SYSTEMATIC ANALYSIS OF RUSSULA IN THE NORTH AMERICAN ROCKY MOUNTAIN ALPINE ZONE by Chance Ray Noffsinger A thesis submitted in partial fulfillment of the requirements for the degree of Master of Science in Plant Pathology MONTANA STATE UNIVERSITY Bozeman, Montana April 2020 ©COPYRIGHT by Chance Ray Noffsinger 2020 All Rights Reserved ii ACKNOWLEDGEMENTS This project would not have been possible without the enthusiasm, intelligence, and guidance of my advisor Cathy Cripps. She has helped me grow as a person, develop as a scientist, and build a foundation for my future. Her teachings will guide me through the rest of my life, in and out of the laboratory, and I will always be grateful for that. I would like to thank Matt Lavin for his assistance with phylogenetics and for our thought- provoking discussions. His dedication to understanding biodiversity along with his impressive active lifestyle have inspired me to balance my career and athletic aspirations. I wish to show my gratitude to Sara Branco for her continued support throughout this project, which has undoubtedly improved its quality. Her dedication to mycology and evolution has shaped my scientific interests. I would like to recognize all of my sources of funding including the John W. Marr Fund, Ben Woo Scholarship, Plant Science and Plant Pathology Department, College of Agriculture, and S.W. Montana Mycology Society. I would like to thank all the herbaria that loaned collections including the Denver Botanic Garden, University of Michigan fungal herbarium, the New York Botanical Garden, the Botanical Museum at the University of Oslo, Arctic University of Norway, and Åbo Akademi University in Finland. I am also indebted to all who have contributed to this project including Sue Brumfield, Egon Horak, Jukka Vauras, Ursula Peintner, and Vera Evenson. I would like to thank Anna Bazzicalupo, Olivia Anderson, Ed Barge, and Marlee Jenkins for their assistance in the laboratory. Lastly, I would like to recognize my mother, Carmen Noffsinger, and my partner, Lauren Baumgardner, for their emotional support and for tolerating my obsessive rants about Russula. iii TABLE OF CONTENTS 1. LITERATURE REVIEW: HISTORY OF ARCTIC AND ALPINE MYCOLOGY IN NORTH AMERICA AND INTRODUCTION TO THE GENUS RUSSULA ................................................................................................. 1 Arctic-Alpine Biome ....................................................................................................... 1 Arctic and Alpine Fungi .................................................................................................. 3 The History of Arctic and Alpine Mycology in North America ..................................... 5 Early Expeditions to the Canadian Arctic (1819–1940) .......................................... 6 Detailed Morphological Studies ............................................................................ 11 Arctic Micromycetes ................................................................................. 12 Arctic Macrofungi ..................................................................................... 13 Arctic and Alpine Fungal Ecology ............................................................ 18 The International Symposium on Arctic and Alpine Mycology (1980–2016) .......................................................................................................... 20 Fungi in the Rocky Mountain Alpine Zone ........................................................... 22 Ecological and Biogeographic Research on Arctic and Alpine Fungi in North America ......................................................................................... 27 Ectomycorrhizal Fungi and Plant Hosts in Arctic-Alpine Environments ..................... 28 The Rocky Mountains ................................................................................................... 31 Russulales ...................................................................................................................... 33 Russulaceae ................................................................................................................... 36 Russula Persoon ............................................................................................................ 40 Morphological Characters ..................................................................................... 43 Chemical Reactions ............................................................................................... 46 Ecology and Hosts ................................................................................................. 48 Russula in Arctic-Alpine Habitats of Europe, Asia, and Arctic Islands .................................................................................................................... 52 North American Studies of Russula ...................................................................... 59 Arctic and Alpine Research on Russula in North America ................................... 61 2. SYSTEMATIC ANALYSIS OF RUSSULA IN THE NORTH AMERICAN ROCKY MOUNTAIN ALPINE ZONE ................................................. 65 Introduction ................................................................................................................... 65 Methods ......................................................................................................................... 74 Study Sites ............................................................................................................. 74 Taxon Sampling and Processing ........................................................................... 77 Morphological Descriptions .................................................................................. 78 Scanning Electron Microscopy .............................................................................. 82 Compound Microscope Photography .................................................................... 82 DNA Extraction, PCR Amplification, DNA Purification, and Sequencing ............................................................................................................ 92 iv TABLE OF CONTENTS CONTINUED Sequence Alignment and Phylogenetic Analyses ................................................. 97 Maps of Russula Species Distributions ............................................................... 101 Results ......................................................................................................................... 101 Phylogenetic Analyses ......................................................................................... 101 Subgenus Russula .................................................................................... 105 Subgenus Brevipes ................................................................................... 112 Ecology of Russula in the Rocky Mountain alpine ............................................. 114 Taxonomy ................................................................................................................... 114 Russula altaica .................................................................................................... 115 R. heterochroa ..................................................................................................... 121 R. laccata ............................................................................................................. 125 R. laevis ............................................................................................................... 132 R. montana ........................................................................................................... 137 R. nana ................................................................................................................. 143 R. cf. pascua ........................................................................................................ 151 R. purpureofusca ................................................................................................. 156 R. saliceticola ...................................................................................................... 162 R. subrubens ........................................................................................................ 168 Key 1. Russula in the central and southern Rocky Mountain alpine zone .......................................................................................... 182 Discussion ................................................................................................................... 183 Addendum – Alaskan species of Russula ................................................................... 198 Key 2. Red-capped Russula from Alaska, mostly with Betula at treeline .......................................................................................... 199 Taxonomy of Alaskan Species .................................................................................... 200 R. cf. alpigenes .................................................................................................... 200 R. intermedia ....................................................................................................... 204 R. cf. sphagnophila .............................................................................................. 209 R. aff. vinosa ........................................................................................................ 212 REFERENCES CITED ................................................................................................... 215 APPENDICES ................................................................................................................. 257 APPENDIX A: C-Tab Extraction Protocol with Lab Notes ............................... 258 APPENDIX B: Preparation of Primers Ordered from Integrated DNA Technologies (IDT) .................................................................. 261 APPENDIX C: Seqtrace Protocol ....................................................................... 263 APPENDIX D: Code Required to Run the Desktop Version of Muscle on Windows ........................................................................................ 265 APPENDIX E: Maximum Likelihood Phylogenies Produced ............................ 267 APPENDIX F: Beast Protocol ............................................................................. 272 v TABLE OF CONTENTS CONTINUED APPENDIX G: Best-fitting Substitution Models Determined by Partition Finder for Bayesian Analyses .......................................................... 274 APPENDIX H: R Code for Maps of Russula Species Distributions ........................................................................................................ 276 vi LIST OF TABLES Table Page 1. Selected surveys of Arctic and alpine fungi in Europe, Asia, and Arctic Islands ....................................................................................................... 4 2. Macromycetes collected during early Arctic expeditions from 1819 through 1931 ............................................................................................ 10 3. Research published in the proceedings of the ten International Symposia on Arctic and alpine mycology focused on North American Fungi ................................................................................................. 21 4. Reports of Arctic-alpine Russula species in Eurasia, on Arctic Islands, and in North America .......................................................................... 53 5. Collections whose micromorphological features were examined or were included in the phylogenetic analyses .................................................. 83 6. Primer sequences used in this study .................................................................. 95 vii LIST OF FIGURES Figure Page 1. Select microscopic features in Russula ............................................................. 