americana (BR3), and Morchella importuna (GB772G) grown on solid media at seven different concentrations of either glufosinate or glyphosate brand herbicides. No isolate grew on media containing 40.0mL/L of either herbicide. Neither Benniella nor Morchella isolates grew on media containing 10.0mL/L of either herbicide. Linnemannia elongata was the most tolerant of the saprotrophic fungi assayed, with growth in 10.0mL/L of both glufosinate or glyphosate herbicide (FIGURE 2.3), whereas, neither Morchella 32 species grew on media with herbicide concentrations above 1.0mL/L. None of the Morchella species nor L. elongata grew at 40.0mL/L, and all isolates appeared to grow equally fast at herbicide concentrations below 0.01mL/L independent of herbicide brand. Experiment 2 Liquid Media In the second experiment, the impacts of glyphosate herbicide on fungal growth in liquid media was tested for nine isolates from nine genera. Among the nine fungal isolates tested, no discernable mycelium was visible at the highest concentration of glyphosate herbicide (4.561mL/L). There was a relatively steady decrease in visible growth from the lowest concentration to the highest concentration for isolates grown in MEB (Fig. 2.9). Linnemannia elongata (GB_AUS_24) grew on the surface in the negative control flask (Fig. 2.9). This fluffy growth is typical and was seen in every replicate, however, no surface growth was observed in any flask containing glyphosate herbicide. Similar growth was observed in all saprotrophic fungi (FIGURE 2.4) but was not observed uniformly amongst the mycorrhizal fungi. The biomass (FIGURE 2.5) produced in the working concentration of 0.451mL/L was significantly less than that of the negative control for B. erionia (P<0.001), L. elongata (P<0.001), M. americana (P=0.0019), M. importuna (P<0.001), T. rugosum (P<0.001), and T. lyonii (P=0.0047). 33 FIGURE 2.4 – BIOMASS OF FUNGI GROUPED BY TROPHIC ECOLOGIES. Dosage responses of T. borchii, T. canaliculatum, T. floridanum, T. lyonii, T. rugosum, B. erionia, L. elongata, M. americana, and M. importuna across four concentrations of glyphosate herbicide. Saprotrophic species (B. erionia, L. elongata, M. americana, and M. importuna) exhibit more 34 FIGURE 2.4 (cont’d) – predictable growth declines as concentration increases whereas the Tuber species show variable reactions to the three lower concentrations. 35 FIGURE 2.5 - BIOMASS OF FUNGAL ISOLATES. Each plot shows the biomass of Tuber borchii (BR25), Tuber canaliculatum (BR7), Tuber floridanum BR61c), Tuber lyonii (GB17), Tuber rugosum (BR64a), Benniella erionia GB_AUS_27b), Linnemannia elongata FIGURE 2.5 36 FIGURE 2.5 (cont’d) - (GB_AUS_24), Morchella americana (BR3), and Morchella importuna (GB772G) when grown in four concentrations of glyphosate herbicide in liquid media. The mycorrhizal species were grown on a medium referred to as Tuber media (TM) whereas the saprotrophic species were grown in malt extract agar. FIGURE 2.6 – AVERAGE HYPHAL LENGTH MEASUREMENTS OVER A 14-DAY PERIOD OF RHIZOPUS ORYZAE, SERENDIPITA INDICA, STROPHARIA RUGOSOANNULATA, AND UMBELOPSIS SP. ON SOLID MEDIA CONSISTING OF 37 FIGURE 2.6 (cont’d) - SEVEN CONCENTRATIONS OF EITHER GLUFOSINATE OR GLYPHOSATE BRAND HERBICIDE. These were not replicated and therefore do not hold statistically significant information, however, the trends amongst these soil inhabiting fungi mirror those seen in FIGURE 2.2 and FIGURE 2.3. Rhizopus oryzae was the only taxa in this study to grow on media containing 10.0 mL/L glyphosate herbicide. 38 FIGURE 2.7 - REPLICATIONS OF BENNIELLA ERIONIA, LINNEMANNIA ELONGATA, MORCHELLA AMERICANA, AND MORCHELLA IMPORTUNA GROWING ON MEDIA WITH GLYPHOSATE HERBICIDE AT 7 DIFFERENT CONCENTRATIONS. 39 FIGURE 2.8 - REPLICATIONS OF BENNIELLA ERIONIA, LINNEMANNIA ELONGATA, MORCHELLA AMERICANA, AND MORCHELLA IMPORTUNA GROWING ON MEDIA WITH GLUFOSINATE HERBICIDE AT 7 DIFFERENT CONCENTRATIONS. 40 FIGURE 2.9 – GROWTH OF LINNEMANNIA ELONGOTA IN BROTH WITH HERBICIDE AT FOUR CONCENTRATIONS. Only the media without any herbicide displayed thick characteristic growth above the liquid surface. Media with herbicide at 0.04561mL/L did have growth, but it remained solely within the broth. Media at 0.4561mL/L and 4.561mL/L had little if any observable hyphal growth. Discussion Globally, an estimated 950,000 tons of chemical herbicides are applied to land annually (Sharma et al., 2019). In fact, glyphosate is one of the most common herbicides used in agriculture. It is a broad-spectrum systemic herbicide, a phosphonate, which interrupts the shikimate pathway. Although the impacts of this herbicide on plants and animals is fairly well studied, impacts of these chemicals on non-target soil biota is less well understood. To address this, in this study we tested the impact of glyphosate and glufosinate on pure culture isolates of 41 diverse soil fungi including saprotrophic morels (Morchella), ectomycorrhizal truffles (Tuber), and beneficial root symbionts (Mortierellaceae; Sebacinales) of agricultural plants. In this study the impacts of herbicides were tested on different fungal guilds including plant growth-promoting fungi, ectomycorrhizal fungi, and soil saprotrophs. Among isolates tested, ectomycorrhizal fungi were the least sensitive to these herbicides, and only showed detrimental growth effects at concentrations at or above the rate of application (0.456mL/L) as per the herbicide label suggestions. This finding supports other studies that found little to no effect in mycorrhizal status or extraradical hyphae density with moderate applications of glyphosate (Bonet et al., 2006; Chakravarty & Chatarpaul, 1990; Gómez-Molina et al., 2020; Olivera et al., 2011). Interestingly, the saprotrophic fungi assayed here were more sensitive to glyphosate and showed deleterious effects at concentrations lower than the recommended application rate. One possible explanation is that ectomycorrhizal species grow slowly in vitro and therefore have more time to either process the herbicide or more time to produce the amino acids disrupted by the disruption of the shikimate pathway. Future work focusing on the effects of glyphosate on the fungal shikimate pathway may yield more clues. In non-agricultural settings where hand applications of glyphosate are commonplace, it is therefore important to prevent overapplication of this herbicide to prevent concentrations where fungistatic or fungicidal outcomes are observed. Our results demonstrate that the herbicide glyphosate can impact fungal vegetative growth when applied at concentrations above the recommended concentration. Glyphosate has also been shown to influence other organisms, such as by decreasing melanization and increasing infection rates among the moth Galleria mellonella L. and mosquito Anopheles gambiae Giles (Smith et al., 2021). There have been numerous studies outlining varied outcomes of glyphosate 42 applications on earthworms such as weight loss (Correia & Moreira, 2010), negative reproductive influences (Domínguez et al., 2016), avoidance response driven migration (Verrell & Van Buskirk, 2004), and decrease mycorrhization by arbuscular mycorrhizal fungi in mesocosms with Trifolium repens L. and earthworms (Zaller et al., 2014). Despite the growing breadth of knowledge into non-target herbicide interactions, there is still clearly more that needs to be studied. Outdoor morel cultivation has garnered much interest recently, particularly since the first successful artificial outdoor cultivation of morels in 2012 in Sichuan, China (Xu et al., 2022). One appealing facet of morel cultivation are improvements to the soil and given morels’ short growing period they fit neatly into certain rotating cropping systems (Su et al., 2022). As morel cultivation further expands into more crop rotations, further work is needed to identify interactions with the pesticides used with the preceding crop. The use of herbicides, such as those containing glyphosate, decrease fungal biomass in the soil as well as cultivable fungi species richness (Vázquez et al., 2021). However, there is still much work to be done regarding pesticides and their implications on plant growth-promoting fungi. In conclusion, we used pure fungal cultures to assess the impact of glyphosate and glufosinate on fungal growth in solid and liquid media. We found that as concentration of glyphosate or glufosinate increased in the growth media, the herbicide decreased fungal network density, branching and biomass. Interestingly, we found that the saprotrophic fungi tested were more susceptible to glyphosate compared to ectomycorrhizal species. Taken together, these results indicate that glyphosate herbicides can impact fungal populations if used at concentrations higher than recommended by the producers. Glyphosate is sometimes used in truffle orchards to help control weed establishment around planted seedlings. It is also used in 43 agricultural systems where plant growth-promoting fungi or other edible fungi may be of interest. Thus, care should be taken if applying glyphosate herbicides in agricultural systems where fungal populations are of interest to limit overapplication. If applied at recommended concentrations, our results indicate that commercial blends of glyphosate or glufosinate may not negatively impact the vegetative growth of the fungal mycelium growing in treated soils in the short-term. However, further research will be needed to determine the fate of herbicides in complex real world environmental ecosystems. 44 Chapter 3 Does ectomycorrhizal inoculum after methyl bromide field applications in bare-root conifer nursery seedlings establish and persist? 45 Abstract Conifer seedlings are widely grown in the midwestern United States for Christmas tree plantations, landscape nurseries and reforestation. Seedlings are typically grown for two years prior to lifting and replanting. Many of these bare-root nurseries have been in business for many decades and typically sow seed in the same soil cycle after cycle. In part to reduce disease pressure, but also to reduce weed seed banks, some nurseries apply methyl bromide with chloropicrin as a fumigant to eradicate these pests. With concern about this fumigant destroying not only pests but also beneficial mycorrhizal fungi, there is interest in whether the application of mycorrhizal spores following fumigation is worthwhile. We used this fumigation-cropping cycle to investigate how introducing local, exotic, or commercial inoculants may influence health, height, and biomass of the seedlings grown in these fumigated soils. Based on investigating two nurseries over two cropping cycles, our findings indicate that there are no disease reduction or growth increases gained regardless of any mycorrhizal inoculants tested. Rather, we found a robust diversity of mycorrhizae, beyond our inputs, infesting the roots of Pseudotsuga menziesii, Picea abies, and Pinus sylvestris irrespective of whether the inoculum was from local, exotic, or commercial sources. These findings suggest that there are ectomycorrhizal fungi either surviving fumigation or blowing in from nearby tree stands. The major finding of this study is that Norway spruce, Douglas-fir, or Scots pine grown in a bare-root nursery do not benefit from the addition of additional ectomycorrhizal spores. Introduction Currently, there are approximately 300,000 acres of cultivated conifer trees grown on 15,000 farms in the United States (USDA, 2017). Combined with Europe, over 80 million trees are grown every year, many of which begin life in bare root nurseries (Chastagner & Benson, 46 2000). As with any high-density cropping system, pests and pathogens can become a major issue in conifer nursery beds. The most concerning pathogens to conifer tree nurseries cause root rots and damping off diseases. These diseases reduce production yields and quality of seedlings and are often seen as chlorosis, stunting, and necrosis (Weiland et al., 2016). Fungal species of Fusarium Link, Cylindrocladium Morgan, and Rhizoctonia DC. and Oomycete species of Phytophthora de Bary and Pythium Pringsh. are the most commonly reported causal agents of seedling death amongst nurseries (Cram, 2015). Oomycetes such as Pythium and Phytophthora, once thought to be fungi, are in the kingdom Chromista. Unlike fungi, oomycetes lack chitin in their cell walls and live primarily as diploid organisms. This group hosts some of the most important pathogens whose motile zoospores cause several damping-off diseases in nurseries. Pythium irregulare Buism. for example, has been noted to kill Pinus patula (Linde et al., 1994) and Picea abies (Kozlowski & Métraux, 1998), and Pseudotsuga menziesii (Weiland et al., 2013). Pythium ultimum var. ultimum and P. cinnamomi were also found virulent on Pinus sylvestris (Chavarriaga et al., 2007). However, pathogens are not the only challenge nurseries must contend with. Weed plants are a concern as they compete for water, nutrients, and sunlight, especially at the nursery stage, but also at each subsequent stage of the conifer tree production