81 2. Maximum likelihood phylogeny of the genus Russula ................................... 104 3. Bayesian posterior probability phylogeny of the Russula crown clade .......... 108 4. Bayesian posterior probability phylogeny of the Russula core clade .............. 111 5. Bayesian posterior probability phylogeny of the Brevipes clade .................... 113 6. Russula altaica ................................................................................................ 117 7. Russula heterochroa ........................................................................................ 123 8. Russula laccata ................................................................................................ 128 9. Russula laevis .................................................................................................. 134 10. Russula montana ........................................................................................... 139 11. Russula nana ................................................................................................. 145 12. Russula cf. pascua ......................................................................................... 153 13. Russula purpureofusca .................................................................................. 159 14. Russula saliceticola ....................................................................................... 165 15. Russula subrubens ......................................................................................... 171 16. Scanning electron microscope photographs of the basidiospores from species in the Russula core clade and Brevipes clade ........................... 176 17. Scanning electron microscope photographs of the basidiospores from species in the Russula crown clade ....................................................... 177 18. Compound microscope photographs of basidiospores .................................. 178 19. Maps of Russula species distributions Set 1 .................................................. 179 viii LIST OF FIGURES CONTINUED Figure Page 20. Maps of Russula species distributions Set 2 .................................................. 180 21. Maps of Russula species distributions Set 3 .................................................. 181 22. Russula cf. alpigenes ..................................................................................... 203 23. Russula intermedia ........................................................................................ 207 24. Russula cf. sphagnophila ............................................................................... 211 25. Russula aff. vinosa ......................................................................................... 213 ix ABSTRACT Russula Pers. (Russulales) is an important ectomycorrhizal fungal genus in alpine and Arctic regions where it occurs in association with Salix, Betula, Dryas, and Polygonum. Despite Russula’s importance and abundance in Arctic and alpine systems there has been no in-depth systematic analysis of the genus in these habitats. This is also true for alpine areas of the Rocky Mountains where only four species of Russula have been casually reported above treeline. The genus Russula is large, diverse, and intraspecific morphological variation makes taxonomic classification difficult, which means verification using molecular techniques is necessary. This research compared Rocky Mountain alpine Russula collections to Arctic and alpine collections from Europe using an in-depth morphological study and a systematic molecular analysis of the nuc rDNA ITS1-5.8S-ITS2 region (ITS barcode) and the second largest subunit of the RNA polymerase II gene (RPB2). Over 130 Russula collections were sequenced including type material. This research confirmed eight species with intercontinental distributions in Arctic and alpine habitats, including R. nana, R. laccata, R. subrubens, R. cf. pascua, R. heterochroa, R. saliceticola, R. purpureofusca, and R. laevis. Two species are reported from subalpine habitats at treeline; R. montana with conifers and R. altaica with Betula. The Russula present in the Rocky Mountain alpine represent a subset of those known from other Arctic-alpine habitats and data show that multiple Russula species independently colonized alpine habitats. This is the first formal report of R. altaica, R. saliceticola, and R. subrubens in the Rocky Mountains and of R. heterochroa and R. purpureofusca in North America. Previous work matched sequences extracted from ectomycorrhiza in Canada to R. laevis, but this is the first work to collect this species and report it in North America. A key for the identification of alpine Russula in North America is provided. A history of Arctic and alpine mycology in North America is included and provides background material for the study. This work contributes to our knowledge of biodiversity in Arctic and alpine systems and will promote future ecological and taxonomic research on alpine Russula because little is known about these species or how to identify them. Key words: Alpine, Betula, ITS, RPB2, Salix, taxonomy, Russulaceae 1 CHAPTER ONE LITERATURE REVIEW: HISTORY OF ARCTIC AND ALPINE MYCOLOGY IN NORTH AMERICA AND INTRODUCTION TO THE GENUS RUSSULA Arctic-Alpine Biome The Arctic zone is defined as the vegetated areas between latitudes 66° and 70° N, which lies beyond the climatic limit of tree growth and south of the permanent ice sheets (Bliss 1988; Chapin & Körner 1995). The Arctic includes northern areas of Russia, Norway, Iceland, Greenland, Canada and the United States of America, which together comprise a circumpolar habitat (Callaghan et al. 2005). The alpine zone is defined as the open, vegetated areas above tree line in mountainous regions worldwide (Körner 2003). The lower limits of alpine regions can be as high as 3,500 m in equatorial regions or as low as 300 m in subpolar regions (Billings 1974). Alpine regions are disjunct habitats found at various latitudes. In North and South America, the alpine zone is a nearly contiguous system of island mountain tops stretching from the Arctic to the Subantarctic (Billings 1974; Körner 1995). Distinguishing Arctic and alpine habitats from those of subarctic and subalpine habitats is often difficult because scattered and variable vegetation can obscure the boundaries between them. Arctic and alpine systems are responsible for retaining and regulating water resources for human use (Körner 2003) and also have the ability to store large amounts of carbon making them efficient carbon sinks (Tarnocai et al. 2009). Due to severe environmental filters, large coniferous and deciduous trees are unable to grow 2 and species richness declines with increasing latitude and altitude (Walker 1995; Billings 1974). Arctic and alpine habitats are similar in that they often experience cold temperatures, high winds, and a majority of their moisture as snow fall; these factors result in a short growing season and poorly developed soils (Billings 1973; Körner 1999, 2003). However, due to difference in geographic locations, there are daily and annual differences in temperature, and in the length of the growing season; there is also considerably more vertical relief in alpine zones (Billings 1973; Chapin & Körner 1995; Körner 1999). Together Arctic and alpine regions make up 5% and 3% of land on earth, respectively (Chapin & Körner 1995; Körner 1999, 2003), and because of their similarities, they are collectively referred to as the Arctic-alpine biome (Bliss 1962, 1988; Billings 1973; Löve & Löve 1974; Chapin & Körner 1995; Murray 1995). Mounting evidence suggests that anthropogenic activity is drastically increasing the atmospheric concentration of carbon dioxide (CO2) and other greenhouse gasses (H2O, CH4, N2O, and O3) leading to increased global warming and variable changes in precipitation (IPPC 2014; Semenova et al. 2016). Research suggests that Arctic and alpine environments are being disproportionately affected by global warming compared to other systems, this is referred to as Arctic amplification (Serreze and Barry 2011; IPPC 2014). The temperature in Arctic-alpine regions is increasing 0.1° C per year, which is among the highest temperature increases recorded (Anisimov et al. 2007; Comiso and Hall 2014). Recent work also suggests that higher elevations will see increased warming (Wang et al. 2016). Another factor that has been historically underestimated, is the thawing of permafrost in Arctic systems, which is releasing large amounts of greenhouse 3 gasses. As these gasses enter the atmosphere, it will create a positive feedback loop leading to increased rates of global warming (Tarnocia et al. 2009). As the climate warms, alpine communities will be restricted to ever-decreasing areas on mountain tops or will even disappear completely (Grabherr et al. 1995). Most alpine regions are highly fragmented, disconnected, and contained within small areas of land, and the species present are often adapted to the harsh conditions found at high elevations (Anthelme and Lavergne 2018). These unique characteristics of alpine habitats have important implications for conservation of biodiversity and ecosystem functioning. A changing climate along with increased warming has the potential to drastically increase extinction rates of Arctic and alpine species. Arctic and Alpine Fungi While much is known about the flora and fauna in Arctic and alpine habitats, less is known about the funga (Dahlberg and Bültmann 2013). Fungi play important roles in nutrient cycling in Arctic environments where they exist as symbionts, decomposers, and pathogens. Fungi are estimated to be one of the most species rich groups in the Arctic, yet most research has focused on lichenized fungi, which only represent a subset of fungal diversity. With climate inevitably changing, Arctic and alpine habitats are highly threatened, which could lead to the loss of understudied fungi that plat important roles in the ecosystem. Therefore, it is important to monitor all Arctic and alpine fungi to gain a better understanding of fungal diversity and the role of fungi in ecosystem functioning (Dahlberg and Bültmann 2013). 4 Research on fungi in Arctic and alpine habitats, has been ongoing for the last two centuries, primarily in Europe. These studies have provided the basis for taxonomic classification of alpine fungi via morphological identification using macro and microscopic features derived from sporocarp surveys. Europe has a rich history of mycology that supports and encourages investigation by amateurs and professionals alike. Selected studies that provide information on fungi in Arctic and alpine habitats of Europe are outlined in TABLE 1. Table 1. Selected surveys of Arctic and alpine fungi in Europe, Asia, and Arctic Islands. Location Reference(s) Alps, Austrian Peintner 1998 Alps, French Kühner 1975; Lamoure 1982; Bon 1985; Kühner and Lamoure 1986; Bon and Cheype 1987; Bon 1991, 2000; Moreau 2002 Alps, German Schmid-Heckel 1988; Bresinsky et al. 2000 Alps, Italian Horak 1960; Bon 1987; Jamoni 1995, 2008 Alps, Swiss Favre 1955; Irlet and Rieder 1985; Senn-Irlet 1987; Graf 1994 Bulgaria Gyosheva and Dimitrova 2011 Faroe Islands Vesterholt 1998 Finland Ohenoja 2000 Iceland Christiansen 1941; Hallgrimsson 1998 Norway Lange and Skifte 1967; Gulden and Lange 1971; Gulden et al. 1985; Gulden 2005 Pyrenees Bon and Noguera 1995; Bon and Ballarà 1996; Corriol 2008 Russia Knudsen and Mukhin 1998; Niezdoiminogo 2003 Scotland Watling 1987 Slovakia Fellner and Landa 1993a, 1993b Southern Carpathians Ronikier 2008 Svalbard Gulden et al. 1985; Skifte 1989; Gulden and Torkelsen 1996 Sweden Kühner 1975 5 In contrast, there have been comparatively fewer studies of Arctic and alpine fungi in North America. This is likely due to that fact that there are fewer mycologists, Arctic and alpine habitats are remote, and a long mycological history is lacking. Arctic and alpine areas of North America are distributed across a large expansive landscape and many areas, particularly in the Canadian Arctic and the northern Rocky Mountains, are remote and inaccessible. Despite these limitations, there is some early information on the fungi present in Greenland (Borgen et al. 2006), Northern Canada (Ohenoja and Ohenoja 2010), Alaska (Laursen and Ammirati 1982a; Miller 1987, 1993, 1998), and the central and southern Rocky Mountains (Cripps and Horak 2008). In general, knowledge of fungal diversity in North America is much less complete than in Arctic-alpine Europe and particular Arctic islands. The History of Arctic and Alpine Mycology in North America Several papers have briefly summarized the history of Arctic mycology in North America (Kobayasi et al. 1967; Hutchison et al. 1988) and a few have provided detailed accounts of the mycological history of Canada, including Arctic regions (Savile 1963; Estey 1994; Redhead and Baillargeon 1999; Väre 2017). However, no historical review has focused on alpine fungi in North America and no comprehensive history of mycological studies in Arctic and alpine regions of North America has been compiled. As a contribution to the field, this section provides a historical overview of Arctic and alpine mycology in North America. Although Greenland is considered part of the North American continent, Arctic and alpine fungi on Greenland has been extensively studied (Lange 1957; Peterson 1977; Knudsen and Borgen 1982; Borgen 2006; Borgen et al. 6 2006) and summarized in Arctic and Alpine Mycology 6 (Boertmann and Knudsen 2006) and therefore, this island is excluded from this work. Early Expeditions to the Canadian Arctic (1819-1940) During the nineteenth century, Arctic expeditions were exploring the northern polar regions of North America. These expeditions were funded by wealthy business men or supported by government agencies (Berkeley 1878; Rostrup and Simmons 1906; Lind 1910; Dearness 1923; Nares 2011) and included crew members with experience in sailing, Arctic travel, craftsmanship, engineering, and a variety of sciences. Arctic expeditions were often exciting, dangerous, and only for adventurous types as members often died during these expeditions, which could last for up to five years. Botanical collections were of primary interest to scientifically-minded individuals on these early expeditions. Nicholas Polunin summarizes the various accounts of these collectors in Botany of the Canadian Eastern Arctic, parts I and II (Polunin 1940, 1947). Historically, mycologists began investigating Arctic fungi by examining the dried vascular plant material brought back by botanists for attached saprophytic or parasitic fungi (Linder 1947; Savile 1963). Occasionally, collectors would bring back large fleshy fungi; however, these were usually not identifiable due to improper handling and storage (Linder 1947; Savile 1963). Starting in 1819, Captain William Edward Parry led two ships through the Canadian Archipelago on what would later be deemed the most successful attempt to find the Northwest Passage. The ships wintered at Melville Island and in 1820 officers of the voyage collected Cantharellus lobatus (Arrhenia lobata), Lycoperdon pratense, and 7 fifteen species of lichen on the island (Brown 1823). According to Redhead and Baillargeon (1999) these two collections are the earliest published record of any agaric in Canada. Dr. John Richardson, a surgeon on Captain Sir John Franklin’s second overland expedition (1825–1827), was the next to collect Arctic fungi; this expedition through the Canadian Arctic crossed the Northwest Territories and reached the Arctic Ocean (Berkeley 1839; Väre 2017). Richardson’s collections were placed in the prestigious herbarium of Sir William Jackson Hooker who was later appointed director of the Herbarium at Kew in 1841 (Desmond 1995). The names and descriptions of theses fungi are found throughout Rev. Miles Joseph Berkeley’s paper published on exotic fungi (Berkeley 1839). Berkeley, an English clergyman with a strong interest in cryptograms and mycology, is considered one of the founders of plant pathology (Massee 1913) and was one of the first mycologists to study fungal material from the Canadian Arctic. Captain Franklin set out again in 1845 with two ships, the HMS Erebus and the HMS Terror, in an attempt to traverse some of the last unknown sections of the Northwest Passage. This final voyage would end tragically with the deaths of 129 men, including Franklin’s, and the loss of both ships (Neatby and Mercer 2008). Franklin’s widow, Lady Jane Franklin, would go on to finance five expeditions in search of her husband’s lost remains. Interestingly, Arctic fungi were collected on two of these expeditions. The one led by Admiral Belcher in 1852 set out with at least four ships and only one returned; it is noteworthy that, Dr. David Lyall, a physician and naturalist who was with the returning expedition, collected Arrhenia lobata (again recorded as Cantharellus lobatus) in 1853 at Wellington Channel (Polunin 1940). Redhead (1984) 8 noted that Lyall’s collection of A. lobata would be the first confirmed collection (based on preserved material) of the fungus from North America and it was placed in the Herbarium at Kew. The fifth and final expedition that set out to search for Franklin’s party, in 1857, was led by Sir Francis Leopold M’Clintock in the Yacht ‘Fox’, a boat purchased by Lady Franklin (M’Clintock 1860). M’Clintock’s expedition included Dr. David Walker, surgeon and naturalist, who was the next documented explorer to collect Arctic fungi. Walker focused primarily on botanical collections; however, he also collected five basidiomycete fungi in the genus Agaricus (agarics) and one ascomycete (Hooker 1860). Walker was not well versed in botany; therefore, all 170 collections were given to Sir Joseph Dalton Hooker, William Hooker’s son who succeeded him as director at Kew (Desmond 1995), for determination. Hooker then passed all of the fungi to Berkeley for review (Hooker 1860). Berkeley continued to contribute to Arctic mycological investigations when he published on 26 species (23 species from Arctic Canada and three from Greenland) of Basidiomycota and Ascomycota collected during the Arctic Expedition from 1875–1876 (Berkeley 1878). The study of Arctic fungi continued with the Harriman Alaska Expedition of 1899 when William Trelease and other botanists collected Arctic fungi from Alaska (Trelease 1904). In 1904 Pier A. Saccardo, Charles H. Peck, and W. Trelease published a list of all known fungi in Alaska. Their list contained the names of fungi collected from the Harriman Alaska Expedition, from which, Saccardo and Scalia describe three new species of Arctic micromycetes. The list also includes four other Arctic species, all of which were collected from Point Barrow (Saccardo et al. 1904). In 1906, Emil Rostrup 9 began careful examination of botanical collections made by Herman George Simmons during the second voyage of the Fram Expedition (1898–1902). These fungi were collected from Ellesmere Island in Northern Canada and included macromycetes and micromycetes, although most were saprotrophs associated with phanerogamous vegetation; eight new species were described (Rostrup and Simmons 1906). The Gjoa expedition under Captain Ronold Amvindsen from 1904–1906 visited King Point on the Yukon Coast and King William Island. Fungal collections extracted from plants were observed by Jens Lind (1910) who noted the difficulty of identifying fungi that were deformed by the cold temperatures and exposed habitats of the Arctic, a problem still facing mycologists today. Lind would continue researching Arctic micromycetes throughout the early 20th century. He examined herbarium material of Arctic vascular plants and produced detailed publications focused on the parasitic and saprophytic fungi present; including their ecology, hosts, dispersal, geographic distribution and diversity (Lind 1927, 1934). In 1909, agrostologist A. S. Hitchcock traveled through the Alaskan interior looking at various grasses. He collected Arctic uredineous rust fungi Near Nome, which he shared with J. C. Arthur (Arthur 1911). In 1923, John Dearness of London, Ontario, examined over a hundred fungi collected by the naturalists of the Southern Party from the Canadian Arctic Expeditions (Dearness 1923). He noted the wide host range and relative lack of parasitic fungi, only recording three rusts and one smut. However, Savile (1963) explained these low numbers by pointing out that botanists avoid diseased plants in order to secure the best collections, leading early mycologists to underestimate parasitic fungal abundance in Arctic 10 environments. Dearness later returned to the Basidiomycota after more than a half century focusing on the fleshy fungi of southern Baffin Island (Dearness 1928). One of the last true Arctic Expeditions was the Oxford Exploration Club’s 1931 expedition to Akpatok Island, after which Nicholas Polunin (1934) published a list of the 24 fungi he collected. The macromycetes collected during the eight major arctic expeditions outlined above are listed in TABLE 2. Overall, research from these early expeditions found relatively few fungi in multiple genera and with a variety of ecological roles. Most if not all of these fungi were originally described from Europe and were similar to the fungi found in previously studied Arctic regions. These studies provide a base for future mycologist to begin exploring fungal diversity in Arctic regions of North America. With a majority of Arctic North America mapped out and explored, expeditions began tapering off; however, detailed investigations of the fungi in these regions were just beginning. Table 2. Macromycetes collected during early Arctic expeditions from 1819 though 1931. Names and authorities as reported. Expedition Species Reported Location Citation Captain William Basidiomycota: Cantharellus lobatus Peck and Lycoperdon Melville Island, Brown Edward Parry's pratense Pers. Canada Arctic 1823 Expedition (1819- Archipelago 1820) Captain Sir John Basidiomycota: Cantharellus canadensis KL. MSS., Northwest Berkeley Franklin's Second Favolus (Pleuropus) humboldtii Berk., Favolus Territories and 1839 overland expedition (Pleuropus) hepaticus Kl., and Favolus (Pleuropus) Arctic regions in (1825-1827) canadensis Kl. l. c. Canada Arctic Expedition led Basidomycota: Cantharellus lobatus Peck Wellington Polunin by Admiral Belcher Channel, 1940 in (1852-1854) Nunavut, Canada Sir Francis Leopold Basidiomycota: Agaricus allosporus Berk., Agaricus Boothia Hooker M'Clintock's cyathiformis Bull., Agaricus furfuraceus P., Agaricus Peninsula, 1860 Expedition in the vaginatus Bull., and Agaricus umbelliferus L. Nunavut, Yacht 'Fox' (1857- Canada 1859) Arctic Expedition Ascomycota: Peziza stercorea P. and Urnula hartii B. Northern Berkeley from 1875-1876 Basidiomycota: Agaricus (Clitopilus) undatus Fr., Nunavut, 1878 Agaricus (Naucoria) bellotianus B., Agaricus (Omphalia) Canada 11 umbelliferus L., Agaricus (Omphalia) umbilicatus Schaeff., Agaricus (Stropharia) feildeni B., Agaricus (Tubaria) furfuraceus P., Agaricus (Tubaria) pellucidus Bull., Cantharellus muscigenus Fr., Hygrophorus miniatus Fr., Hygrophorus virgineus Fr., Lycoperdon atropurpureum Vitt., Lycoperdon cretaceum B., Merulius aurantiacus Fr., and Russula integra Fr. Second voyage of the Ascomycota: Mollisia graminis (Desm.)., Niptera Ellesmere Rostrup Fram Expedition melatephra (Lasch.)., and Sclerotinia vahliana Rostr. Island, Nunavut, and (1898-1902) Basidiomycota: Cantharellus lobatus (Pers.)., Collybia Canada Simmons dryophila (Bull)., Galera hypnorum (Batsch)., Hebeloma 1906 fastibilis (Fr.)., Lycoperdon gemmatum (Batsch)., Mycena pumila (Bull)., Naucoria festiva (Fr.)., Naucoria melinoides (Fr.)., Naucoria nimbosa (Fr.)