cycle. Growers have many options to control weeds. Non-chemical means include mowing, pulling weeds by hand, and even employing sheep to graze on the non-target crop (Saha et al., 2020). With over 100,000 people employed in the conifer tree industry within the United States alone (Saha et al., 2020), growers have incentive to reduce labor expenditures and chemical weed reduction is often chosen. In bare-root tree nurseries, preplant soil fumigation is a common method employed to reduce both weeds and pathogens in the soil (Shrestha et al., 2008; Weiland 47 et al., 2016). A popular choice among growers for preplant fumigation is methyl bromide with chloropicrin as it has proven effective at controlling pathogens as well as weed seed germination (Weiland et al., 2016). Methyl bromide is naturally produced in marine environments by many phytoplankton species (Sœmundsdóttir & Matrai, 1998) although it remains unclear what role it plays. It has been suggested that halogenated metabolites act as chemical defense or as antifouling agents (Paul & Pohnert, 2011). However, methyl bromide is also a powerful broad-spectrum pesticide produced commercially for use in food crops such as peppers, strawberries, grapes and structural fumigation applications for treating pests such as termites (Piccirillo & Piccirillo, 2010). Typical agricultural application of methyl bromide is injection into the soil followed by covering the ground with plastic for several days to seal in the fumigant (Piccirillo & Piccirillo, 2010). Since methyl bromide is a colorless and odorless gas, chloropicrin is often added to help applicators detect exposure since methyl bromide is neurotoxic in humans (Piccirillo & Piccirillo, 2010). Used as a chemical warfare agent in World War I (Sciuto & Kodavanti, 2015), chloropicrin also has biocidal and fungicidal applications (Wilhelm et al., 1996) similar to methyl bromide. Additionally, as with methyl bromide the exact mode of action is unclear (O’Malley, 2010; Sparks et al., 1997). In unison, these two fungicides are effective in controlling nematodes, fungi, oomycetes, and many more soil-borne pathogens though methyl bromide is being phased out and restricted due to its role in ozone depletion (Weiland et al., 2011; Zasada et al., 2010). Another unintended consequence of methyl bromide used is the impact it has on non- target organisms. Beneficial soil microbiota are known to significantly influence plant fitness and broad spectrum fumigants reduce this critical facet of soil health in our agricultural systems (Astudillo-García et al., 2019; Castellano-Hinojosa et al., 2022). Broad-spectrum fungicidal soil 48 fumigants have the potential to reduce and effectively reset the community structure leading to unique opportunities for studying priority effects. An ever broadening assemblage of studies provide strong evidence as to the importance of priority effects on microbiome assembly, diversity, host and ecosystem function (Debray et al., 2022). However, some edible ectomycorrhizal fungi cultivation systems aim to restrict mycorrhizal diversity. Early dominating establishment is critical for cultivating targeted ectomycorrhizal fungi, such as truffles, whereby seedlings are raised to be well-colonized by select Tuber species before planting out into prepared fields. These highly prized hypogeous fruiting bodies of Tuber spp. P. Micheli form in association with various hosts including Quercus L., Pinus L., Picea A. Dietr., and Pseudotsuga Carrière. It is estimated that over 500,000 trees inoculated with truffle spores are produced annually in nurseries where they are grown in individual pots for later sales and establishment by truffle plantation owners (Murat, 2014). Beyond marketing edible mycorrhizal fungi, many companies choose to focus on growth and health aspects of various crops in an ever growing industry. Research strongly implicates the importance of fungi in the rhizosphere of plants. However, despite this growing field of interest and an ever expanding commercial presence, the results have been largely lackluster as there are numerous reports of little to no growth or health benefits observed (Duell et al., 2022). There are a multitude of challenges these products must overcome. For example, one study found that Sphagnum peat moss, commonly used in container nurseries, contains many ectomycorrhizal species which are still viable and were able to colonize the roots of Pinus montezumae (Ángeles-Argáiz et al., 2016). However, most mycorrhizal products sold primarily contain arbuscular mycorrhizae; though some recent research suggests that the advertised arbuscular inocula is rarely accurate. One study found that the products they 49 sampled had unaccounted for taxa and were nearly all missing taxa claimed on the product label (Vahter et al., 2023). Two of these products contained none of the taxa listed on the packaging and five products only contained one of the listed inoculants (Vahter et al., 2023). This study was conducted to investigate whether introducing ectomycorrhizal inoculum increases tree growth and reduces disease incidence in three species of conifers at two bare-root nurseries. We utilized locally acquired ectomycorrhizal fungi, exotic Tuber species, and readily available commercial blends of mycorrhizal inoculum in this field experiment. Specifically, we wanted to investigate the following: can roots of Pseudotsuga menziesii, Picea abies, and Pinus sylvestris grown in methyl bromide fumigated soils be colonized by targeted ectomycorrhizal fungi. We also wanted to assess how well locally sourced, exotic (Tuber spp.), and commercial inoculum establishes and persists in methyl-bromide fumigated soils, and similarly how host species, inoculum treatment, and location impact fungal and oomycete diversity in the roots of conifer tree seedlings. Methods Two variations of an experimental design were carried out over two complete bare-root conifer nursery cropping cycles. They will be referred to in this manuscript based on the year of inoculation as either experiment 1 or experiment 2 for inoculation 2018 or inoculation 2019 respectively. Tables 2-4 provide the complete experimental design for both farms in both years. Site Selection The two bare-root conifer tree nurseries 20 miles apart in Michigan were selected for this field experiment. These sites share a temperate continental climate with cold winters. The average low-temperature in the coldest month is -7.2 C, and the average high-temperature in the warmest month is 26.4 C. There are an average of 100.8 rainfall days and 186 mm of 50 precipitation annually. Both nurseries utilize Tri-Brom 80-20 soil fumigant at 240lb/ac and are composed of the following active ingredients: 80% Methyl Bromide, 19.9% Chloropicrin. These two sites are referred to as Allegan and Gobles. Soil at Allegan is a Chelsea loamy fine sand with a pH of 7.0 and the soil at Gobles is Ottokee loamy fine sand with an average pH of 6.8. The pH of homogenized soil was collected from 5 points approximately 12.0 cm deep for each tree species at each farm. Each was prepared by mixing two parts soil into one part double distilled water by mass and letting it rest for 30 min. Readings were collected using a calibrated pH Meter (AB15 Accumet Basic, Fisher Scientific). Soil Preparation Both farms fumigated their fields early September in the year prior to planting. See Supplementary table 3.1 for the complete pesticide application schedule. For experiment 1, seeds were sown on May 15th, 2018 at Allegan and May 22nd, 2018 at Gobles. For experiment 2, seeds were sown on May 16th, 2019 at Allegan and June 5th, 2019 at Gobles. Prior to sowing, grower-cooperators prepared seedbeds using their standard practices (FIGURE 2.1A). Once seedbeds were formed (1.0 m wide), the cooperator sowed seeds at the following rates: Norway spruce 650 / m2 ; Scots pine 550 / m2 ; Douglas-fir 370 / m2 . Inoculum Preparation Application rates for all inoculum are listed in Table 2.2. Tuber indicum Cooke & Massee, Tuber aestivum Vittad., and Tuber borchii Vittad. ascocarps were weighed and surface sterilized using 3% hydrogen peroxide for 2 minutes and rinsed with double distilled water. Tuber indicum ascocarps were pulverized using a Waring commercial blender (Model 4324X) on the high setting for 15 minutes; T. borchii and Tuber aestivum for 10 minutes. Ascocarps, crushed ice, and double distilled water were added to the blender and after running for the 51 aforementioned times, a compound light microscope (Leica DM750) was used to verify that approximately 50% of ascospores were released from their asci. This spore suspension was brought to a concentration of 0.2g/mL using additional double distilled water and then stored at 4C overnight for application the following day. While in transport to the field sites, inocula remained in a cooler on ice until application to preserve spore viability. Laccaria bicolor (Maire) P.D. Orton, and Scleroderma citrinum Pers. basidiocarps were each also prepared the same as with Tuber, however blended for only 5 minutes. L. bicolor, and S. citrinum suspensions were each brought to 0.1g/mL. Field inoculation 2018 At both Allegan and Gobles, beds were formed in the morning just prior to inoculation. At Allegan the study was installed as a fully randomized block design (Table 3.3). However, Gobles did not have a randomized design to reduce treatment blending by farm equipment. Allegan spanned 85 treatment blocks and Gobles included 75 treatment blocks. Each treatment block was 1.0 m2 and a PVC frame was utilized to ensure even application over each block. Specific locations for each block were identified using an iron spike with an identification number such that locating exact locations for subsequent sampling during the fall of the following year could be made precisely where spore application was made. Spore suspensions were further diluted to facilitate even spraying using a 4.0 L portable pump sprayer (Chapin brand). Each inoculum was applied at three concentrations; T. aestivum, T. borchii, and T. indicum were applied at 0.2g/m2 , 2.0g/m2 , and 20g/m2 . Similarly, L. bicolor, and S. citrina were each applied at 0.1g/m2 , 1.0g/m2 , and 10g/m2 . Negative control blocks were sprayed with water from the same distilled water as was used to make the fungal spore slurries. Each treatment block was replicated five times at both farms. 52 At Allegan and Gobles, Picea abies (Norway spruce), Pinus sylvestris (Scots pine), and Pseudotsuga menziesii (Douglas-fir) were subjected to these inoculation treatments. Treatment blocks at Allegan were spread evenly across the total length of the bed for each tree species; P. abies 274.32 meters, P. sylvestris and P. menziesii each 122 meters. At Gobles, treatment blocks were grouped (supplementary table 4) to reduce treatment mixing effects. Spore slurries were applied at both sites between bed formation and seed sowing by spraying the slurry directly onto the soil surface. After seeds were sowed into the soil, a roller press finished the formation of the bed. Field inoculation 2019 The inoculation in 2019 was carried out in a similar manner as the 2018 inoculation with key differences being inoculum type and timing. Here, spore application was performed before bed formation such that spores could be rototilled into the soil. Allegan spanned 40 treatment blocks and Gobles included 20 treatment blocks. In 2019 T. borchii, T. indicum, and a commercial mycorrhizal fungi (MycoApply® Endo/Ecto, Mycorrhizal Applications, Grants Pass, Oregon USA) blend were used. The commercial product used is a mix of endomycorrhizal and ectomycorrhizal fungal spores homogenized in a fine powder designed for water suspension. According to the label the commercial inoculum includes spores of Glomus intraradices, G. mosseae, G. aggregatum, G. etunicatum as the included endomycorrhizal inoculants and Rhizopogon villosulus, R. luteolus, R. amylopogon, R. fulvigleba, Pisolithus tinctorius, Scleroderma cepa, and S. citrinum as the included ectomycorrhizal fungal inoculants. 53 Collection dates Trees inoculated in the Spring of 2018 were collected from Allegan and Gobles on the 9th and 13th of September 2019, respectively. Trees inoculated in the Spring of 2019 were collected from Allegan and Gobles on the 11th and 15th of June 2020, respectively. Collection Methodology At both sites for both years the collection methodology was the same. A point-intercept frame was utilized for each block such that trees closest to points on the frame were selected in a non-biased way. This frame was placed over the treatment block being sampled and the tree nearest to a point was carefully dug out of the soil using a hand trowel. Five trees from each block were collected and placed into a plastic bag which was stored on ice in a cooler while in the field. The samples were stored that day in a 4°C walk-in cooler until further processing the following day. Plant growth metrics Tree height was measured from the root collar to the apical tip using a ruler for all trees, both years. We measured root collar diameters of trees collected in 2020 from both sites using digital calipers. Overall tree health was assessed based on chlorosis and necrosis on a scale of 1- 4, where 1 indicated more than 75% necrosis/chlorosis; 2 indicates 50-75% necrosis/chlorosis. Each rating level represents a 25% rating, for example, a rating of 2 indicates 50-75 necrosis/chlorosis; 3 indicates 25 to 50 necrosis/chlorosis whereas a 4 indicates less than 25% necrosis/chlorosis. Root colonization levels were determined by analysis conducted under a compound light microscope (Leica Model S9i,1700 Leider Lane, Buffalo Grove, IL 60089 United States) (FIGURE 3.1C). 