., Omphalia umbellifera (L.)., Psalliota campestris (L.)., Psalliota rodmani (Peck)., Psathyrella polaris n. sp., Russulina lutea (Huds.)., and Tricholoma caelatum (Fr.). Canadian Arctic Ascomycota: Peziza micropus Pers. var. flavida Phil., Northern Dearness Expedition (1913- Scleroderris fuliginosa (Fr.) Karst., and Scutellina Regions of 1923 1918) scutellata (L.). Basidiomycota: Boletus scaber Fr., Nunavut, Cantharellus muscigenus Fr., Calvatia creatacea (Berk.) Northwest Lloyd, Galera hypnorum Batsch., Hebeloma fastibile Fr., Territories, and Hygrophorus cantharellus Fr., Hygrophorus sp., Inocybe the Yukon, flocculosa Berk., Lycoperdon umbrinum Pers., Naucoria Canada sp., Omphalla umbellifera Fr., and Russula sp. Oxford Exploration Ascomycota: Helvella lacunosa Afz., Peziza acetabulum Akpatok Island, Polunin Club's 1931 L., and Sphaerospora asperior (Nyl.) Sacc. Basidiomycota: Nunavut, 1934 Expedition to Boletus reticulatus (Schaeff.) Boud., Boletus sp., Canada Akpatok Island Cantharellus cibarius Fr., Cortinarius spp., Dictyolus muscigena (Bull.) Quél., Hygrophorus sp., Hypholoma sp., Inocybe violacea Pat., Inocybe spp., Laccaria laccata (Scop.) B. & Br., Lactarius vellereus Fr., Lycoperdon perlatum Pers., Lycoperdon sp., Mycena polygramma Quél., Mycena spp., Omphalia sp., Psilocybe semilanceata Fr., Russula emetica Fr., Russula ochroleuca Fr., and Russula sp. Detailed Morphological Studies Detailed mycological work that focused on Arctic fungi in North America includes researchers who focused on micromycetes, fungi that produce small, inconspicuous fruiting bodies, or researchers who focused on macrofungi, which produce large, conspicuous fruiting bodies. Usually mycologist focused their studies on 12 micromycetes or macrofungi, with little overlap between the two. Therefore, Arctic micromycetes and macrofungi are separated here to provide an overview of each group. Arctic Micromycetes. Margaret Newton and J. P. Anderson undertook the first detailed morphological studies of cold-climate micromycetes in 1940. Their work expanded our understanding of plant parasitic fungi, mainly rusts, in the Northwest Territories and Northern Alaska (Estey 1994; Anderson 1940). Shortly after, in 1953, Edith K. Cash compiled a list of all known fungi and Myxomycetes in Alaska, which consisted of 843 species names that included plant parasites, saprobic micromycetes, and Basidomycetes collected in the northern Arctic regions of the state (Cash 1953). As research into Arctic fungi began ramping up in the 1960s, Douglas Barton Osborne Savile, a prominent Canadian mycologist, focused on understanding the diversity of parasitic fungi in the Canadian Arctic (Savile 1961, 1963). His work highlighted the evolutionary history of these organisms by connecting plant parasitic fungi like Puccinia and Ustilago to their hosts, which contributed significantly to the taxonomic understanding of these groups. Savile’s research would later be praised for initiating detailed mycological investigations into fungi of the Canadian Arctic (Kobayasi et al. 1967). Savile went on to collaborate with John A. Parmelee on parasitic micromycetes of Queen Elizabeth Island (Savile and Parmelee 1964) and later wrote a review of Arctic fungi (Savile 1972). Parmelee continued to describe micromycetes in the Canadian Arctic (Parmelee 1968), which further increased our understanding of the fungi on Central Baffin Island (Parmelee 1969) and in the Yukon (Parmelee and Ginns 1986). Parmelee’s research on Arctic fungi culminated with his 1989 paper focused on the Uredinales (now 13 Pucciniales) of Arctic Canada, which contained excellent figures and a key to the order and all major genera (Parmelee 1989). Dr. Margaret Barr was the next mycologist to study micromycetes in the Canadian Arctic. She published a landmark paper on Pyrenomycetes that provided taxonomic clarification for the group and simplified identification (Barr 1959). Next, William Bridge Cooke surveyed soil and water for fungi in the Alaskan tundra near the Eskimo village of Napaskiak (Cooke and Fournelle 1960). Shortly after, Wehmeyer (1961) published a monograph of Pleospora and its allies, which included Arctic fungi. Japanese mycologist Yosio Kobayasi yearned to visit the biologically diverse Arctic after a trip to Antarctica in 1963. In 1965, his opportunity arose and Kobayasi organized a small party to perform field work in the Alaskan Arctic. Kobayasi’s team arrived in Point Barrow, Alaska on August 1, 1965 and began three weeks of exploration and experimentation at various sites in the region. During their research, they isolated fungi from soil, dung, water (water molds), plant material, animal bones, and insects (in an unsuccessful search for Trichomycetes). Kobayasi’s team estimated that over 450 taxa of micromycetes were present in the Alaskan Arctic (Kobayasi et al. 1967). In all, the party collected 230 fungi representing 61 families and 136 genera; six species were new to science. Just as Lind (1910) noted, Kobayasi’s party recognized that the harsh environmental factors and corresponding altered fungal appearance complicated identification. Arctic Macrofungi. Up until the 1950’s little research had focused on the larger fruiting bodies produced by the Ascomycota and the Basidiomycota in the Arctic, 14 otherwise known as macrofungi. In 1908 Elias J. Durand first confirmed Geoglossaceae in Arctic and subarctic regions of North America when he identified Mitrula gracilis in Labrador and Newfoundland (Durand 1908); others would also study Geoglossaceae in Arctic regions (Mains 1955; Kankainen 1969). Durand noted that M. gracilis also had been collected in Greenland (Durand 1908) and appeared to have a large distribution in Arctic regions. Hugh S. Spence and O. E. Jennings investigated the agarics of the Northwest Territories and on Southampton Island, respectively, and found species in the genera Boletus, Calvatia, Cortinarius, Hydnum, Hygrophorus, Lactarius, Psathyrella, Russula, and Tubaria (Spence 1932; Jennings 1936). David H. Linder (1947) summarized the current knowledge regarding fungi found in the Canadian eastern Arctic, coming to some conclusions that were ahead of his time. Linder recognized that fungi found in Arctic habitats appeared to have circumpolar distributions and shared similarities with their alpine counterparts at lower latitudes. He also noted the parallels between distributions of Arctic-alpine fungi and flowering plants, indicating that the study of fungi may substantiate theories concerning distributions of phaneograms. During the 1950’s several researchers began focusing on fungi in Northern Canada. James Walton Groves and colleagues published a series of articles starting with the Hypocreales and Discomycetes (Groves and Hoare 1954), followed by one on the Boletaceae (Groves and Thomson 1955), and lastly with a paper focused on the Amanitaceae, Hygrophoraceae, Rhodophyllaceae, and Paxillaceae (Groves et al. 1958). Groves research continued into the 1960s when he published a paper focused on the Gasteromycetes of Northern Canada with Constance A. Bowerman (Bowerman and 15 Groves 1962). Groves final paper, published one year after his death in 1970 in collaboration with M. E. Elliott, was on the Discomycetes of Northern Canada (Groves and Elliott 1971). During the same period, Howard E. Bigelow studied collections secured by government biological survey parties, which included some Arctic collections in the Tricholomataceae (Bigelow 1959). Bigelow also contributed to our knowledge of Arctic Omphalina in Alaska and Canada (Bigelow 1970). Towards the end of the 1960s, Orson K. Miller Jr., a mycologist trained under Alexander Smith, became interested in Arctic fungi after a visit to Alaska with Robert L. Gilbertson in 1967 and 1968 (Cripps 2010). Miller’s interest in Arctic fungi would last for the next 30 years, and Miller would inspire several mycologists to examine the understudied fungi in these remote areas. Miller published three papers focused on gasteromycetes from the Yukon territory and adjacent Alaska; he was considered an expert on this group by his peers (Miller 1968, 1969; Miller et al. 1980). Miller then turned his attention to Arctic and subarctic Basidiomycota in Canada and Alaska publishing on Coprinus with Roy Watling (Watling and Miller 1971), the genera Omphalina, Laccaria, and Coprinus with David F. Farr (Farr and Miller 1972), and on Melanoleuca with Linnea S. Gillman (Gillman and Miller 1977). Miller had several students who focused on the study of Arctic fungi. Robert K. Antibus studied fungal ecology in the Arctic and two other Miller students, Gary Laursen and T. G. Rau, who focused on soil fungi and fungal decomposition respectively in the Alaskan tundra (Laursen 1975; Rau 1977). Miller went on to publish a series of papers with Laursen, first on Arctic and alpine agarics in Alaska and Canada (Miller et al. 1973; Laursen et al. 16 1976); then on fungal hyphae belowground-biomass and distribution in Alaska (Miller and Laursen 1974; Laursen and Miller 1977); and lastly, on the function, distribution, and known plant associates of mycorrhizal fungi of the Alaskan tundra (Miller and Laursen 1978). Laursen would focus a large part of his early career on Arctic fungi, collaborating with several other mycologists. Laursen and Harold H. Burdsall Jr. found Geopora cooperi in association with Salix alaxensis in the Alaskan tundra, expanding its distribution to include Arctic regions (Laursen and Burdsall 1976). In the 1980s, Laursen and Joe Ammirati studied Lactarius, Cortinarius, and Hygrophoraceae in the Alaskan Arctic tundra (Laursen and Ammirati 1982a; Ammirati and Laursen 1982; Laursen et al. 1987a). Miller continued publishing heavily on Arctic fungi into the ‘80s and ‘90s and this includes a review of the current taxonomic understanding of Arctic fungi at Point Barrow, AK (Bunnell et al. 1980). Miller studied higher fungi in the subarctic tundra of Alaska and the Yukon (Miller 1982a; Miller 1987), Marasmius epidryas with Canadian Mycologist Scott Redhead (Redhead et al. 1982), Phaeogalera and Galerina with prominent European mycologist Egon Horak (Horak and Miller 1992), Cystoderma (Miller 1993), and Hebeloma (Miller 1998). Miller’s in-depth knowledge of Arctic fungi provided him with the basis to recognize the large intercontinental distribution of some fungal species (Miller et al. 1982). Miller continued to publish into the early 2000s when he focused on three Hebeloma species in the alpine tundra of Colorado, recognizing that these species were also present in the Arctic tundra in Europe (Miller and Evenson 2001). 