54 Plant Tissue Preparation After morphological analysis concluded, roots were separated from the above ground growth at the root collar. Root sections were then soaked briefly in a 0.01% solution of Tween20 then rinsed with tap water until any loose soils had been removed. Surface cleaned roots were then wrapped in paper towels and allowed to air dry overnight on. The following day, roots in paper towels were stored in a plastic bag with silica desiccant beads until further processing. Process controls consisting of the same paper towels were prepared with water and dried but without roots inside. DNA extraction Dried roots placed in paper towel bundles were rolled against a hard surface such that root tips were dislodged and freed from the larger root sections from which they arose. Loosened root pieces were then collected and placed into a 2.0 mL microcentrifuge tube with two stainless steel ball bearings. Each sample was ground up and homogenized utilizing the Qiagen Tissuelyser II bead mill for 30 RPS for one minute (QIAGEN Sciences Inc, 19300 Germantown Rd, Germantown, MD 20874). DNA was extracted from the plant tissue using the Mag-Bind Plant DNA Plus Kit (Omega Bio-tek, United States). Process controls were included with each extraction and also included the paper towel controls used in the drying process. Illumina MiSeq Sequencing and Library Preparation Amplicon libraries for MiSeq sequencing were produced based on the same three step PCR protocols used in (Benucci, et al., 2019; Lundberg et al., 2013). The ITS1F and ITS2 primer sets were used for the Fungal library and the ITS-6 (5’ GAAGGTGAAGTCGTAACAAGG 3’) and ITS-7 (5’ AGCGTTCTTCATCGATGTGC 3’) primer sets were used for the Oomycetes library. The products of the PCR were visualized utilizing a QIAxcel Advanced capillary 55 electrophoresis device (QIAGEN Sciences Inc, 19300 Germantown Rd, Germantown, MD 20874). PCR products from successful amplifications were normalized using the Invitrogen SequalPrep™ Normalization Plate (96) kit (Fisher Scientific International L.L.C., Wyman Street, Waltham, Massachusetts 02454-9046). Normalized amplicons were then pooled using the Amicon® Ultra 0.5 mL 50 K filters (EMD Millipore, Germany) then purified to remove primer dimers as well as remaining fragments using the Agencourt AMPure XP magnetic beads kit (Beckman Coulter, USA). The purified amplicons were then sequenced at the Research Technology Support Facility (RTSF) Genomics Core at Michigan State University utilizing the Illumina MiSeq V3 analyzer with the 600 cycles kit (Illumina Inc., 5200 Illumina Way, San Diego, CA). Bioinformatic and Statistical analyses Sequences were initially processed using the same pipeline as in (Benucci, et al., 2019b) up to the taxonomic assignments. Here, CONSTAX2 taxonomic classifier was utilized (Liber et al., 2021) as well as FUNGuild (Nguyen et al., 2016) for ecological classification. Operational taxonomic unit (OTU) tables and metadata files were imported to R Studio (build 372) running R version 4.1.2 (Bunn & Korpela, 2015). The scripts used in this pipeline are available at https://github.com/rennickb. 56 Results FIGURE 3.1 - FIELD SITES, ROOT INSPECTION, AND COLONIZED ROOT TIP. A) Gobles seed bed formation. B) Allegan conifer seedlings before lifting. C) Norway spruce under compound light microscope with root system suspended in water. D) Extramatrical 57 FIGURE 3.1 (cont’d) - hyphae on Douglas-fir typical of Basidiomycota ectomycorrhizae. E) Example of Tuber borchii mycorrhizae on Pinus. F) Ectomycorrhizal mantle on Pinus roots. Tree growth Inoculation did not affect (p>0.05) tree height (FIGURE 3.3), biomass (FIGURE 3.4), or root collar diameter (FIGURE 3.5 and 3.6) regardless of tree species or field location. With an average health rating of 3.9, overall tree health was high and similarly, there was very little to no disease noted for any inoculated or uninoculated blocks. Visual inspection of the roots for exotic mycorrhizae did not reveal any roots colonized by Tuber species (FIGURE 3.1). Irrespective to the field location, tree species, or inocula there was a great diversity of mycorrhizal fungi on the roots of conifer trees in soils previously fumigated with Tri-Brom 80-20 (Fig 3.1). High-throughput sequencing results The total reads after filtering for quality were 20,975,165 for the fungal library and 19,839,935 reads for the oomycete library. Across 314 samples, the average read depth in the fungal library was 66,800 and 63,185 in the oomycete library. There were 1,680 OTU’s in the fungal library and 117 OTU’s in the oomycete library. Fungal Library Community Composition The fungal library was dominated by two taxa, Sphaerosporella (Svrček) Svrček & Kubička (33.4% relative abundance across dataset) and Wilcoxina Chin S. Yang & Korf. (32% relative abundance across dataset) Other notable taxa included Suillus Gray, Peziza Dill. ex Fr., Hebeloma (Fr.) P. Kumm., and Geopora Harkn. Suillus spp. (14.3% relative abundance) are well represented across the roots of P. sylvestris at Gobles during the first experiment. Successful Tuber borchii inoculation seems to have occurred on the roots of Douglas-fir trees at Gobles within experiment 1 (12.7% relative abundance), though read counts were low, T. borchii was 58 detected on Norway spruce and Scots pine here as well. Interestingly, Laccaria Berk. & Broome was found in the control blocks placed with Douglas-fir at Allegan, experiment 1 (1.2% relative abundance) and 2 (1.0% relative abundance), however, it was also found on Scots pine at Gobles in low abundances as well. Scleroderma Pers.was not found in any treatment, tree, location, or experiment. Pathogenic taxa varied greatly between field locations and the community structure (FIGURE 3.7). With experiment 1, the pathogen fungal library at Allegan was dominated by Fusarium, Trichoderma Pers., Diaporthe Fuckel, Clonostachys Corda, and a Ceratobasidiaceae G.W. Martin (FIGURE 3.7). However, Paraphoma Morgan-Jones & J.F. White, Phoma Sacc., Nectriaceae Tul. & C. Tul., Diplodia Fr., and Rhizoctonia DC. had the most reads from the Gobles roots during the first experiment. However, experiment 2 resulted in higher Macrophomina Petr. and Paraphoma reads from Allegan and Nectria (Fr.) Fr., Clonostachys, and similarly, Paraphoma at Gobles (FIGURE 3.7). Oomycete Library Community Composition Many other important pathogenic taxa in the Oomycetes lineage were also present at both locations. One Pythium OTU dominated the oomycete library at Allegan during the first experiment (FIGURE 3.3). As a genus, Pythium comprises nearly all reads in the dataset (92.6% relative abundance). Notably tied to damping-off disease in nursery seedlings, P. dissotocum (9.1% relative abundance), P. irregulare (11.1% relative abundance), P. rostratifingens (11.2% relative abundance), and P. ultimum var ultimum (5% relative abundance) were each widespread across the dataset. The Phytophthora genus only represents 1.3% relative abundance in the dataset. 59 FIGURE 3.1 – MYCORRHIZAL FUNGAL TAXA STACKED BAR PLOT. Mycorrhizal fungi in roots of Picea abies (L.) H.Karst. (Norway Spruce), Pinus sylvestris L. (Scots pine), and Pseudotsuga menziesii (Mirb.) Franco (Douglas-fir) after one year of growth in soils inoculated with either a commercial mycorrhizal blend (MycoApply®), local inocula (Laccaria Berk. & Broome and Scleroderma Pers.), Tuber P. Micheli inocula (T. aestivum Vittad., T. borchii Vittad., and T. indicum Cooke & Massee), or no inocula (control). Experiment 1 at Gobles has the strongest Tuber borchii reads at 12.7% relative abundance within Tuber inocula blocks in P. menziesii across both experiments. Suillus species (14.3% relative abundance) are well represented across the roots of P. sylvestris at Gobles during the first experiment. Nearly all tree 60 FIGURE 3.1 (cont’d) – species were largely dominated by Wilcoxina Chin S. Yang & Korf, Sphaerosporella (Svrček) Svrček & Kubička, and unclassified Pyronemataceae taxa. FIGURE 3.2 – OOMYCETE TAXA STACKED BAR PLOTS. Oomycete taxa in roots of Picea abies (L.) H.Karst., Pinus sylvestris L., and Pseudotsuga menziesii (Mirb.) Franco after one year of growth in soils inoculated with either a commercial mycorrhizal blend (MycoApply®), local inocula (Laccaria Berk. & Broome and Scleroderma Pers.), Tuber P. Micheli inocula (T. aestivum Vittad., T. borchii Vittad., and T. indicum Cooke & Massee), or no inocula (control). Pythium dissotocum Drechsler, P. irregulare Buisman, P. rostratifingens De Cock & Lévesque, and P. ultimum var ultimum Trow are notable pathogens linked to damping- off disease and comprise a significant portion of all reads in this dataset. 61 FIGURE 3.3 – INFLUENCE OF INOCULUM ON HEIGHT. Picea abies (L.) H.Karst., Pinus sylvestris L., and Pseudotsuga menziesii (Mirb.) Franco height after one year of growth in soils inoculated with either a commercial mycorrhizal blend (MycoApply®, local inocula (Laccaria Berk. & Broome and Scleroderma Pers.), Tuber P. Micheli inocula (T. aestivum Vittad., T. borchii Vittad., and T. indicum Cooke & Massee), or no inocula (control). There is no significant difference in height among trees grown in control blocks (no inoculation) and those grown with any of the inocula. 62 FIGURE 3.4 – INFLUENCE OF INOCULUM ON BIOMASS. Picea abies (L.) H.Karst., Pinus sylvestris L., and Pseudotsuga menziesii (Mirb.) Franco biomass after one year of growth in soils inoculated with either a commercial mycorrhizal blend (MycoApply®, local inocula (Laccaria Berk. & Broome and Scleroderma Pers.), Tuber P. Micheli inocula (T. aestivum Vittad., T. borchii Vittad., and T. indicum Cooke & Massee), or no inocula (control). There is no significant difference in biomass among trees grown in control blocks (no inoculation) and those grown with any of the inocula. 63 FIGURE 3.5 – INFLUENCE OF INOCULUM ON ROOT COLLAR DIAMETER. Picea abies (L.) H.Karst., Pinus sylvestris L., and Pseudotsuga menziesii (Mirb.) Franco root collar 64 FIGURE 3.5 (cont’d) – diameter after one year of growth in soils inoculated with either a commercial mycorrhizal blend (MycoApply®), Tuber P. Micheli inocula (T. borchii Vittad., and T. indicum Cooke & Massee), or no inocula (control). There is no significant difference in root collar diameter among trees grown in control blocks (no inoculation) and those grown with any of the inocula. 65 FIGURE 3.6 – ROOT COLLAR DIAMETER AND TREE HEIGHT SAMPLE DISTRIBUTION. Distribution of root collar diameters of Picea abies (L.) H.Karst., Pinus sylvestris L., and Pseudotsuga menziesii (Mirb.) Franco grown in soils inoculated with either a commercial mycorrhizal blend (MycoApply®), Tuber P. Micheli inocula (T. borchii Vittad., and T. indicum Cooke & Massee), or no inocula (control), or in soils that received no inoculation. 66 FIGURE 3.7 – PATHOGENIC FUNGAL TAXA STACKED BAR PLOT. Pathogenic fungi in roots of Picea abies (L.) H.Karst., Pinus sylvestris L., and Pseudotsuga menziesii (Mirb.) Franco after one year of growth in soils inoculated with either a commercial mycorrhizal blend (MycoApply®), local inocula (Laccaria Berk. & Broome and Scleroderma Pers.), Tuber P. Micheli inocula (T. aestivum Vittad., T. borchii Vittad., and T. indicum Cooke & Massee), or no inocula (control). Experiment 1 Allegan is dominated by Fusarium Link, Trichoderma Pers., Diaporthe Fr., Clonostachys Corda, and a Ceratobasidiaceae G.W. Martin operational taxonomic unit, whereas Gobles was dominated by Paraphoma Morgan-Jones & J.F. White, Phoma Sacc., Nectriaceae (Fr.) Fr., Diplodia Fr., and Rhizoctonia DC. Experiment 2 Allegan trees were dominated by Trichoderma Pers., Macrophomina Petr., and Paraphoma Morgan-Jones & J.F. 67 FIGURE 3.7 (cont’d) – White, whereas Gobles was dominated by Nectria (Fr.) Fr., Clonostachys Corda, and similarly, Paraphoma. FIGURE 3.8 – FUNGAL AND OOMYCETE RAREFACTION CURVES. 68 Table 3.1 – Pesticide application rates. 69 Table 3.2 – Inoculum application rates. 70 Table 3.3 – Experiment 1 inoculation block design. 71 Table 3.4 – Experiment 2 inoculation block design. 