17 While Miller and colleagues were exploring the Alaskan Tundra, Scott Redhead provided clarification for the Arctic species Gerronema pseudogrisella (Redhead 1980), published a detailed overview of Arrhenia in Arctic North America (Redhead 1984), and briefly recounted early agaricology for each Canadian Territory, which included information on early Arctic fungal collections (Redhead and Baillargeon 1999). Redhead (1989) also focused on the biogeographical patterns of Canadian fungi, he noted that fungal species found in the high Arctic and in Arctic-alpine habitats have intercontinental distributions that match Arctic and alpine floristic patterns. Around the same time, Hutchinson, Summerbell, and Malloch (1988), who were intensively studying the fungi of Quebec, noticed that a majority of the fungi they collected in Northern Quebec were also present in Arctic regions of Greenland and Northern Europe (Gulden et al. 1985; Bresinsky 1987; Watling 1987). The collections of Durand (1908), Miller et al. (1982), Redhead (1984), and Hutchinson et al. (1988) further support the hypothesis that a majority of Arctic fungi have large intercontinental distributions with disjunct distributions in the alpine, an idea that would continue to dominate Arctic and alpine mycological and botanical research. Several European mycologists have studied Arctic-alpine fungi in North America. H. Heikkila and P. Kallio from Finland showed that Omphalina is one of the most common genera in Arctic-alpine habitats of Northern Canada (Heikkila and Kallio 1966, 1969) and Kallio (1980) surveyed the subarctic fungi of Schefferville, noting similarities to Finnish species. Seppo Huhtinen (1982, 1985) reported on the Pezizales and Helotiales in Northern Labrador and Quebec, finding 16 new species in North America. Esteri 18 Ohenoja and Martti Ohenoja focused on the macromycetes in Arctic Canada during trips to the Hudson Bay in 1971 and 1974 (N.W.T. and Manitoba). Working independently and in collaboration with various authors they studied Lactarius (Ohenoja and Ohenoja 1993), Inocybe (Ohenoja et al. 1998), Marasmius epidryas (Redhead et al. 1982), Ascomycetes in Finland noting some distributions extending into North America (Ohenoja 1975), and the fungal diversity at Rankin Inlet (Ohenoja 1972). Ohenoja and Ohenoja (2010) then summarized all of their findings in the Canadian Arctic. Prominent Norwegian Mycologist Gro Gulden also contributed knowledge on Arctic and alpine species of Lepista from Alaska (Gulden 1983) and Galerina from Schafferville (Noordeloos and Gulden 1992). Arctic and alpine Fungal Ecology. In the 1960s, research was also expanding to include ecological investigations in Arctic regions. Sprague and Lawrence (1959, 1959– 1960, 1960) published a three-part series with the goal of understanding the effects of deglaciation on pedological, botanical, and fungal development. Flanagan and Scarborough (1973, 1974) investigated fungal decomposition and Laursen and Chmielewski (1982) focused on the ecological significance of mutualistic and saprophytic soil fungi, all in the Arctic Tundra. During this same time mycorrhizal fungi started getting the attention of mycologists in Arctic and alpine regions (See Ectomycorrhizal Fungi and Hosts in Arctic-alpine environments). A. E. Linkins and Robert K. Antibus studied the ectomycorrhizal fungi of Salix rotundifolia following oil contamination of the soil. They found reduced numbers of viable mycorrhizae, an altered fungal mantle (Antibus and Linkins 1978), decreased root respiration rates, and reduced 19 adaptation to cold temperatures (Linkins and Antibus 1978); however, these effects were less prominent at a natural oil seep in Cape Simpson. Antibus and colleagues went on to investigate ectomycorrhizal synthesis of local and non-native fungi with Salix rotundifolia in Alaska (Antibus et al. 1981). Orson K. Miller Jr. also studied fungal biomass in Alaska in natural (Miller 1982b) and oil contaminated soils (Miller et al. 1978). The mycorrhizal status of plants in the Arctic Tundra and in alpine regions of Montana and Wyoming were assessed by Linkins and Antibus (1982) and Lesica and Antibus (1986a, 1986b). Antibus later reported on the physiology of ectomycorrhizal fungi from his earlier research, focusing on the effects of temperature, at Point Barrow (Antibus 2010). Shortly after the turn of the century, Kernaghan and Harper (2001) began assessing edaphic factors, i.e. how ecological habitat change from subalpine to alpine affected ectomycorrhizal diversity in the Canadian Rockies. They reported that in alpine areas of the Front Range of the Rocky Mountains, fungi appeared to be non-host specific, forming mycorrhizal associations predominantly with conifers below treeline and with angiosperms like Salix and Dryas above treeline. They hypothesized that these interactions would facilitate upward movement of treeline due to pre-existing mycorrhizal fungi. The research of Schadt et al. (2003) marked a significant shift in the theoretical understanding of the biogeochemical processes in the alpine tundra of Colorado. Their research showed evidence of microbial communities, primarily consisting of fungi, maintaining significant biological activity under snow-cover, this 20 finding challenged our previous understanding of biogeochemical cycling in cold environments. The International Symposium on Arctic and Alpine Mycology (1980 – 2016) On August 16th, 1980, during the most productive fruiting period for Arctic and alpine fungi, an esteemed group of 25 mycologists from 9 countries met at the Naval Arctic Research station in Point Barrow, Alaska. The First International Symposium on Arcto-Alpine Mycology (ISAM) was organized by acting president Gary Laursen. Many pertinent Arctic and alpine mycologists were in attendance including Savile, Miller, Kobayasi, Moser, Lamoure, Lange, Knudsen, and Horak. The symposium’s goal was to better understand fungi in Arctic and alpine systems and the week-long meeting was an overwhelming success, providing detailed knowledge of the fungal community at Point Barrow (Laursen and Ammirati 1982a). Mycologists in attendance were required to submit a paper focusing on Arctic and/or alpine fungi for the proceedings. The format of the first meeting consisted of several days collecting fungi in the field followed by evening lectures to discuss proposed contributions. Because of the relative lack of knowledge on Arctic-alpine fungi, the first ISAM, focused on determining the fungi present. Fungal taxonomy would continue to dominate all subsequent meetings. After the first ISAM, it was decided that a select group of professional Arctic and alpine mycologists would meet every four years on varying continents to advance the knowledge of Arctic and alpine fungi. At the end of each symposium a representative would be selected to organize and act as sitting president for the next meeting. The format of the first ISAM has persisted through all ten symposia conducted thus far and 21 more that 100 different researchers have been involved in ISAM since its inception (Gulden and Høiland 2008, Cripps and Ammirati 2010). Each ISAM was held at an iconic Arctic or alpine location and all have improved our understanding of fungi in these environments. TABLE 3 summarizes the research published in each proceeding that contributes information on Arctic and alpine mycology in North America. Table 3. Research published in the proceedings of the ten International Symposia on Arctic and alpine mycology focused on North American fungi. Symposium Research on Arctic and alpine mycology in North Acting Editor(s) America President 1st ISAM, Basidiomycetes in subarctic Alaska (Miller 1982a), Gary Laursen Laursen and August 1980. Cortinarius in Alaska (Ammirati and Laursen 1982), Ammirati Barrow, Alaska, Lactarius in Alaska (Laursen and Ammirati 1982b), (1982a) U.S.A. Lower fungi in the Arctic (Kobayasi 1982), Soil fungi in Arctic tundra (Laursen and Chmielewski 1982), and Mycorrhizal associations with Salix rotundifolia (Linkins and Antibus 1982) 2nd ISAM, Basidiomycetes in the Rocky Mountian alpine (Moser Egon Horak Laursen et al. August 1984. and McKnight 1987), Cortinariaceae in the Yukon (1987b) Swiss National Territory of Alaska (Miller 1987), and Hygrophoraceae in Park, Switzerland the Arctic-alpine tundra of Alaska (Laursen et al. 1987a) 3rd ISAM, August 1988. The 3rd and 4th ISAM where combined into one Sigmund Petrini and Svalbard proceeding and the following research related to Arctic Sivertsen Laursen (1993) and alpine mycology in North America was included: 4th ISAM, Cystoderma in Alaska (Miller 1993), Lactarius in Arctic August 1992. Canada (Ohenoja and Ohenoja 1993), and Myxomycetes Denise Petrini and Lanslebourg, in the Alaskan tundra (Stephenson and Laursen 1993) Lamoure Laursen (1993) France 5th ISAM, Hebeloma from Alaska (Miller 1998), Inocybe species Viktor Mukhin Mukhin and August 1996. from Arctic Canada (Ohenoja et al. 1998) Knudsen Polar Urals, (1998) Russia 6th ISAM, Arrhenia auriscalpium in Colorado (Cripps and Horak Torbjorn Boertmann and August 2000. 2006) and Agrocybe praemagna in the Rocky Mountain Borgen Knudsen Greenland alpine region (Horak and Moser 2006) (2006) 7th ISAM, Cortinarius in the Rocky Mountain alpine (Peintner Gro Gulden Gulden and August 2005. 2008) and A checklist of agarics from the Rocky Høiland (2008) Finse, Norway Mountain alpine zone (Cripps and Horak 2008) 22 8th ISAM, Amanita found on the Beartooth Plateau (Cripps and Cathy Cripps Cripps and August 2008. Horak 2010), Marasmius epidryas in the Rocky Mountain Ammirati Beartooth alpine (Ronikier and Ronikier 2010), Hebeloma hiemale (2010) Plateau, in the Rocky Mountain alpine (Becker et al. 2010), Montana/Wyomi Larger fungi in Arctic and subarctic Canada (Ohenoja and ng, U.S.A. Ohenoja 2010), Inocybe (Mallocybe) in the Rocky Mountian alpine (Cripps et al. 2010), Lycoperdaceae on the Beartooth Plateau (Kasuya 2010, Jalink 2010), and Mollisoid Ascomycetes on the Beartooth Plateau (Nauta 2010) 9th ISAM, Inocybe leiocephala, a species with an intercontinental Esteri Ohenoja Ohenoja et al. August 2012. distribution (Larsson et al. 2014) and Lactarius from the (2018) Utsjoki, Finland Rocky Mountain alpine zone (Cripps and Barge 2013) 10th ISAM, Inocybe species distributed in the North American Alpine Tamotsu Hoshino et al. August 2016. (Larsson et al. 2018) Hoshino (2018) Kanazawa, Japan Fungi in the Rocky Mountain Alpine Zone Many mycologists have focused their attention on Arctic fungi in Alaska and Canada over the last century; however, little attention was paid to the alpine fungi in the mountainous regions of the continent. At the beginning of the 20th century, researchers began reporting macromycetes from the Rockies (Overholtz 1919; Kauffman 1921; Seaver and Shope 1930; Solheim 1949); however, no research on true alpine fungi above treeline occurred until the 1980s when Meinhard Moser made a few brief visits to the North American alpine. Moser made several collecting trips to the Greater Yellowstone Area, which included alpine regions of Yellowstone National Park, the Beartooth Plateau in Montana and Wyoming, and the Windriver Mountains of Wyoming. Moser primarily described Cortinarius from these regions but also reported Russula nana for the first time from the Rocky Mountain alpine (Moser and McKnight 1987; Moser 1993; Moser et al. 1994, 1995). Redhead also reported Arrhenia lobata from Colorado after examining several of Alexander H. Smith’s collections from the Rocky Mountain alpine; A. lobata is 23 a common Arctic-alpine species known from the Alps, Greenland, and Iceland (Redhead 1984). Research into the diversity of lichens in the Rocky Mountain alpine started much earlier than research into the macro or micromycetes of the region. William A. Weber (1955) and Henry A. Imshaug (1957) provided historical accounts of lichenology in the Rocky Mountain alpine, starting as early as 1861. Imshaug also compiled a list of all the lichens present in alpine regions of the western United States and Canada, provided keys to alpine lichen identification and maps of their distribution (Imshaug 1957). Weber, a trained botanist who became obsessed with lichens in 1951, published a book on lichens of the Rocky Mountains that included alpine collections from the central and southern regions of the range (Corbridge and Weber 1998). Other researchers were also exploring the diversity of lichens in the Rockies; Eversman (1995) compiled a list of over 300 species of lichens in the alpine of the Beartooth Plateau and Erik D. Ahl’s Master’s Thesis focused on understanding how lichen diversity and abundance changes from alpine to subalpine sites at Rocky Mountain National Park in Colorado (Ahl 2016). With the start of the new millennium, research into the fungi present in Alpine regions of North America, and especially the southern Rocky Mountains, would increase substantially. In 1996 Monique Gardes and Anders Dahlberg wrote a review on the diversity of mycorrhizal fungi in Arctic and alpine regions. They focused on the major mycorrhizal relationships in these systems including arbuscular mycorrhizae, dark septate fungi, ectomycorrhizae, and ericoid