72 Discussion This study was designed to assess whether mycorrhizal inoculum is effective or necessary in bare-root conifer tree nurseries that manage seedling beds with methyl bromide and chloropicrin. Our field experiments were carried out during two growing seasons and indicate that in-field ectomycorrhizal inoculum supplements are not effective or necessary. Despite the use of local, exotic, or commercial inocula, there were no significant increases in the growth or health of trees observed in this study. While there may be cropping systems that benefit from these sorts of inputs, the robustness of the mycorrhizae either persisting through fumigation or blowing in from nearby tree stands may be all that is needed to meet the requirements of these tree species. However, it is likely that wind dispersed spores are the primary source as one study found Glomus intraradices spores were no longer viable after fumigation by methyl bromide (Bendavid-Val et al., 1997). Yet, more work is needed to determine the effects of methyl bromide on spores of ectomycorrhizal fungal species. Fungal community In fact, native infield inoculum is sufficiently high enough that it displaces most of the ectomycorrhizal spore inoculum provided in our treatments. Certain ectomycorrhizal groups including Suillus were particularly pervasive (Fig 3.1). Suillus is strongly associated with Pseudotsuga, Pinus, and Abies (Smith & Thiers, 1964) it is therefore not surprising that the locations this study took place within would be inundated with spores blowing in from nearby conifer tree stands. Additionally, there is some evidence that suggests certain ectomycorrhizal species such as Laccaria laccata (Scop.) Cooke produce methyl bromide raising questions about natural tolerance to this compound when used as a fumigant (Redeker et al., 2004). Potential production and tolerances to methyl bromide by ectomycorrhizal fungi have not been widely 73 studied and may yield fruitful insights given future research. The wide diversity of ectomycorrhizal species found on the roots within these nurseries also suggests that interest in applying inoculants may not be necessary or beneficial. There were differences in fungal community composition among treatments, however, these differences were not reflected in growth metrics or overall tree health, hinting at the importance of robust and diverse fungal communities. Oomycete community The other community investigated in this study, oomycetes, are among the most concerning pathogen groups within our dataset. Unlike mushrooms, oomycetes produce motile spores that travel in water and not in the air. Oomycete spores spread largely through irrigation and splashing. Methyl bromide application typically reaches 20 to 25 cm in depth, however, soil composition, tilth, moisture, and temperature each factor into the efficacy and penetration of this fumigant (Dunlap, 2009). Some Pythium species, such P. splendens Hans Braun are most abundant at soil depth of 15 to 30 cm, others have been recorded at depths of up to 355 cm (Plaats-Niterink, 1981). Our analysis of the oomycetes found Pythium ultimum var. ultimum and Pythium irregulare, two of many concerning pathogens amongst Pinaceae (Kozlowski & Métraux, 1998; Chavarriaga et al., 2007; Weiland et al., 2013). While methyl bromide may be effective at controlling Pythium populations near the surface at the time of application, deeper populations combined with irrigation or other mass water flow events can easily return species to the fumigated zone. Persistence of introduced taxa Our work here demonstrates that the fungal community dynamics of these fumigated bare-root nurseries are not conducive to addition of inoculants at the time of planting. By using 74 exotic truffle inocula easily discernible from local mycorrhizal species, we aimed to have both an easily identifiable mycorrhizal morphology for visual inspection and a unique genetic sequence for identification within the MiSeq library. Though the visual inspection did not reveal any Tuber mycorrhizae, we did uncover Tuber borchii in the MiSeq library. While this does indicate the presence of the exotic inocula both establishing and persisting on trees grown in methyl bromide fumigated soils, the relative abundance was rather low. One of the goals of this work was to investigate priority effects in a real-world dynamic system, and the persistence Tuber borchii indicates that introduced fungi can persist even if relatively rare. This is not very surprising given what is known about how well fungi have adapted spore dispersal. There are many challenges to calculating spore dispersal among mushrooms (Dam, 2013), however, there is sufficient evidence that conducive conditions may allow spores to travel hundreds of kilometers in the wind (Viljanen-Rollinson et al., 2007). Open-field bare-root nurseries surrounded by mature trees of the same species being cultivated such as those utilized in this study are no doubt showered in an abundant siege of mutualistic fungal spores, but with the inflow of beneficial species, so too come pathogens even trans-continentally (Brown & Hovmøller, 2002). Incomplete picture Biases inherent to amplicon sequencing mean one must use caution when relating read counts to actual environmental abundance or diversity. The metabarcoding in this study was applied to surface-cleaned roots from living trees, however, this does not guarantee the amplicons belong to active living organisms. As an additional limitation, we know that every locus, primer set, and reference database have their own intrinsic biases as well so it was no surprise that the detection of Tuber aestivum, for example, did not occur as the primer set used 75 does not select for this species. Similarly, as made evident by the dominating Pythium OTU not identified to species level (FIGURE 3.3), there are yet many challenges faced when metabarcoding Oomycetes. A known limitation of the ITS6 and ITS7 primers often used in oomycete community amplification (Cooke et al., 2000), are their propensity to bind and amplify fungal and plant sequences (Coince et al., 2013). Additionally, these primers have been shown to preferentially amplify Pythium over Aphanomyces de Bary ( Taheri et al., 2017) leading to unbalanced abundancies. Summary In summary, through looking at two bare-root nurseries over two cropping cycles and three conifer tree species (Pseudotsuga menziesii, Picea abies, and Pinus sylvestris), and using local (Laccaria bicolor or Scleroderma citrinum), exotic (Tuber aestivum, Tuber borchii, or Tuber indicum), or a commercial blend (MycoApply®) of mycorrhizal inoculants, we did not observe statistically significant reduction in disease or increase in tree height, root collar diameter, or biomass. We did not see the exotic mycorrhizae during a visual inspection but did find T. borchii in low abundance on the roots of P. menziesii during the first experiment. We also observed over 12 species of Pythium and four species of Phytophthora in the oomycete MiSeq library dataset we generated. Conclusions Based on the lack of disease reduction, the lack of growth increases, and the diversity of ectomycorrhizal fungi on the roots of P. menziesii, P. abies, and P. sylvestris in these bare-root nurseries, we cannot recommend the use of mycorrhizae inoculants. Should bare-root nurseries decide to continue using methyl bromide with chloropicrin, spores blowing in from fungi 76 growing in nearby tree stands will likely be sufficient to provide the health and growth benefit s that mycorrhizal fungi like Suillus, Sphaerosporella, and Wilcoxina are known to provide. 77 Chapter 4 Tuber rugosum, a new species from Northeastern North America: Slug mycophagy aides in electron microscopy of ascospores Source: This chapter has been published in Mycologia: Rennick, B., Benucci, G. M. N., Du, Z.-Y., Healy, R., & Bonito, G. (2023). Tuber rugosum, a new species from northeastern North America: Slug mycophagy aides in electron microscopy of ascospores. Mycologia, 1–17. https://doi.org/10.1080/00275514.2023.2184983 78 Abstract Species in the genus Tuber are ascomycetous fungi that produce hypogeous fruiting bodies commonly called truffles. These fungi are ecologically relevant owing to the ectomycorrhizal symbiosis they establish with plants. One of the most speciose lineages within Tuber is the Rufum clade, which is widely distributed throughout Asia, Europe, and North America and is estimated to include more than 43 species. Most species in this clade have spiny spores, and many still have not been formally described. Here, we describe Tuber rugosum based on multigene phylogenetic analysis and its unique morphological characters. Tuber rugosum (previously designated in literature as Tuber sp. 69) has been collected throughout the Midwest, USA, and Quebec, Canada, and is an ectomycorrhizal symbiont of Quercus trees, as confirmed through morphological and molecular analyses of root tips presented here. We also present a novel method for preparing Tuber ascospores for scanning electron microscope imaging that includes feeding, digestion, and spore excretion by the slug Arion subfuscus. Following this method, spores become free from ascus and other mycelial debris that could obscure morphological traits during their passage through the snail gut while maintaining ornamentation. Finally, we report the fatty acid analysis, a fungicolous species association, and we provide an updated taxonomic key of the Rufum clade. Introduction Although first described in 1780, truffles in the genus Tuber have a much longer history of being sought after due to their culinary value (Wang and Marcone, 2011), which can be attributed to their unique aroma (Martin et al., 2010). The white truffle T. magnatum Picco and the black truffle T. melanosporum Vittad, for example, are amongst the most well-known and highly prized fungal species with unique aromas (Pelusio et al., 1995). Beyond their aroma, 79 truffles are rich in carbohydrates, proteins, and unsaturated fatty acids (Bouatia et al., 2018; Yan et al., 2017), but it is still unknown how variable these traits are between Tuber species. In addition, the ecology and microbiology few Tuber species has been extensively studied (Splivallo et al., 2011), often linking fungivory by animals to the aromatic lure produced by a mature ascocarp (Hochberg et al., 2003; Maser et al., 2008). Successful truffle spore dispersal relies on mycophagous animals detecting, consuming, and defecating mature sporocarps, as hypogeous fungi are not able to actively discharge their spores. Numerous studies have highlighted Tuber spores found in animal scat, including that of pika (Cázares and Trappe, 1994), northern flying squirrels (Gabel et al., 2010), crested porcupines (Ori et al., 2018), and wild boar (Piattoni et al., 2013). Passage of spores through animal digestive tracts such as the crested porcupine removes asci, may lead to some degradation of the ornamentation, and often will promote spore germination (Ori et al., 2018). Observations made while collecting specimens for this study indicated slugs, which are known to be mycophagous (Beyer and Saari, 1978; McGraw et al., 2002), regularly consume truffles and other fungi that grow beneath the leaf litter of the forest floor. These observations led to the question of whether slugs could be used as an alternative to chemical preparations (e.g., Puliga et al., 2020) to obtain clean ascospores for unobstructed and improved scanning electron imaging. Further field observations of truffles as they matured in situ led to observations of fungal infections. In 2017, Leonardi et al. reported on fungi living within eight species of Tuber where they found 58.6% were infected by Clonostachys rosea (Link) Schroers, Samuels, Seifert & W. Gams (= Bionectria ochroleuca (Schwein.) Schroers & Samuels) (Leonardi et al., 2018). Other studies have indicated that yeasts including Candida saitoana Nakase & M. Suzuki, Rhodotorula mucilaginosa (A. Jörg.) F.C. Harrison, and Trichosporon moniliiforme E. Guého & M.T. Sm. 80 isolated from Tuber melanosporum and Tuber magnatum ascomata produce, and may contribute to, the characteristic aroma profile of the truffles in which they are found (Buzzini et al., 2005). Mycelial fungi such as Trichopezizella nidulus (J.C. Schmidt & Kunze) Raitv., Absidia cylindrospora Hagem, and Peniophora cinerea (Pers.) Cooke are also known to be associated with truffle fruiting bodies (Pacioni et al., 2007). Additionally, Aspergillus, Cladosporium, Fusarium, Penicillium, and Trichoderma have been isolated from other truffle taxa, such as Tuber aestivum Vittad. and Tuber melanosporum ascomata (Rivera et al., 2010). Some mycoparasites are known to produce mycotoxins; thus, precautionary care should be taken to avoid consuming parasitized truffles. It has been suggested that truffles exposed on the surface of the soil are more prone to disease, although moisture levels in the environment likely play an equally important role in completing this disease triangle (Eslick, 2012). In concurrence with Eslick (2012), ascomata of the species that we describe here, T. rugosum, sp. nov., observed with disease was subhypogeous. The genus Tuber contains over 200 species, with most species diversity residing within the Rufum, Puberulum, and Maculatum clades (Bonito et al., 2010; Healy et al., 2016). Many species within these clades have yet to be formally described (Bonito et al., 2010; Healy et al., 2016). Truffles in the Rufum clade can be distinguished from those in other lineages by their smooth to slightly verrucose pale to reddish-colored peridium, stemmed ascus, and the often spiny or spinose-reticulate ornamentation of their ascospores (Healy et al., 2016). Another unique facet of the Rufum clade is the absence of cystidia on the mycorrhizal mantle they form (Healy et al., 2016). Spore ornamentation, size, shape, and dimension remain the cornerstone morphological characteristics used to describe Tuber species. 81 From 2009 through 2021, we collected truffles with morphological characteristics of those in the Rufum clade. An internal transcribed spacer (ITS) meta-analysis of Tuber has provided a framework by which many new species have been described (Bonito et al., 2010). Sequences of the ITS region from our specimens matched with the sequence Bonito et al. (2010) designated in the literature as Tuber sp. 69 (GenBank HM485428), which we formally describe here as Tuber rugosum, sp. nov. In support of this new species to science, we provide (i) multigene phylogenies based on the ITS, the elongation factor 1α (EF1α), and the second -largest subunit of RNA polymerase II (RPB2) genes; (ii) a morphological comparison of its peridium, gleba, and spores; and (iii) a characterization of its fatty acid profile. Further, we describe an improved method for preparing spores for scanning electron microscopy (SEM) study, we identify a fungicolous species found on T. rugosum, sp. nov., and present a dichotomous key for the Rufum clade. Materials and Methods Collection and isolation Truffles were collected with the aid of a hand-held four-pronged garden cultivator to remove leaf litter and explore within the upper 10 cm of forest soils. Photographs and field notes, including date, location, habitat, and fresh attributes, were made for each specimen. Specimens were stored at 4 C for a maximum of 24 h prior to morphological observations and pure culture isolation. Using forceps and sterile technique, small pieces of freshly exposed internal gleba hyphae from younger specimen were sampled and submerged into an agar medium composed of 8.0 g/L agar, 5.0 g/L potato dextrose broth, 1.5 g/L malt extract, 5.0 mL/L glycerol, and 0.82 g/L calcium nitrate. Prior to autoclaving, the pH was adjusted to 7.5 with 5.0 M sodium hydroxide. Once the postautoclave temperature fell below 50 C, 1.0 mL/L biotin (0.5 g/L stock), 1.0 mL/L 82 chloramphenicol (60.0 mg/mL stock), and 1.0 mL/L ampicillin (50.0 mg/mL stock) were added. After initial growth on antibiotic-containing medium, a subculture was made on the same medium lacking antibiotics. These cultures were incubated at room temperature (20–22 C). Morphological analyses Analysis of truffle micromorphological characters was conducted under a compound light microscope (Leica model DM750; Buffalo Grove, Illinois). Ascospores were collected by scraping a razor blade across the gleba and mounting the fungal tissue collected on the blade on a microscope slide with 3% KOH. In total, 85 spores from 33 asci were measured and imaged at 400× magnification against the long and short axes, excluding ornamentation (Leica Application Suite 4.0). Length, width, and Q (length:width) measurements of the spores were then calculated, as these metrics have been informative in distinguishing species of Oregon white truffles (Bonito et al., 2010). One immature Tuber sp. 69 (RH999) ascocarp used for recording developmental characters was sectioned in quarters and fixed for 2 h (4 C) in 2% glutaraldehyde + 2% paraformaldehyde in 0.1 M sodium cacodylate buffer (pH 7.2); rinsed three times in 0.1 M sodium cacodylate buffer for 20 min each; postfixed for 1 h (4 C) in 1% osmium tetroxide in 0.1 M sodium cacodylate buffer; rinsed in fresh buffer followed by three changes of deionized water for 10 min each; dehyd rated in a graded ethanol series (25%, 50%, 75%, 95%, and 100%, 3× for 1 h each); infiltrated in Spurr’s resin (Spurr, 1969) and embedded in an aluminum dish; and polymerized for 2 days at 74 C. Sections 2 µm thick were cut with a glass knife and placed on a drop of water on a clean glass microscope slide on a warming tray. After drying, sections were stained with 0.5% toluidine blue O and preserved with a drop of per mount and a drop of xylene and a cover glass was placed on 83 top. Images were digitally captured on a Nikon Optiphot compound microscope (Tokyo, Japan) mounted with a QImaging MicroPublisher 3.3 RTV camera (British Columbia, Canada). Hollowed out ascocarps in close proximity to abundant slug populations of the dusky Arion slug, Arion subfuscus, were observed. To test whether the slugs would consume truffles, we placed a single slug in a plastic container with a fresh truffle sporocarp for two sessions each lasting 4 h prior to observation. Frass contents collected from slugs that had consumed truffles were observed with a compound light microscope (Leica model DM750, Buffalo Grove, Illinois) to assess ascospore morphology. To prepare ascospores for scanning electron microscopy (SEM), slugs were maintained in a plastic box for 24 h and frass was observed after the complete T. rugosum, sp. nov., ascocarp had been consumed (SUPPLEMENTARY FIG. 1). The Arion frass was collected and visualized under a compound microscope. The ascospore-containing frass was dried at room temperature and then rinsed with phosphate-buffered saline (PBS). Ascospores not subjected to slug digestion were collected by scraping a scalpel blade across dried and rehydrated gleba and rinsed with PBS. Both sets of samples were fixed in a 4% (v/v) glutaraldehyde solution, dried with a critical point dryer (Balzers model 010; Balzers Union), and then mounted on aluminum stubs using high-vacuum carbon tabs (SPI Supplies, West Chester, PA). After the samples were coated with osmium using NEOC-AT osmium coater (Meiwafosis, Osaka, Japa), the samples were observed using a JSM-7500 F scanning electron microscope (Japan Electron Optics Laboratories (JEOL) USA, Peabody, Massachusetts). After primary character data collection commenced, specimens were cut into sections and dried with activated silica beads. Curated holotype and paratype collections have been deposited in the Michigan State University (MSU) Herbarium, with the MSU collection accession numbers MSC408482–MSC408486. These data have also been deposited into Mycobank MB838884. 84 Fungicolous species isolation Tissue supporting orange conidia growing from infected specimens of T. rugosum, sp. nov., was photographed with a Canon EOS Rebel T6 camera (Canon Inc., Tokyo, Japan) with the Laowa 24 mm f/14 2× Macro Probe lens (Venus Optics, Hefei, China) in the field prior to further processing. Photographs were taken with a shallow depth of field and were imported, aligned, and blended using Adobe Photoshop (Adobe Inc, 2019b) for FIG. 3A. The fungal growth supporting the orange conidia was then placed in malt extract agar (MEA) medium composed of 10.0 g/L agar, 10.0 g/L malt extract, and 1.0 g/L yeast extract with 1.0 mL/L chloramphenicol (60.0 mg/mL stock), 1.0 mL/L streptomycin (100.0 mg/mL stock), and rifampicin (50.0 mg/mL stock). This isolate (BR428b) was incubated at room temperatures (20– 22 C) and maintained on MEA with no antibiotics. Confocal microscopy Confocal microscopy was performed to visualize lipid droplets within the ascocarp. Samples were sliced with a surgical scalpel and stained with 10.0 μg mL−1 BODIPY 493/503 (Thermo Fisher Scientific, Pittsburgh, USA) in a phosphate-buffered saline (PBS) buffer for 2 days at 23 C. After two washes with a PBS buffer, the samples were then observed using an Olympus FV10i microscope (Olympus Scientific Solutions Americas, Waltham, Massachusetts). An argon (488 nm) laser was used for BODIPY (emission: 510–530 nm). Lipid extraction and analysis Mycelium was incubated on the agar medium described above until 40.0 mm (50% colonization of Petri dish) of growth from the inoculation point was reached. The total lipid fraction was extracted from the mycelium by placing methanol-chloroform-88% formic acid (1:2:0.1 by volume) in glass tubes, followed by a wash with half volume of 1.0 M KCl and 0.2 M 85 H3PO4. After phase separation by centrifugation (2000 × g for 3 min), total lipids were collected to prepare fatty acid methyl esters (FAMEs) with 1.0 M methanolic HCl at 80 C for 25 min. FAMEs were then extracted with hexane and analyzed by gas chromatography and flame ionization detection (Agilent, CA, USA). Molecular analyses DNA was extracted from all specimens and isolated using a rapid alkaline extraction method (Liber et al., 2022), as previously described. Ascocarp DNA was extracted by removing a small amount of the peridium and placing a 1.0-mm2 piece of sterile gleba into 40.0 μL extraction solution (ES). Colonized root tips were imaged, rinsed with deionized (DI) H2O, and placed into 20 μL ES and crushed using a pipette tip. Samples were then placed into a thermocycler set to 95 C for 10 min to lyse the cells. Following lysis, bovine serum albumin (BSA) was added at a rate of 3 times the volume of ES to help neutralize and suspend the DNA extraction. One microliter of the extracted DNA was used as template for subsequent polymerase chain reaction (PCR) amplification reactions. Fungal rDNA was amplified with universal fungal primers ITS1F and LR3 (TABLE 1). Tuber-specific primers were used to amplify protein-coding genes, including the second-largest subunit of RNA polymerase II (RPB2_Tuber_f, RPB2_Tuber_r) and elongation factor 1α (EF1α_Tuber_f, EF1α_Tuber_r) (TABLE 1) following methods of Bonito et al. (Citation2010, 2013). 86 Table 4.1 - List of primers and sequences used in this study for phylogenetic analyses. Primer Name Sequence (5′ to 3′) First Reported ITS1-F CTTGGTCATTTAGAGGAAGTAA Gardes & Bruns, 1993 LR3 CCGTGTTTCAAGACGGG Vilgalys & Hester, 1990 EF1α Tuber_f AGCGTGAGCGTGGTATCAC Bonito et al., 2013 EF1α Tuber_r GAGACGTTCTTGACGTTGAAG Bonito et al., 2013 RPB2 Tuber_f YAAYCTGACYTTRGCYGTYAA Bonito et al., 2013 RPB2 Tuber_r CRGTTTCCTGYTCAATCTCA Bonito et al., 2013 Amplicon products were Sanger sequenced bidirectionally at the Research Technology Support Facility (RTSF) Genomics Core at Michigan State University on the Applied Biosystems 3730XL capillary sequencer (Waltham, Massachusetts). Sequences were trimmed with SnapGene 4.3.7 to remove low-quality regions (GSL Biotech, Chicago, Illinois). Sequences were then compared with others in the National Center for Biotechnology Information (NCBI) database with the BLASTn algorithm to verify that they were Tuber and to identify other entries of this taxon in the database. Phylogenetic analyses Sequence alignments of taxa in the Rufum clade were made with the MUSCLE alignment algorithm (Edgar, 2004) within Mesquite (Maddison and Maddison, 2019). Sequence ends and highly ambiguous regions of ITS1 were excluded to eliminate ambiguous regions in the alignment. Aligned sequences were used to infer the phylogeny with maximum likelihood (ML) and Bayesian inference (BI). All ML searches were generated with Randomized Axelerated Maximum Likelihood (RAxML), and 1000 bootstrap replicates were carried out with the GTRGAMMA nucleotide substitution model on the CIPRES Science Gateway (Miller et al., 2010; Stamatakis, 2014). All BI searches were generated utilizing MrBayes on the CIPRES Science Gateway (Huelsenbeck and Ronquis, 2001; Ronquist and Huelsenbeck, 2003). The Markov chain Monte Carlo (MCMC) ran for 40 000 000 generations with the Metropolis- 87 coupled Markov Chain Monte Carlo (MCMCMC) set to run four chains in parallel, sampled every 1000 cycles, and had a burn-in rate of 25% for each BI search (Geyer, 1991). The model for among-site rate variation was set to INVGAMMA (inverse gamma distribution). Character sets for the ITS BI search were based on an alignment made to an annotated Tuber brumale Vittad. (GenBank AF106880) sequence extending from the 18S ribosomal RNA gene to the 28S ribosomal gene. The quality of the BI search was verified using MCMC files viewed with Tracer 1.7 to ensure parallel runs converge and to quantify the effective sample size (Rambaut et al., 2018). Visualization of the phylogenetic trees was performed using FigTree 1.4.4 (FigTree, 2018) updated to reflect the ML bootstrap support value and BI posterior probability within Adobe Illustrator (Adobe Inc, 2019a). Dichotomous key A dichotomous key of described species in the Rufum clade was generated based on available species descriptions used in TABLE 3 (Butters, 1903; Cao et al., 2011; Vittadini, 1831; Chen et al., 2005; Deng et al., 2009; Eberhart et al., 2020; Elliott et al., 2016; Lancellotti et al., 2016; Fan et al., 2012, 2013; Frank et al., 2006; Granetti et al., 1988; Grunow and Rabenhorst, 1884; Harkness, 1899; Hu and Wang, 2005; Leonardi et al., 2019; Suwannarach et al., 2016; Trappe et al., 1996; Uecker and Burdsall, 1977; Wang, 1988; Yan et al., 2018). 