associations. The occurrence of these mycorrhizal types is highly variable and certain Arctic-alpine plants lack mycorrhizal associations completely 24 (Gardes and Dahlberg 1996). Gardes and Dahlberg (1996) identified mycorrhizal associations in cold-dominated environments as a potential model for understanding the evolution of mycorrhizal symbioses. They outlined future research questions requiring investigation, focused on the use of molecular tools to answer questions about the fungal species present, and mycorrhizal community structure and dynamics. Gardes and Dalberg’s review set the stage for future research into mycorrhizal fungi in Arctic and alpine areas. In 1999 a survey of alpine fungi in the Rocky Mountains was initiated by Cathy Cripps and Horak with funding from a National Science Foundation Grant. Cripps is a professor at Montana State University in Bozeman and another former student of Miller’s who was interested in Arctic and alpine fungi. Cripps and Horak were the first to survey alpine fungi in the Rocky Mountains on a large scale and their work focused on the central and southern Rockies. Later that same year a preliminary report on the agarics found in the Rocky Mountain alpine zone was presented at the International Botanical Congress (Cripps and Horak 1999). Cripps and Horak’s investigation of the alpine fungi continued throughout the early 2000s and information on the distribution of ectomycorrhizal genera and their associated plant hosts in the Rocky Mountain alpine was disseminated at various conferences; genera discussed included: Amanita, Inocybe, Russula, Lactarius, and Hebeloma (Cripps and Horak 2002, 2005, 2007; Cripps 2003; Cripps et al. 2008). A review contributed information on the mycorrhizal status of alpine plants, including those of the Beartooth Plateau (Cripps and Eddington 2005). This review reported that 68% of alpine vascular plant species form mycorrhizal associations. 25 These associations included ectomycorrhizae, arbuscular mycorrhizae (AM), ericoid fungi, and arbutoid mycorrhizae; arbuscular mycorrhizal plant-fungal associations were the most common. Previously, Lesica and Antibus (1986a, 1986b) had only assessed arbuscular mycorrhizal root colonization on fell-field plant communities and on hemiparasitic vascular plants in alpine regions of the Rocky Mountains. Cripps and Horak (2006) reported the well-known Arctic-alpine species Arrhenia auriscalpium from Colorado; this species is known for its intercontinental distribution, and they reported the highest elevation and the furthest latitude south recorded for the fungus. In 2008 Cripps and Horak published a preliminary list of alpine fungi in the Rocky Mountains in the proceedings of the seventh ISAM (Cripps and Horak 2008). At this date, their research revealed over 165 species from 46 genera and 11 families. They estimated that at least 75% of the macromycetes present in the Rocky Mountain alpine are known from other Arctic-alpine environments; the remaining 25% could be endemic. Based on these findings, the most diverse mycorrhizal groups in the Rocky Mountain alpine zone are the Cortinariaceae, Inocybaceae, and Hydnangiales with over 74 species. They hypothesized that the diverse geology, habitat, and mesic conditions of the southern Rockies led to more variation in habitat and thus greater fungal diversity than observed further north on the Beartooth Plateau (Cripps and Horak 2008). Around this time, molecular DNA methods and phylogenetic analysis became cheaper and easier to use, providing researchers with a powerful and independent way to verify taxonomic determinations based on morphology. Armed with large molecular data sets and a basic understanding of the fungal diversity mycologists began investigating the 26 fungi in the Rocky Mountain alpine zone, focusing primarily on ectomycorrhizal genera. Cripps student Todd Osmundson investigated Laccaria in the Rockies (Osmundson et al. 2005). He utilized ribosomal DNA from the ITS region, along with morphological and cultural data; systematic analysis revealed five species in the Rocky Mountain alpine zone. Laccaria laccata var. pallidifolia and L. nobilis were reported for the first time in Arctic-alpine habitats and L. pseudomontana was new to science. Another Cripps student, Ed Barge, focused on the distribution and morphology of the genus Lactarius (Russulales, Russulaceae) in the Rocky Mountain alpine zone. Together they confirmed six species in the Rocky Mountain alpine zone, one new to science, found that most species were intercontinentally distributed in Arctic-alpine regions, and that species distribution may have been shaped by glaciation, host ranges, and long-distance dispersal (Barge et al. 2016; Barge and Cripps 2016). Additional molecular and morphological analyses have investigated the ectomycorrhizal genera Hebeloma (Beker et al. 2010; Cripps et al. 2019a), Cortinarius (Peintner 2008), and Inocybe (Cripps et al. 2010; Larsson et al. 2014; Larsson et al. 2018; Cripps et al. 2019b) in the Rocky Mountain alpine. All of the molecular studies listed above have confirmed intercontinental distributions and disjunct populations of numerous species, with the exception of Osmundson et al. (2005), in which this was not addressed. Although a few alpine Russula species have been reported from the Rocky Mountains (Cripps and Horak 2007; Cripps and Horak 2008; Cripps et al. 2016), detailed investigation is still needed to understand the true diversity of the genus in this region. 27 Ecological and Biogeographic Research on Arctic and Alpine Fungi in North America In the last decade, there has been a dramatic increase in the number of studies concerned with biogeography and distribution of Arctic fungi in North America. József Geml, a mycologist from Hungary, assessed the biodiversity of Lactarius in Arctic tundra and boreal forests of Alaska using 95% and 97% internal transcribed spacer sequence similarity. He found strong habitat preference and a high diversity in the genus, noting that species richness appeared to decrease with increasing latitude (Geml et al. 2009). A few studies have hypothesized that long distance dispersal plays an important role in the distribution of Arctic fungal genera (Geml et al. 2012; Timling et al. 2014). Other recent studies have assessed how these fungi are affected by climate change and a warming Arctic (Deslippe and Simard 2011; Dislippe et al. 2012; Morgado et al. 2015; Geml et al. 2015, 2016; Semenova et al. 2016). Although not strictly concerned with North America, several reviews have addressed fungi in the Arctic. These reviews focus on cold adaptation (Robinson 2001), Basidiomycota decomposers (Ludley and Robinson 2008), the current status of fungal knowledge and species richness (Dahlberg and Bültmann 2013), the taxonomic and ecological structure of basidial macromycetes (Shiryeaev et al. 2018), and fungi in polar regions (Tsuji and Hoshino 2019). Prominent microbial ecologist, Steve K. Schmidt and colleagues have assessed the presence of dark-septate fungi and arbuscular mycorrhizal fungi in the Front Range of the Colorado Rocky Mountains (Schmidt et al. 2008). More recently Schmidt et al. (2012) examined the diversity of fungi in-between the alpine zone and the zone of permanent ice, which lacks plants and is referred to as the periglacial zone. They 28 compared fungal communities in the periglacial zone in Colorado, the Andes, and the Himalayas and found that zoosporic fungi in the order Spizellomycetales dominate (Schmidt et al. 2012). They go on to hypothesize that these fungi complete their lifecycle with the available free-moisture from snowmelt and then remain dormant for a majority of the year. Working closely with Schmidt, Ph.D. student Clifton P. Bueno de Mesquita, was the main author on multiple papers that investigated the arbuscular mycorrhizal fungi and root endophytes on Niwot Ridge in Colorado. They characterized the development of these fungi throughout the plant growing season (Bueno de Mesquita et al. 2018a) and found that colonization by these two groups of fungi was highly variable between plant species. They concluded that these fungi play important roles in the cycling of nitrogen and phosphorus in alpine systems (Bueno de Mesquita et al. 2018b). Ectomycorrhizal Fungi and Plant Hosts in Arctic-Alpine Environments Currently, the dominant hypothesis explaining the distribution of Arctic and alpine floras states that the alpine flora is derived from an ancient Arctic flora, which has moved up and down mountains due to changes in glaciation across North America (Billings 1974). Alternatively, Weber (2003) hypothesized that North American alpine floras have been around since the Tertiary period and therefore predate modern Arctic floras. This would imply that mountainous floral communities migrated into low-land Arctic communities following deglaciation. If Weber’s (2003) hypothesis is true, large- scale migration and an Arctic origin of alpine floras may not explain the modern Arctic- alpine floral and fungal communities as proposed by Billings (1974). Based on Weber’s (2003) hypothesis the term ‘oroboreal’ is a more appropriate descriptor than 29 ‘circumpolar’ for describing the distribution of floral and fungal communities throughout the northern hemisphere because it implies a mountainous rather than Arctic origin. Either way, plant communities in the alpine are in some ways similar to those in the Arctic and there is considerable overlap of species within the two regions (Billings 1974). However, Billings (1974) hypothesizes that alpine plant species are ecotypically different due to isolation during glacial phases of the Pleistocene and Chapin and Körner (1995) believe alpine communities have higher species richness due to niche differentiation. Woody plants are particularly important in the alpine and in combination with grasses comprise the largest biomass in the biome (Körner 2003). A majority of the vascular-plant biomass is made up of less than ten species in most Arctic and alpine zones and most of the plant diversity will be present in 1 km2 (Chapin & Körner 1995). These Arctic and alpine plants have adapted to the harsh conditions present in these cold- dominated regions (Körner 1999). Most Arctic and alpine vascular plants form mutualistic relationships with fungi in what is called mycorrhizal symbiosis. The most abundant woody plants in these regions are the shrubs Salix L., dwarf Betula L. (Körner 2003), and Dryas L., which are known to form ectomycorrhizae (Trappe 1962; Miller et al. 1974; Chapin and Körner 1995; Cripps and Eddington 2005). A few plants such as Bistorta vivipara (L.) Delarbre and the sedge Kobresia Willd. are also known to form ectomycorrhizae (Trappe 1962, 1987; Lipson et al. 1999; Schadt 2002; Cripps and Eddington 2005; Ronikier and Mleczko 2006; Bjorbækmo et al. 2010; Thoen et al. 2019). In general, ectomycorrhizal fungi increase plant nutrient uptake (primarily nitrogen), ameliorate water relations, 30 protect roots from pathogen invasion, and can prevent heavy metal uptake. In return, the plant provides the fungus with photosynthetically derived sugars (Halling 2001; Smith and Read 2008). Hobbie and Hobbie (2006) reported that ectomycorrhizal fungi can provide up to 86% of the nitrogen required by the plant host in Arctic tundra and Lipson et al. (1999) showed that ectomycorrhizal fungi in association with Kobresia play an important role in the transfer of nitrogen in the form of amino acids. Ectomycorrhizal fungi also provide indirect benefits in Arctic and alpine regions where the mycelium aggregates and stabilizes soil (Graf and Brunner 1995). Ectomycorrhizal fungi are present in most major forest ecosystems in North America and Eurasia (Smith and Read 2008; Tedersoo et al. 2012), where they play an important role in seedling establishment, survival, and growth (Tedersoo et al. 2012). Concordantly, without these mutualistic fungi, most plants could not survive in harsh Arctic and alpine environments due to a short growing season, low temperatures, and poor soil nutrition (Haselwandter and Read 1980; Read and Haselwandter 1981; Gardes and Dahlberg 1996; Bjorbækmo et al. 2010). Shrubs are expanding into Arctic (Sturm et al. 2001, 2005) and alpine habits (Formica et al. 2014) in response to changes in climate. Research has shown that fungal community composition can change under simulated climate change scenarios that increase temperature, alter nutrients, and increase snow cover, all of which can lead to a decline in species richness (Sturm et al. 2001, 2005; Semenova et al. 2016). Deslippe et al. (2011) reported that following warming of dwarf Betula in the Arctic, some groups of ectomycorrhizal fungi like Cortinarius (Pers.) Gray increased in abundance while others like the Russulaceae (Russula Pers. and Lactarius Pers.) decreased in abundance. 