88 Results FIGURE 4.1 - TUBER RUGOSUM GROWTH IN VITRO. Growth shown on the pH 7.0 adjusted medium containing 1.0 mL/L biotin as described in Materials and Methods. A. A typical right-angled hyphal branch with simple septa. B. A hyphal coil that is seen regularly on surface growth in cultures as they age. Bars = 20.0 μm. 89 FIGURE 4.2 - TUBER RUGOSUM DESCRIPTION PLATE. A. Tuber rugosum (BR64, holotype) ascocarp showing both the rugose peridium and exposed sterile gleba. B. A cross- 90 FIGURE 4.2 (cont’d) – section of T. rugosum (BR64) showing the gleba and its marbling. C. A cross-section (stained with toluidine blue O) of T. rugosum (RH999) showing the distinctive long, narrow cells of the immature peridium. D. Hymenium (stained with toluidine blue O) of an immature specimen showing the developing asci and paraphyses. E. Ascii of T. rugosum (BR64) containing 1, 2, 3, 4, 5, and 7 ascospores. F. Mantle of T. rugosum (GenBank MW579340) on the root tip of a red oak (Quercus rubra). G. SEM image of a T. rugosum ascospore (BR64) showing the echinate surface. H. SEM image showing details of the uncinulate spines on the ascospore of T. rugosum (BR64, holotype; GenBank MW579336). Bars: A, B = 6.0 mm; C, F = 50.0 μm; D = 25.0 μm; E = 30.0 μm; G = 10.0 μm; H = 2.0 μm. Scanning Electron Microscopy. Spores that had passed through the digestive tract of the Dusky slug (Arion sp.) were free of nearly all of the asci remnants, leaving relatively clean and intact ascospores for imaging. SUPPLEMENTARY FIG. 1 shows clean, ascus-free T. rugosum, sp. nov., spores after passage through the slug digestive tract. As seen in FIG. 5, the delicate uncinulate spines are well preserved through this digestive process, which is an improved process for obtaining high- quality SEM opportunities. 91 FIGURE 4.5 - SCANNING ELECTRON MICROSCOPY OF TUBER RUGOSUM ASCOSPORES. A. SEM image of Tuber rugosum ascospore having been passed through the digestive system of the dusky Arion slug. B. SEM image of T. rugosum ascospore cleared from ascus by drying down ascocarp and scraping gleba. C. Representative cluster of T. rugosum ascospores subjected to the slug digestive system. D. Example of remaining ascus, digestive remnants, or other unknown tissue observed on a few of the T. rugosum ascospores SEM-imaged after slug digestion. Bars: A, B, D = 10.0 μm; C = 100.0 μm. 92 Confocal microscopy, Lipid extraction, and analysis Confocal microscopy revealed that Tuber rugosum, sp. nov., BR64 and T. floridanum A. Grupe, Sulzbacher & M.E. Sm. contain spores that are rich in lipid droplets (green fluorescence by BODIPY staining, FIG. 4A, B). This staining made visualizing the high lipid content of the spores respective to the surrounding hyphae of the gleba evident. Further fatty acid analyses of in vitro mycelial growth showed that T. rugosum, sp. nov. (BR64, holotype), has about 60% of polyunsaturated fatty acids (18:2 and 18:3; FIG. 4C), whereas T. lyonii Butters has the highest content of unsaturated fatty acids, including 29% 18:1 (oleic acid), 34% 18:2, and 10% 20:4 (arachidonic acid) (FIG. 4D). Molecular analyses Tuber rugosum, sp. nov., DNA sequences, including for ITS, elongation factor 1α, and RNA polymerase II have been deposited into NCBI GenBank; see TABLE 4 for accession numbers. FIGURE 4.3 - TUBER RUGOSUM ASCOCARP FOUND IN 2021 INFECTED WITH CLONOSTACHYS ROSEA. A. The centermost ascocarp showing the darker red brown peridium seen when infected by C. rosea compared with the peridium color of noninfected ascocarps. B. Orange conidia from C. rosea growing from the gleba of a T. rugosum ascocarp. Bars: A = 20.0 mm; B = 2.0 mm. 93 Phylogenetic analyses. The ITS (FIG. 6), elongation factor 1α (FIG. 7A), and RNA polymerase II (FIG. 7B) phylogenetic trees all place T. rugosum, sp. nov., within the Rufum clade as one of the more early-divergent species. The ITS rDNA data suggest that T. rugosum, sp. nov., is a sister species to T. spinoreticulatum Uecker & Burds. with a maximum likelihood score of 91 and a Bayesian posterior probability score of 98.9 (FIG. 6). Both the elongation factor 1α and the RNA polymerase II phylogenetic trees show the distinct placement of Tuber rugosum, sp. nov., near T. spinoreticulatum and more basal to the other taxa in the Rufum clade (FIG. 7). The elongation factor 1α phylogenetic data provide a maximum likelihood score of 75 for the placement of T. rugosum, sp. nov. (FIG. 7A), and the RNA polymerase II data fall below the threshold of significance with a score of 69 (FIG. 7B). TAXONOMY Tuber rugosum Rennick B., Benucci G.M.N., Du Z., Healy & Bonito, sp. nov. FIG. 2 MycoBank MB838884 Diagnosis Unique to T. rugosum are highly rugose zones across the peridium frequently with tight peridial folds revealing exposed gleba, characteristic echinate ascospores (mean Q = 1.1) that variably have hooked apices with an occasional subreticuate framework sloping gently away basipetally from the spines, and gene sequences. Typification USA. MICHIGAN: Ingham County, Onondaga township, elevation 277 m, found in soil between Quercus rubra and Q. alba in a mixed hardwood forest, 27 Aug 2018, Bryan Rennick 94 BR64 (holotype MSC408483, designated here). GenBank: ITS = MW579343; EF1a = MW584660; RPB2 = MW584657. Etymology The proposed species name references the wrinkly, or rugose, appearance of the ascocarp. Morphology Ascomata irregular to subglobose, irregular or lobate, 7.0–28.1 (x̄ = 12.7) × 10.0– 29.7 (x̄ = 16.8) mm diam, from opaque to translucent beige to whitish, with faint mottled gleba in mature specimen. Glebal marbling, white sterile veins and melanized yellowish beige fertile tissue. Odor mild and almost nutty; flavor mild. Table 4.2 - Ascospore length, width, and shape (Q) measurements based on the number of spores per asci. Spores per asci Spore count (n) Length µm Width µm Q ( x̅ ) 1 11 24–32.5(–37) (19.5–)20.5–25(–26.5) 1.1–1.5 (1.2) 2 12 (19–)20.5–25(–25.5) (17–)18.5–21.5(–22) 1.0–1.4 (1.2) 3 20 (19.5–)20–23(–24.5) (18–)18.5–20.5(–22) 1.0–1.2 (1.1) 4 20 (16–)18–22(–23) (15.5–)16–20(–22) 1.0–1.2 (1.1) 5 22 (18–)18.5–21.5(–22.5) (16.5–)17–18.5(–19) 1.0–1.3 (1.1) Peridium 267.6 ± 41.6 µm thick, glabrous, large zones of rugose pellis. Pellis 139.9 ± 7.5 µm thick. Outermost layer 39.5 ± 6.1 µm thick-walled beige cells subtended by hyaline cells, isodiametric to pseudoparenchymal. Subpellis 97.3 ± 18.9 µm thick. In immature specimen, subpellis distinct from pellis with long, narrow, interwoven cells running perpendicular to the peridial surface. Clavate hyaline asci, 1–7 yellow-brown ascospores, most frequently with 4 ascospores. The main ovoid section of the asci 63.3– 89.2 × 28.6–62.86 μm (X̄ = 75.7 × 42.7 μm), Q = 1.2–2.8 (X̄ = 1.9); peduncle from which the main ovoid section of the ascus arises 39.1–83.9 μm (X̄ = 50.3 μm) in length. 95 Ascospores high in lipids (FIG. 4A, B), subglobose, covered in well-spaced, uncinulate to corniform spines, having a low-sloping ridge extending away from the base and variably fusing with the ridge of adjacent spines, rarely reticulated. Ascospores in 1- spored asci are 24–32.5 × 20.5–26.5 μm with a shape (Q) of 1.2 and in 4-spored asci spores are 18.0–22.0 × 16.0–20.0 μm (Q = 1.1). Additional ascospore size and shape data are in TABLE 2. While growing in vitro, hyphae with simple septae, common branching at right angles (FIG. 1A), rarely producing hyphal coils (FIG. 1B). Hyphae have a high content of unsaturated fats (FIG. 4B, C). Distribution Northeastern North America: Quebec (GenBank HM485428), Minnesota, and Michigan (see TABLE 4). The holotype of Tuber rugosum, sp. nov., was found within meters of an expansive collection of T. floridanum and T. brennemanii A. Grupe, Healy & M.E. Sm. Habitat, Distribution, and Phenology Northeastern North America from Michigan and Minnesota, USA, through Quebec, Canada. Collected Jul–Nov; hypogeous to subhypogeous in previously disturbed soil of mixed hardwood forest dominated by Quercus rubra and Q. alba. Additional specimens examined USA. MINNESOTA: Stearns County, found hypogeous in mixed conifer and deciduous forest, 17 Oct, 2009, Rosanne Healy RH1030 (FLAS-F-61987); MICHIGAN: Ingham County, found hypogeous in mixed hardwood dominated by Q. rubra and Q. alba, 17 Jul 2017, Bryan Rennick BR48 (MSC408482); ibid., found subhypogeous in mixed hardwood dominated by Q. rubra and Q. alba, 7 Aug, 2019, Bryan Rennick BR145 (MSC408484); ibid., found hypogeous in mixed hardwood dominated by Q. rubra and Q. alba, 18 Aug 2019, Bryan Rennick BR159 96 (MSC408485); ibid., found subhypogeous in mixed hardwood dominated by Q. rubra and Q. alba, 11 Sep 2020, Bryan Rennick BR378 (MSC408486); ibid., found subhypogeous in mixed hardwood dominated by Q. rubra and Q. alba, 4 Sep 2021, Bryan Rennick BR428a (MSC409443). Notes Both T. rugosum and T. spinoreticulatum share a distinctive small cavity revealing gleba on most ascoma, as seen in FIG. 2A. Additionally, they share similar habitats among oak trees, found in northeastern North America, with pseudoparenchyma cells forming the pellis and interwoven cells forming the subpellis. However, T. rugosum has a smooth, white to tan peridium surface, whereas T. spinoreticulatum has a leathery, brownish gray peridium. Their aroma also differs in that T. rugosum has a nutty aroma but T. spinoreticulatum smells of rotten cabbage (Uecker and Burdsall, 1977). Finally, T. rugosum has small, spiny ascospores at 16.1– 25.7 × 15.4–22.0 μm, but T. spinoreticulatum has larger spiny-reticulate ascospores at 30–35 × 22–25 μm. On the same agar medium described in Materials and Methods, Tuber rugosum grows vegetatively with right-angled branching and simple septae, as shown in FIG. 1A. In older cultures (FIG. 1B), hyphal coils can be seen once the culture becomes stressed. We found some collections of T. rugosum that were infected by a fungicolous species that we identified as Clonostachys rosea. When infected by C. rosea, the aroma was particularly smoky and the peridium turned a darker shade with more pronounced red -brown hues, as shown in FIG. 3. The Clonostachys rosea isolate (BR428b) maintained a faint smoky aroma from the pure isolate after more than two transfers beyond the initial isolation plate, but directed 97 experiments will be needed to test the impact of this mycoparasite on truffle aroma. Cultures of the holotype (BR64) as well as C. rosea (BR428b) are available upon request. 98 Table 4.3 - Characteristics of morphologically similar Tuber species within the Rufum clade as reported in primary literature. Peridium Pellis Thickness Species Peridium surface Peridium color Thickness (μm) Pellis Cell Type (μm) Subpellis Cell Type Tuber lyonii Smooth, Slightly Pruinose Light Chestnut Brown 300–500 Interwoven 20–40 Interwoven Tuber liaotongense Verrucose Brownish Yellow Not Reported Pseudoparenchyma Not Reported Prosenchyma White Yellow, Pale Yellow or Tuber microspiculatum Glabrous 200–250 Pseudoparenchyma 50–100 Interwoven Light Brown, Reddish Brown Tuber quercicola Verrucose Dark Red to Brownish Red 250–500 Interwoven 20–100 Interwoven Light Yellowish Brown to Reddish Irregularly Compact to Tuber candidum Smooth 100–300 30–100 Interwoven Brown Elongated Cells Tuber ferrugineum Papillose Reddish 215–390 Interwoven 15–50 Interwoven Tuber nitidum Glabrous Reddish Yellow Not Reported Not Reported Not Reported Not Reported Tuber rufum Minutely Warty to Smooth Reddish Brown Not Reported Pseudoparenchyma Not Reported Interwoven Tuber melosporum Small Warts Reddish Brown to Reddish Black Not Reported Pseudoparenchyma Not Reported Interwoven Tuber wenchuanense Smooth Grey Brown 200–250 Pseudoparenchyma 50–100 Interwoven Pseudoparenchyma Tuber malacodermum Smooth Light Brown 300–450 Pseudoparenchyma Not Reported to Interwoven Pale Yellow Brown to Yellow Tuber piceatum Smooth and Glabrous 200–350 Pseudoparenchyma Not Reported Interwoven Brown Interwoven To Tuber crassitunicatum Smooth Brown to Yellow Brown 250–300 200–250 Interwoven Pseudoparenchyma Tuber spinoreticulatum Leathery Brownish Gray Not Reported Pseudoparenchyma 150–600 Interwoven Pseudoparenchyma Tuber theleascum Smooth Reddish 160–250 Pseudoparenchyma 45–150 to Interwoven Pale Yellow, Yellow Brown or Tuber taiyuanense Smooth 150–300 Pseudoparenchyma Not Reported Interwoven Brown Tuber pustulatum Low Pyramidal Warts Reddish 370–550 Pseudoparenchyma 160–220 Prosenchyma Tuber lishanense Smooth Yellow White to Yellow Brownish 250–350 Pseudoparenchyma 150–200 Interwoven Subglobose or Tuber luomae Verrucose Light Orange Brown ± 500 ± 150 Interwoven Subpolyhedral Tuber rugosum Smooth White to Tan 267.6 ± 41.6 Pseudoparenchyma 139.9µm ± 7.5µm Interwoven Pale Yellow, Yellow Brown or Tuber umbilicatum Smooth to Minute Papillae 320–500 Pseudoparenchyma 90–250 Interwoven Brown Tuber furfuraceum Smooth Brown 340–480 Pseudoparenchyma 170–270 Interwoven Verrucose, Slightly Tuber huidongense Yellow-Brown to Red-Brown 150–300 Pseudoparenchyma 80–150 Interwoven Furfuraceous, Pubescent Tuber lannaense Smooth Yellow-Brown to Dark-Brown 130−260 Pseudoparenchyma 35–80 Interwoven Tuber wanglangense Smooth and Glabrous Yellow White 200–250 Pseudoparenchyma 50–100 Interwoven 99 Table 4.3 - (cont’d) Subpellis Spore Shape (Q or Spores per Species Average Spore Size (μm) Ornamentation Aroma Source Thickness (μm) Description) Ascus Spinose To Pungent, Nutty, (Butters, 1903; Trappe et al., Tuber lyonii Not Reported 21–31 x 17–21 Ellipsoid 1–5 Subreticulate Malted Milk 1996) Alveolate Tuber liaotongense Not Reported 29–40 x 26–35 Ellipsoid 2–4 Not Reported (Wang, 1988) Reticulate Alveolate Tuber microspiculatum Not Reported 22.5–35 x 17.5–22.5 Ellipsoid 1–4 Slight, Not Pungent (Fan et al., 2012) Reticulate Earthy, Fresh Green Tuber quercicola 130–400 20–45 x 15–35 Ellipsoid 1–5 Curved Spines (Frank et al., 2006) Beans Mild to Slightly (Harkness, 1899; Frank et al., Tuber candidum 70–200 19–42 x 14–34 Globos to Ovoid 1–5 Curved Spines Earthy 2006) (Carlo Vittadini, 1831; Elliott Tuber ferrugineum 200–350 28.2 x 20.5 Ellipsoid 2–4 Spiny Silkworm / Pleasant et al., 2016) (Carlo Vittadini, 1831; Tuber nitidum Not Reported Not Reported Ellipsoid 1–4 Echinulate Nauseous Granetti et al., 1988) (Carlo Vittadini, 1831; Tuber rufum Not Reported 28–42 x 18–28 Ellipsoid 4–5 Spiny Insipid Grunow & Rabenhorst, 1884) Tuber melosporum Not Reported 40–45 x 20–25 1.7 1–6 Smooth Garlic (Enrico et al., 2016) Tuber wenchuanense 150–200 25–45 x 17.5–30 Ellipsoid 1–5 Spiny-Reticulate Not Reported (Fan et al., 2013) Tuber malacodermum Not Reported 26.0 ± 4.6 x 21.1 ± 3.5 1.23 ± 0.12 1–4 Spinose Celery (Leonardi et al., 2019) Tuber piceatum Not Reported 26.7 ± 2.7 x 19.2 ± 1.4 1.4 ± 0.1 1–6 Spinose Not Reported (Yan et al., 2018) Tuber crassitunicatum Not Reported 29.1 ± 4.8 x 18.8 ± 2.5 1.5 ± 0.1 1–5 Spiny-Reticulate Mild (Yan et al., 2018) Tuber spinoreticulatum 30–125 30–35 x 22–25 1.4 1–5 Spiny-Reticulate Rotting Cabbage (Uecker & Burdsall, 1977) Spinose To Tuber theleascum 85–120 26.6 ± 5.18 x 16.8 ± 2.63 1.59 ± 0.19 1–6 Not Reported (Leonardi et al., 2019) Subreticulate Tuber taiyuanense Not Reported 20–45 x 18–30 Ellipsoid 1–5 Spiny-Reticulate Light (Cao et al., 2011) Acidulous To Tuber pustulatum 210–350 27.4 ± 5.7 x 22.6 ± 4.55 1.22 ± 0.3 1–6 Spiny-Reticulate (Leonardi et al., 2019) Rancid Tuber lishanense 100–150 25.5 ± 3.8 x 21.6 ± 3.7 1.2 ± 0.2 1–5 Spiny Inconspicuous (Yan et al., 2018) Tuber luomae ± 350 23–30 x 18.5–23 1.21–1.32 1–5 Spiny Mildly Acrid (Eberhart et al., 2020) Tuber rugosum 97.3 ± 18.9 21.5 x 19.4 1.2 1–7 Spiny Nutty Spiny Alveolate Tuber umbilicatum 150–400 23–33 x 17–23 1.4 ±0.13 1–6 Not Reported (Chen et al., 2005) Reticulum Slight, Not Tuber furfuraceum 170–210 25–46 x 14–27 1.7 2–5 Spiny-Reticulate (Hu & Wang, 2005) Distinctive Tuber huidongense 90–150 27–35 x 18–22 1.50 ± 0.18 1–5 Spiny-Reticulate Not Reported (Deng et al., 2009) Tuber lannaense 100–175 25−29 x 17−21 1.17 ±0.14 1–5 Spiny-Reticulate Not Reported (Suwannarach et al., 2016) Tuber wanglangense Not Reported 29.9 ± 3.6 x 24.5 ± 2.6 1.2 ± 0.1 1–5 Spiny-Reticulate Not Reported (Yan et al., 2018) 100 Table 4.4 - Tuber rugosum collections used in this study with their herbarium and GenBank accession numbers and collection dates. Accession Number Date Species Collection Source Locale Collected Herbarium ITS1-F EF1α RPB2 Tuber rugosum RH999 Ascoma USA:MN 8 Aug 2009 MW584702 Tuber rugosum RH1030 Ascoma USA:MN 17 Oct 2009 FLAS-F-61987 MW584701 Tuber rugosum RH1330 Ascoma USA:MN 3 Sep 2011 MW584700 Tuber rugosum BR48 Ascoma USA:MI 17 Jul 2017 MSC408482 MW579335 Tuber rugosum BR64 Ascoma USA:MI 27 Aug 2018 MSC408483 MW579343 MW584660 MW584657 Tuber rugosum BR145 Ascoma USA:MI 7 Aug 2019 MSC408484 MW579344 MW584661 MW584658 Tuber rugosum BR159 Ascoma USA:MI 18 Aug 2019 MSC408485 MW579345 MW584662 MW584659 Tuber rugosum BR339 Root tip USA:MI 5 Aug 2020 MW579346 Tuber rugosum BR340 Root tip USA:MI 5 Aug 2020 MW579347 Tuber rugosum BR342 Root tip USA:MI 5 Aug 2020 MW579348 Tuber rugosum BR343 Root tip USA:MI 5 Aug 2020 MW579349 Tuber rugosum BR378 Ascoma USA:MI 11 Sep 2020 MSC408486 MW579975 Tuber rugosum BR428a Ascoma USA:MI 4 Sep 2021 MSC409443 OL438889 Clonostachys rosea BR428b Anamorph USA:MI 4 Sep 2021 MSC409443 OL438890 101 FIGURE 4.4 - LIPID ANALYSIS OF TRUFFLE SPORES. A, B. Confocal microscopy of BODIPY-stained (A) Tuber rugosum (BR64) and (B) Tuber floridanum ascospores revealing lipid content as green. C, D. FAME analysis of (C) T. rugosum and (D) T. lyonii in vitro mycelial growth showing distinct variation in fatty acid concentrations. Bars: A = 0.5 mm, 100.0 μm, 25.0 μm, respectively; B = 0.5 mm, 100.0 μm, and 30.0 μm, top to bottom. 102 FIGURE 4.6 - ITS RDNA PHYLOGENY OF THE RUFUM CLADE. This most likely phylogenetic tree reconstructed from ITS rDNA data shows that T. rugosum is basal in the 103 FIGURE 4.6 (cont’d) – Rufum clade and supported as sister species to T. spinoreticulatum . Maximum likelihood bootstrap support values over 70 are shown above the nodes, whereas Bayesian posterior probabilities above 95 are shown below nodes. Tuber regimontanum, T. indicum, and T. melanosporum were included as outgroups as identified by Bonito et al. (2010). Taxa are shown with specimen, isolate, or collection number as listed in the NCBI database. FIGURE 4.7 - ELONGATION FACTOR 1Α (A) AND RNA POLYMERASE II GENE (B) PHYLOGENETIC TREES. Both show maximum likelihood bootstrap support values over 70. Tuber indicum was chosen as an outgroup as identified by Bonito et al. (2010). Taxa are shown with specimen, isolate, or collection number as listed in the NCBI database. 104 FIGURE 4.8 - TRUFFLE SPORES IN SLUG FRASS. A) Dusky Arion slug near T. rugosum ascoma. B, C) Slug frass showing high amount of T. rugosum ascospores. D) Slug frass mounted on slide with 3% KOH showing T. rugosum ascospores free of their ascus. Bars: A = 20.0 mm, B = 200.0 μm, C = 500.0 μm, D = 200.0 μm Discussion Here, we described a new truffle species, T. rugosum, as supported by three independent phylogenetic markers, as well as by morphological characters. Phylogenetic reconstructions demonstrate this species as a novel North American species in the Rufum clade. Tuber rugosum is most closely related to the North American species T. spinoreticulatum, and as seen in TABLE 3, the two species differ macroscopically in the color and texture of their peridium and microscopically by the size, shape, and ornamentation of their ascospores. The two species also 105 have a different aroma, with Tuber spinoreticulatum being particularly unpleasant and originally described as smelling like rotten cabbage (Uecker and Burdsall, 1977). Animals play an important role in the spore dispersal of hypogeous fungi. This has been well documented in the case of small mammals, which consume truffles as food (Cázares and Trappe, 1994; Gabel et al., 2010). Other animals including Diptera and Stylommatophora have fungivorous members that have been shown to enhance mycorrhizal spore dispersal, including that of truffles (Kitabayashi and Tuno, 2018; McGraw et al., 2002). As documented previously, slugs in the genus Arion are mycophagous (Beyer and Saari, 1978); however, their role in spore dispersal has not been investigated. Although this paper did not set out to determine truffle spore viability in slug excreta, we did demonstrate that T. rugosum is consumed by Arion slugs and that spores that pass through the slug digestive tract are released from the asci in good condition for visualization. In fact, we found this to be a useful pretreatment for cleansing spores prior to preparation for SEM. Recent work by Ori et al. (2021) demonstrated that slug digestion of T. aestivum ascospores exhibit an altered episporial texture and increased mycorrhization of Quercus robur roots compared with those ascospores consumed by mice or uningested spores. Our observations that T. rugosum is found just beneath leaf litter and is often partially consumed raise questions on the role of Arion slugs and other gastropods in truffle spore dispersal in nature. Truffles damaged by small animals or other mechanical means may also become more susceptible to infection by fungicolous organisms, which may alter its aromatic profile (Eslick, 2012). It is still unknown whether C. rosea is a primary pathogen of truffles, although the literature suggests that C. rosea can be a mycoparasite of many fungal species and thus may have potential use as a biocontrol against fungal crop pathogens such as Fusarium graminearum Schwabe (Gimeno et al., 2021). In a prior report, C. rosea was isolated from Tuber magnatum 106 but was not found to be chitinolytic (Pavic et al., 2013). We were unable to demonstrate C. rosea pathogenicity on T. rugosum, as our in vitro assays were inconclusive. While growing C. rosea on agar, we noted that it maintained a faint smoky aroma, which supports the hypothesis that it contributed to the odor profile of the ascocarps it was found growing on. Other truffle-inhabiting organisms, particularly α- and β-proteobacteria, have been found to be principal contributors to sulfur-containing volatiles characteristic of T. borchii Vittad. ascocarp aroma (Splivallo et al., 2011). Further work is needed to ascertain the involvement of other fungi and bacteria in the aroma development and profiles of Tuber ascoma, as well as truffle development and disease. Fatty acid profiles have been used when trying to distinguish Tuber species within species complexes and provide some insight into their physiology and nutrition (Angelini et al., 2015). However, Tuber fatty acid profiles may vary across geographic regions and under different environmental and growth conditions (Shah et al., 2020). To attempt to control for these environmental variances, we obtained pure culture isolates of T. rugosum and the closely related species T. lyonii (FIG. 4), to assess and compare FAME profiles from similarly aged mycelium grown in the same medium and environment. Truffle species had distinct fatty acid profiles from one another, with T. rugosum being particularly reduced in 18:1 (oleic acid) and T. lyonii enriched in 20:4 (arachidonic acid). Fatty acid profiles are not available for most Tuber species, but they could provide insights into variation in Tuber physiology and nutrition. Similar to previously published ITS phylogenies (Bonito et al., 2010), T. rugosum is found on an early-divergent branch of the Rufum clade and appears to be sister to T. spinoreticulatum (FIG. 6). Our ITS ML- and BI-based phylogenies also conform to more recently published Tuber phylogenies (Yan et al., 2018), which placed T. lishanense L. Fan & X.Y. Yan and T. piceatum L. Fan, X.Y. Yan & M.S. Song basal to T. spinoreticulatum within the 107 Rufum clade. The general structure of the phylogenetic trees analyzed from the two protein- encoding loci (FIG. 7) also agree with earlier studies (Bonito et al., 2013) but with stronger support for T. rugosum within the Rufum clade and its close relationship to T. spinoreticulatum . However, it should be noted that there are relatively few sequences available within the Rufum clade for both the RPB2 and EF1α protein-encoding regions. The lack of species representation within these protein-encoding regions is such that concatenating and condensing the phylogenetic trees into a single, better-supported phylogram is not possible at this time. Further work is needed to generate these protein-encoding sequences in other Rufum clade species in order to reconstruct a more comprehensive phylogeny. Conclusion In conclusion, we have described Tuber rugosum, a pale, wrinkly, spiny-spored truffle endemic to northeastern North America, supported by morphological and phylogenetic analyses of multiple loci. We provide the fatty acid profile of this species, describe a fungicolous species association, and present a method that involves the use of slugs to assist in cleaning ascospores prior to SEM imaging. Finally, a dichotomous key for truffles in the Rufum clade is provided below. Together, these results expand on the knowledge base of Tuber biodiversity and microbiome diversity in North America. A key to Tuber spp. in the Rufum clade 1. Ascospore smooth with no ornamentation T. melosporum 1’. Ascospore with ornamentation 2 2. European species 3 2’. Not European species 7 108 3. Peridium smooth 4 3’. Peridium not smooth 5 4. Peridium color reddish yellow T. nitidum 4’. Peridium color light brown T. malacodermum 5. Ascospore globose in shape T. pustulatum 5’. Ascospore ellipsoid in shape 6 6. Pellis cells pseudoparenchyma T. rufum 6’. Pellis cells interwoven T. ferrugineum 7. North American species 8 7’. Asian species 14 8. Peridium smooth 9 8’. Peridium verrucose or leathery 12 9. Ascospore spines commonly curved or hooked 10 9’. Ascospore spines not commonly curved or hooked 11 10. Found west of the Rocky Mountains T. candidum 10’. Found east of the Rocky Mountains T. rugosum 11. Pellis 20–40 μm thick T. lyonii 11’. Pellis 45–150 μm thick T. theleascum 12. Outer pellis of pseudoparenchyma or globos cells 13 12’. Outer pellis of longer interwoven cells T. quercicola 13. Ascospore globose in shape T. luomae 13’. Ascospore ellipsoid in shape T. spinoreticulatum 109 14. Peridium verrucose or with small papillae 15 14’. Peridium smooth 17 15. Ascospore