31 Semenova et al. (2016) found significant decreases in ectomycorrhizal species richness under experimentally increased snow-cover in the Arctic. Their research suggests that ectomycorrhizal fungi may not be entirely driven by vegetation, as previously thought (Dahlberg and Bültmann 2013) and that they may also be influenced by complex biotic and abiotic interactions in the soil. As climatic warming continues to effect Arctic and alpine systems, the loss of entire plant functional groups could have large impacts on ecosystem function and health (Chapin & Körner 1995) leading to reduced stability due to decreases in community heterogeneity (Graae et al. 2018). These factors have important implications for the diversity of mycorrhizal species in Arctic and alpine regions. As climate change continues, ectomycorrhizal fungal communities will begin to shift or even decline. This could alter ecosystem functioning and highlights the importance of understanding ectomycorrhizal fungi in Arctic (Timling et al. 2014) and alpine environments (Formica et al. 2014). The Rocky Mountains The Rocky Mountains extend from northern Alaska to northern New Mexico and form a semi-contiguous alpine zone (Billings 1988). The Rocky Mountains were thrust up through thick sedimentary rocks and erosion exposed ancient granites and metamorphic rock. However, sedimentary rocks including shale, limestone, sandstone, and quartzite still remain, and make up entire subranges of the Rocky Mountains, basalt, andesite, and rhyolite can also be found (Retzer 1956). The semi-contiguous alpine zone of the Rockies creates isolated island populations (Kuchler 1964) where classic island 32 biogeography theory could apply to populations of animals, plants, and fungi; species richness is correlated with mountain top size and proximity of other alpine zones (MacArthur & Wilson 1967; Hadley 1987). It has been hypothesized (Weber 2003) that the Rocky Mountain flora is derived from Middle Asiatic flora and that these regions were connected by Greenland during the early Tertiary age. If true, it suggests that dry conditions in parts of the Rocky Mountains may have led to the attrition of the diverse Middle Asiatic floral communities, which could be true for fungi as well. Many of the ectomycorrhizal plants that occur in the Rocky Mountain alpine zone have broad circumpolar distributions with disjunct components in alpine areas to the south of the Arctic (Hultén 1968). Historically, changes in climate have altered the geographic distributions of alpine floras; as warming continues, alpine floral communities are shifting upwards in elevation and may disappear completely as subalpine communities move toward the tops of mountains (Fagre 2009). As in other Arctic and alpine regions, Salix species are the most abundant ectomycorrhizal host in the central and southern Rocky Mountains, and dwarf Betula is found in restricted areas. Rocky Mountain ectomycorrhizal woody hosts include: Betula glandulosa Michx., Dryas octopetala L., Salix arctica Pall., S. glauca L., S. planifolia Pursh, and S. reticulata L.; Betula glandulosa is a close relative of the B. nana L. complex, which has a circumpolar distribution (Hultén 1968). Salix reticulata also has a circumpolar distribution in Arctic areas, excluding Greenland (Hultén 1968). Worldwide these species are 100% mycorrhizal with ectomycorrhizal fungi being present on roots a majority of the time (Newman and Reddell 1987; Cripps and Eddington 2005). In one Colorado study site, it 33 was shown that Salix species have expanded 441% in the last 62 years (Formica et al. 2014), highlighting the importance of gathering baseline data prior to large shifts in alpine ecosystem function. In the Rockies, the vegetation and geology have been well studied, but little is known of the ectomycorrhizal fungi associated with the alpine flora. Over 200 macro-fungal species have now been documented in the Rocky Mountain alpine zone since Gardes and Dahlberg’s (1996) review and over half are ectomycorrhizal species (Cripps and Horak 2008). The main diversity of ectomycorrhizal fungi with woody plants in the Rocky Mountain alpine are in the genera Cortinarius (Peintner 2008), Entoloma P. Kumm., Hebeloma (Fr.) P. Kumm. (Beker et al. 2010; Cripps et al. 2019a), Inocybe (Fr.) Fr. (Cripps et al. 2010; Larsson et al. 2014; Larsson et al. 2018; Cripps et al. 2019b), Laccaria Berk. & Broome (Osmundson et al. 2005), Lactarius (Barge et al. 2016), and Russula. The diversity of several of these genera has already been addressed. Here we propose to examine the diversity and ecology of the important ectomycorrhizal genus Russula in the Rocky Mountain alpine zone. This work will first provide an overview of the order (Russulales) and family (Russulaceae), in which Russula is classified, in order to better understand the importance of the genus. Russulales Kreisel ex P.M. Kirk, P.F. Cannon & J.C. David (2001) The order Russulales is one of the largest orders within the Agaricomycetes. Even though the Russulales contains one family that produces gilled mushrooms, it is distantly related to the Agaricomycetidae, which is the subclass containing most of the gilled fungi. The Russulales contains 98 genera, 10 families, and 4,436 accepted species (He et al. 2019). The Russulales are now understood to be an old lineage that began with a 34 polyporous habit, in which species radiated into an array of morphological forms. Basidiocarps can be resupinate, effused-reflexed, discoid, clavarioid, or pileate-stipitate structures (Miller et al. 2006). Hymenophore types are also heterogenous and include smooth, hydnoid, poroid, and lamellate forms. Families now in the Russulales were once thought to be unrelated because of this diversity of features. The order was first described by Kreisel (1969) and at this time only included Russula, Lactarius, and related hypogeous taxa; also, it was not validly published. The modern Russulales was initiated by Donk (1964, 1971) who noticed that genera with vastly different macro-morphology such as lamellate Russulas and crust-like Gloeocystidiellums Donk shared unique microscopic characters such as gloeocystidia (cystidia with oily contents staining gray or black in sulfovanillin) and spores with amyloid ornamentation. Donk (1971) was later focused on the Hydnaceae s.l. and found that the genus Hericium Pers. also shared these features. He felt that gloeocystidia and amyloid spore ornamentation were important taxonomic characters for future classification. Oberwinkler (1977), on examining the microscopic characters of basidiomycetes, determined that the classification system at that time, did not reflect natural relationships. Building off of Donk’s work, Oberwinkler (1977) grouped various families containing gloeocystidia and amyloid spores into the Russulales. The order was validly published by Kirk et al. (2001), and at that time included 11 families and 64 genera. This grouping is now widely accepted and strongly supported to be monophyletic using various taxon sampling methods and multiple genes (Hibbet and Donoghue 1995; Hibbett et al. 1997; Binder and Hibbett 2002; Larsson and Larsson 2003; Larsson et al. 35 2004; Binder et al. 2005; Miller et al. 2006; Larsson 2007; Matheny et al. 2007). Researchers were drawn to sort out relationships within the Russulales using molecular techniques and focused on agaricoid, gasteroid, pleurotoid, and corticioid taxa (Miller et al. 2001, 2006; Larsson and Larsson 2003; Larsson 2007). Recently, the number of families has been reduced from eleven (Kirk et al. 2001) to ten, which includes the Albatrellaceae, Auriscalpiaceae, Bondarzewiaceae, Echinodontiaceae, Hericiaceae, Hybogasteraceae, Peniophoraceae, Russulaceae, Stereaceae, and Xenasmataceae (He et al. 2019). Currently, fifteen genera within the Russulales are classified as incertae sedis (He et al. 2019). Phylogenetic analysis of the order has helped confirm the defining characteristics of the Russulales. The presence of gloeocystidia is now accepted as a synapomorphic trait defining the russuloid clade. Amyloid spore ornamentation is frequently thought to be a synapomorphic character; however, this trait is not found in all of the Russulales and is found outside the order (Miller et al. 2006). Most fungi in the Russulales are saprotrophs, but two families are ectomycorrhizal (Russulaceae and Albatrellaceae), a few genera are root parasites (Heterobasidion and Echinodontium), and some species are insect symbionts (Klepzig et al. 2001). Recent research has examined the evolutionary shift from a saprophytic to mycorrhizal ecology and it is thought that the ectomycorrhizal habit evolved independently in the only two ectomycorrhizal families, Russulaceae and Albatrellaceae (Looney et al. 2018). These two families are also believed to harbor the highest diversity of hymenophore types (Miller et al. 2006). Thus, phylogenetics and 36 molecular techniques have provided systematists with a better understanding of the evolutionary relationships within the order, but much work still remains to be done. Russulaceae Lotsy The family Russulaceae is an old, highly specious, and morphologically diverse lineage of ectomycorrhizal fungi that has undergone rapid diversification (Looney et al. 2016). The family contains around 2,000 species (Kirk 2017; Miller et al. 2006; Hyde et al. 2016; Buyck et al. 2018) and seven genera (He et al. 2019). Four genera form pileate- stipitate structures including Lactarius (Persoon 1797), Lactifluus (Pers.) Roussel (Buyck et al. 2008), Multifurca Buyck & V. Hofst. (Buyck et al. 2008), and Russula (Persoon 1796) and three genera are crust fungi including Boidinia Stalpers & Hjortstam, Gloeopeniophorella Rick, and Pseudoxenasma K.H. Larss. & Hjortstam. The two most recognized genera, Russula and Lactarius, have been known since Persoon (1796, 1797). Both were originally described by Fries (1821) as tribes under the genus Agaricus L.: Russula under the tribe Russula and Lactarius under the tribe Galorrheus (Fr.) Fr. During the same year, Gray (1821) lifted both Russula and Lactarius to the generic level and both have remained stable. However, recent phylogenetic analyses have shown that these two genera as known do not form monophyletic groups; this led to the creation of the genus Multifurca and revision of the genus Lactifluus (Buyck et al. 2008), which are primarily tropical groups (De Crop et al. 2017; Wang et al. 2018). Traditionally, the Russulaceae has been considered to be comprised primarily of pileate-stipitate species, other morphological forms are now included. Macowanites Kalchbr., Gymnomyces Masse & Rodway, Cystangium Singer & A.H. Sm., and Martellia 37 Mattir. are hypogeous and gasteroid genera within the family that were once grouped together, separately from gilled taxa. Phylogenetic analysis has placed these genera within the pileate-stipitate Russula clade and all names have been synonymized with Russula (Elliot and Trappe 2018). The genera