reticulated but with distinct curved spines T. umbilicatum 15’. Ascospore not as above 16 16. Ascospore alveolate reticulate T. liaotongense 16’. Ascospore spiny-reticulate T. huidongense 17. Pellis consisting of pseudoparenchyma 18 17’. Pellis consisting of interwoven cells T. crassitunicatum 18. Ascospore reticulate 19 18’. Ascospore not reticulate 24 19. Ascospore alveolate reticulate T. microspiculatum 19’. Ascospore spiny-reticulate or spiny 20 20. Ascospore globose in shape 21 20’. Ascospore ellipsoid in shape 22 21. Peridium color yellow brown to dark brown; 130–260μm thick T. lannaense 21’. Peridium color yellow white; 200–25 μm thick T. wanglangense 22. Peridium thickness > 300 μm T. furfuraceum 22’. Peridium thickness < 300 μm 23 23. Ascoma deeply and densely furrowed T. taiyuanense 23’. Ascoma mostly smooth with few furrows T. wenchuanense 24. Ascospore globose in shape T. lishanense 24’. Ascospore ellipsoid in shape T. piceatum 110 Chapter 5 Synthesis 111 Synthesis Objectives The three primary goals of this thesis were to: 1) to investigate how glyphosate and glufosinate herbicides interact with truffles, morels, and plant growth promoting fungi in-vitro to inform use in fungal cultivation, 2) to investigate methyl bromide fumigated bare-root conifer tree nurseries as a potential candidate for targeted inoculation of local, exotic, or commercial ectomycorrhizal inoculants, and if these inoculants improve seedling growth, and 3) to confirm and describe a new truffle species utilizing morphology, a unique genetic sequence, a unique fatty acid profile, a novel electron microscopy spore cleaning method, and to build a new taxonomic key for Rufum clade. Efforts toward each of these goals are detailed through chapters 2, 3, and 4. Each of the major results and future directions of these chapters are synthesized here. Herbicide interactions In Chapter 2, I investigated the growth implications of several fungi cultured with media made with serial dilutions of glyphosate or glufosinate herbicides. I focused on Benniella erionia, Linnemannia elongota, Morchella americana, M. importuna, Tuber borchii, T. canaliculatum, T. floridanum, T. lyonii, and T. rugosum. Additionally, I assessed impacts of Rhizopus oryzae, Serendipita indica, Stropharia rugosoannulata, and an Umbeloposis species to the same growth assay with fewer replications. The results of this work revealed that as both herbicides increased in concentration to levels near or above the manufacturers recommended field application rate, fungal growth diminished. It also showed that ectomycorrhizal species were less sensitive to herbicides than saprotrophic fungi. This study demonstrated fungistatic properties of herbicides commonly used in truffle orchards and other environments. These effects were evident when overapplied; something easy to do when applying by hand. 112 Additional work may aim to investigate a wider array of pesticides including bactericides, insecticides, and oomyticides. Beyond investigating more pesticides, pesticide residuals and adjuvants may interfere with more than just mycelial growth. Future work elucidating how pesticides impact development of overwintering structures, fruiting bodies, and mitospores should be carried out. Additionally, as there are many truffle plantations using herbicides and morels cultivated between agronomic crops, we should investigate which herbicide residuals end up in these food items and how concentrated these pesticides may become. Added inoculants to bare-root nurseries In Chapter 3, I assessed how adding various inoculants to methyl bromide fumigated bare-root Christmas tree nurseries may impact seedling health and growth. Specifically, through two experiments at two nurseries over two years, Pseudotsuga menziesii, Picea abies, and Pinus sylvestris were inoculated with local (Laccaria bicolor or Scleroderma citrinum), exotic truffle (Tuber aestivum, Tuber borchii, or Tuber indicum), or a commercial inoculant (MycoApply®) at the time of sowing. Prior to the nursery's harvest the following year, seedlings from each treatment block were collected. Seedling height, root collar diameter, above ground biomass were collected. Additionally, fungal and oomycete MiSeq libraries were constructed for community analysis. The results of this work show no increases in height, diameter, or biomass over uninoculated control blocks. I did find low levels of Tuber borchii reads in the community analysis, but they were well below a threshold that would interest truffle plantations. Additionally, the oomycete library revealed over 12 species of Pythium and four species of Phytophthora, highlighting how well these motile pathogens can persist in or recolonize methyl bromide fumigated soils. Overall, it seems that fungi either persisting or blowing in from nearby 113 tree stands dominate the roots of these Christmas trees. The fungal diversity found on these roots combined with the lack of growth improvements suggest that adding inoculants may not be beneficial or necessary. This work could be expanded by looking at other tree nursery species and other inoculant sources. Additionally, investigating the community of spores introduced from nearby tree stands as they relate to living mycorrhizae in nursery seedlings may reveal one avenue by which these robust communities establish. Christmas trees are grown for two years at the bare-root nursery before getting transplanted into a larger space where it will grow to harvestable size. It would also be beneficial to look at how the fungal community adapts to transplanting and any treatments which may occur at the finishing plantation as there is interest in adding inoculants at this phase as well. A new truffle species Chapter 4 highlights my work done in describing a new truffle species found in Michigan, and the greater north eastern portion of North America. Tuber rugosum, was described using a multigene phylogeny, a fatty acid profile, and morphological characters. Assisting with the morphological analysis, observations of a slug (Arion subfuscus) eating subhypogeous truffles led to the development of a novel ascus clearing technique for scanning electron microscopy. Slug fungivory removes asci from ascospores with delicate ornamentation, such as with spiny-spored T. rugosum, without damaging spore morphology. Additionally, I produced an updated taxonomic key for the Rufum clade. This work describing rare truffle taxa removes ambiguity in future taxonomic efforts and aids in expanding our knowledge of truffle ranges, diversity, and ecology. 114 Further work is still needed to describe other rare truffle species as they directly interact with tree species in truffle cultivation systems. Beyond their ecological roles, some yet to be described truffle species may also have important culinary value. Additionally, there is very little in the literature about slug fungivory and how it relates to ascospore viability. Developing this knowledge could yield significant insights to both truffle and slug ecologies as well as having the potential to improve truffle tree colonization. Conclusions Truffles have captivated our attention for centuries and with recent progress in truffle cultivation, these rare gems of the earth are more available and are gaining even greater interest as they make their way to restaurants around the world. There are still many questions on how best to manage their plantations such as how using herbicides may impact truffle development. I have shown that truffles, but also morels and plant beneficial fungi, have reduced mycelial growth at herbicide application rates above the recommended dosage. If truffières decide to use herbicides, they should not apply glyphosate or glufosinate carelessly or at rates above the manufacturers label. Additionally, at least in terms of outdoor bare-root conifer trees, adding inoculants at the time of sowing does not improve growth and therefore not recommended. Adding inoculants not only requires the purchase of costly products, but also consumes valuable labor during an already tight schedule. Finally, as was the case with Tuber rugosum, there may be new species of truffle in ones backyard. Careful observations of this new species led to the development of a new technique for clean electron microscopy. 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Journal of Forest Research, 20(6), 493–500. https://doi.org/10.1007/s10310-015- 0506-1 132 APPENDIX CURRENT CURRICULUM VITAE Bryan Rennick Michigan State University | Department of Plant, Soil and Microbial Science 3285 Molecular Plant Sciences Building | 1066 Bogue St. East Lansing, MI 48824 rennickb@msu.edu Education Michigan State University, East Lansing, Michigan Present • Master of Science in Plant Pathology; GPA 4.0 Michigan State University, East Lansing, Michigan 2018 • Bachelor of Science in Horticulture; Horticultural Science concentration. • Summa Cum Laude; GPA 4.0 Lansing Community College, Lansing, Michigan 2016 • General Studies Community College of the Air Force, Maxwell Air Force Base, Alabama 2009 • General Studies, Applied Science with focus in Ecological Controls United States Air Force, Sheppard AFB, Wichita Falls, Texas 2004 • Utility Systems Maintenance Technical Training PUBLICATIONS Peer reviewed • Rennick B, Benucci GMN, Du Zhi-Yan1, Healy R, Bonito G. (2023) Tuber rugosum , a new species from Northeastern North America: Slug mycophagy aides in electron microscopy of ascospores. Mycologia 133 • Dissanayake A, Mills G, Bonito G, Rennick B, Naira M. (2021) Chemical Composition, Anti-inflammatory and Antioxidant Activities of Extracts from Cultivated Morel Mushrooms, Species of Genus Morchella (Ascomycota). International Journal of Medicinal Mushrooms. • Benucci GMN, Rennick B, Bonito G (2020). Patient propagules: Do soil archives preserve the legacy of fungal and prokaryotic communities? PLoS ONE. In Preparation for Peer Review Journals • Rennick B, Benucci GMN, Bonito G. Impacts of Liberty and Roundup herbicides on mycelial growth of truffles, morels, and plant growth promoting fungi. • Rennick B, Benucci GMN, Bonito G. Does ectomycorrhizal inoculum after methyl bromide field applications in bare-root conifer nursery seedlings establish and persist? Trade Journal • Rennick B, Benucci GMN, Bonito G. (2022) How do methyl bromide field applications and mycorrhizal inoculants impact bare-root conifer nursery seedlings ectomycorrhizal development? Great Lakes Christmas Tree Journal. PROFESSIONAL PRESENTATIONS Guest Lectures • Rennick, B., (February 20, 2023) Growing Small Scale Farms by Growing Gourmet Mushrooms. Guest Lecture for HRT 341: Vegetable Production and Management, MSU, East Lansing, MI, United States • Rennick, B., (February 21, 2022) Growing Small Scale Farms by Growing Gourmet Mushrooms. Guest Lecture for HRT 341: Vegetable Production and Management, MSU, East Lansing, MI, United States 134 • Rennick, B., (March 1, 2021) Growing Small Scale Farms by Growing Gourmet Mushrooms. Guest Lecture for HRT 341: Vegetable Production and Management, MSU, East Lansing, MI, United States • Rennick, B., (January 21, 2020) Cultivating the Mushroom of Immortality. Guest Lecture for PLP 405: Plant Pathology, MSU, East Lansing, MI, United States • Rennick, B., (February 25, 2019) Growing Small Scale Farms by Growing Gourmet Mushrooms. Guest Lecture for HRT 341: Vegetable Production and Management, MSU, East Lansing, MI, United States Poster Presentations • Rennick, B., Bennuci, G. N., Zhi-Yan Du, Bonito GM (October 23, 2019) Tuber rugosum sp. nov.: A new spiny-spored truffle species from North America. Presented poster at IWEMM10, Nagano, Japan • Rennick, B., Bennuci, G. N., Bonito GM (October 23, 2019) Impact of the herbicide Roundup® on mycelial growth of truffles, morels, and mold. Presented poster at IWEMM10, Nagano, Japan • Rennick, B., Bennuci, G. N., Bonito GM (July 18, 2018) Impact of glyphosate and glufosinate on mycelial growth of truffles, morels, and molds. Presented poster at MSA annual meeting, San Juan, PR, United States. Professional Talks • Rennick, B., (2018) Target Mycorrhizal Fungi in Nursery and Forestry Cropping Systems. Department of Plant, Soil, and Microbial Sciences Plant Pathology Seminar, MSU, East Lansing, MI, United States • Benucci, G., Rennick, B., (2017) Strategies for inoculating chestnut seedlings with truffles. 135 Midwest Nut Producers Council. Clarksville, MI, United States AWARDS AND ACHIEVEMENTS • Plant, Soil, and Microbial Sciences Endowed Graduate Assistantship • Mycological Society of America James M. Trappe Travel Award • American Society for Horticultural Science 2018 Outstanding Undergraduate Student Award • 8 Semesters on Dean’s List • Merrill S. Fuller Scholarship • Victor Ray Gardner Scholarship • Terry L. Schlichter Endowed Scholarship • Carl J. Sellner Scholarship • Air Force Longevity Service Award • Twice awarded the Air Force Achievement Medal • Twice awarded the Air Force Good Conduct Award • Air Force Expeditionary Service Ribbon with Gold Border • Global War on Terrorism Expeditionary Medal 136