Zelleromyces Singer & A.H. Sm. and Arcangeliella Cavara were also erected to describe hypogeus taxa, but research has shown them to be phylogenetically within Lactarius (Miller et al. 2001); they have not yet been synonymized with Lactarius. The family Russulaceae is well-supported as a monophyletic group by multiple molecular phylogenetic studies (Miller et al. 2006; Larsson 2007; Phookamsak et al. 2019) and it is likely that this phylogenetic classification will remain stable. Because the Russulaceae is highly specious and has undergone rapid diversification many species still remain undescribed. To complicate the matter further, species recognition and delineation is difficult. A synapomorphic trait for the Russulaceae is the presence of sphaerocysts (sometimes referred to as sphaerocytes); these are large round (isodiametric) cells that swell during basidiocarp development. Sphaerocysts are responsible for the brittle texture of mushrooms in the family (Buyck et al. 2018). Interestingly, sphaerocysts are only found in this family of the Russulales. Because both sphaerocysts and filamentous hyphae are present in the pileus, stipe, and lamellar trama, tissues of the Russulaceae are termed heteromerous. The Russulaceae are easily defined based on a unique set of morphological characters but placing species in either Russula or Lactarius can be difficult. Verbeken (1996) looked at developmental differences between Russula and Lactarius and found 38 that Lactarius is composed of only one primary rosette complex of sphaerocysts; whereas, Russula is composed of many “agglomerate” rosettes. To clarify, Lactarius species do not contain sphaerocysts in the lamella (Heilmann-Clausen et al. 1998). This distinction has been used to place unknown taxa in either Russula or Lactarius. Although, other researchers have found this distinction to be of little use in tropical species (Henkel et al. 2000). The presence of pseudocystidia in Lactarius and their absence in Russula appears to be a more reliable character for differentiation between the two genera (Miller et al. 2006). Pseudocystidia are cystidia located on the hymenium that lack basal septa because they are connected to the lactiferous system in basidiocarps of Lactarius (Heilmann- Clausen et al. 1998). Species in the genus Lactarius produce a milk or latex on cutting; whereas, Russula species do not, but this character can be obscured in drier climates. Research has determined that the acrid or hot tastes present in some Russula and Lactarius species are caused by sesquiterpenoid unsaturated dialdehydes (Vidari and Vita-Finzi 1995; Clericuzio and Sterner 1997; Clericuzio et al. 1998; Wang et al. 2006; Hanson 2008; Malagòn et al. 2014). The acrid/hot taste has been used as a character for placement within infrageneric classifications of Russula (Miller and Buyck 2002), but is variable in some species. Interestingly, the compounds responsible for the acrid or hot tastes in Lactarius have been shown to produce an anti-feeding response in some insects (Daniewski et al. 1993, 1995). Members of the Russulaceae can be found from the Arctic (Knudsen et al. 2012) to the tropics (Henkel et al. 2000). They form ectomycorrhizal associations with various 39 plant hosts and are also known to be broadly associated with mycoheterotrophic plants in the genus Hexalectris Raf. (Kennedy et al. 2011). The Russulaceae are considered to be one of the most dominant families in ectomycorrhizal communities (Horton and Bruns 2001; Tedersoo et al. 2010). The family contains species that are thought to be late-stage colonizers of forests (Twieg et al. 2007). There is some debate regarding the temporal partitioning of species of Russula and Lactarius, some research shows that the latter is more abundant during the fall and the former is present year-round (Koide et al. 2007); however, there is evidence that contradicts this (Courty et al. 2008). In the tropics pleurotoid species of Russula and Lactarius were thought to be lignicolus based on the observation that these fungi fruited directly from tree trunks in elevated positions (Singer 1952, 1984; Dennis 1970; Pegler and Fiard 1979; Redhead and Norvell 1993; Verbeken 1998). However, more current research has matched sequences of ectomycorrhizae in the soil to basidiocarps, providing evidence that these species are ectomycorrhizal (Henkel et al. 2000). The Russulaceae are widely distributed, ecologically valuable, and form mutualistic associations with important plant hosts. These characteristics indicate that the Russulaceae could be used as a model study system to understand the history, functionality, and role of ectomycorrhizal fungi in the environment (Looney et al. 2018). However, the fact that many species within the family still remain undescribed hinders the value of this model system and highlights the importance of first studying the species diversity within the Russulaceae, especially within the large genus Russula. 40 Russula Persoon The ectomycorrhizal genus Russula contains hypogeous, sequestrate, and pileate- stipitate basidiocarps and is commonly cited as containing 750 species worldwide; however, this is known to be an underestimate of the true species diversity, which is estimated to extend to 3,000 species (Buyck 2007; Buyck et al. 2015; Kirk 2017; He et al. 2019). The type species of the genus is R. emetica (L.) Pers. Russula species have a colorful convex to funnel shaped pileus, attached lamellae that are not usually decurrent, a white to dark yellow spore print, no clamp connections, no latex, amyloid ornamented basidiospores, and sphaerocysts in a heteromerous trama (Romagnesi 1967; Singer 1986; Sarnari 1998–2005). Russula is an obligate mycorrhizal genus that requires a complex set of nutrients from its host plant for growth. Researchers still do not understand the growth requirements of Russula species and because of this they are considered unculturable. However, Hintikka and Niemi (1999) reported growing four different Russula species on commonly used media, but growth was slow. Historically, the taxonomy of Russula is complex and numerous publications produced by many authors have attempted to provide a usable and coherent classification for the genus. Three major monographs on Russula have been written based on the morphological characters present in basidiocarps (Romagnesi 1967; Singer 1986; Sarnari 1998–2005). These monographs have produced complex infrageneric classifications that include subgenus, section, and subsection levels. Eleven subgenera within Russula are currently recognized, although the validity of some subgenera is still under debate; these include Russula subgenus Russula Buyck & V. Hofst., R. subg. Compactae (Fr.) Bon, 41 emend. Buyck & V. Hofst., R. subg. Heterophyllidia Romagnesi, R. subg. Ingratula Romagnesi, R. subg. Amoenula Sarnari, R. subg. Incrustatula Romagnesi, R. subg. Archaea Buyck & V. Hofst., R. subg. Brevipes Buyck & V. Hofst., R. subg. Malodora Buyck & V. Hofst., R. subg. Crassotunicata Buyck & V. Hofst., and R. subg. Glutinosae Buyck & X.H. Wang (Sarnari 1998–2005; Hongsanan et al. 2015; Das et al. 2017; Buyck et al. 2018, 2020). There are also many sections and subsections within the infrageneric classification of Russula and the most recent revision of the classification, based on morphology, was performed by Sarnari (1998–2005). The infrageneric classifications produced by Romagnesi (1967), Singer (1986), and Sarnari (1998–2005) all differ slightly and even the most recent classification has not resolved all observed clades in molecular phylogenetic analyses. Miller and Buyck (2002) showed that while some infrageneric classifications are well supported, others are not, and do not appear to be natural phylogenetically related assemblages of species. Due to these inconsistencies, the infrageneric classifications at the section and subsection level within Russula will not be listed here. Recently, molecular phylogenetic analysis using the ITS, LSU, mtSSU, RPB1, and RPB2 regions has resolved eight major clades in the genus Russula (Looney et al. 2016; Buyck et al. 2018; Adamčík et al. 2019). These clades include the Russula crown clade, Russula core clade, Malodora, Compacta, Archaea, Heterophyllidia, Crassotunicata, and Brevipes. The Russula crown clade and Russula core clade together represent R. subgenus Russula and the remaining six clades phylogenetically represent the subgenera from which their names are derived. The ultimate goal of Russula 42 taxonomy is to create an infrageneric classification that matches strongly supported, multi-gene phylogenies (Adamčík et al. 2019). The morphological and the molecular complexity within Russula has confused taxonomists who have had difficulty applying the correct names to taxa. This has led to the creation of many erroneous or superfluous names in the literature, further complicating Russula taxonomy. Buyck (2007) and Buyck et al. (2015) have discussed many of the issues facing the North American Russulologist, including the lack of specialists and the need for high quality type descriptions. There is no doubt that the genus Russula needs taxonomic and systematic revision within North America. Mushrooms produced by certain species of Russula are economically important as edibles for humans. They are harvested around the world as a food source (Hu and Zeng 1992; Guo 1992; Rammeloo and Walleyn 1993; Buyck 1994) and are an important source of essential fatty acids (Sande et al. 2019). Worldwide there are 128 species of Russula that are considered edible or medicinal (Boa 2004). Russula mushrooms are unculturable; therefore, the medicinal and edible mushrooms produced hold substantial economic value. In small and rural communities, especially in Asia, mushroom foragers sell species like R. virescens (Schaeff.) Fr. for economic gain (Sitta and Davoli 2012). Russula virescens and other Russula species bring in high prices and a large market share, similar to highly regarded edibles like truffles (Tuber P. Micheli ex F.H. Wigg. spp.) and Matsutake (Tricholoma matsutake (S. Ito & S. Imai) Singer). Russula mushrooms also play important ecological roles in the diets of insects and larger mammals (Fogel 1975; Fogel and Trappe 1978; Lebel and Tonkin 2007). 43 Morphological Characters Morphological observation of Russula species has continued since the time of Fries (1874) when he wrote Hymenomycetes Europaei. As with many other genera, Fries based his descriptions on characters found in/on the pileus, stipe, and lamellae. Unlike most genera these characters proved to be less informative or more difficult to interpret in Russula. Burlingham (1944) discussed how traditional characters like pileus color, which shows considerable color variation, might be useful in species delineation. However, she stressed the importance of taste, odor, and spore ornamentation in iodine for distinguishing species of Russula, indicating that these characters hold just as much importance as characters originally used by Fries. Taste, odor, and spore ornamentation have indeed proven to be important characters in Russula and are now commonly used in descriptions and mapped onto molecular phylogenies (Romagnesi 1967; Singer 1986; Sarnari 1998–2005; Miller and Buyck 2002; Adamčík et al. 2019). The most well-known odors are the fishy smell that characterizes R. subsection Xerampelinae (Adamčík and Knudsen 2004) and the foul smell characteristic of the foetid Russula species (Shaffer 1972). Malagòn et al. (2014) noted that the origin of the compounds responsible for pungent or peppery taste in Russula may be useful in taxonomic classification. It is worth noting that species producing these hot or acrid tastes also tend to also produce a dark purple or black reaction in sulfovanillin when chemically tested (Favre-Bonvin and Bernillon 1982). Chemical reactions in Russula will be discussed later in more detail. Another character that has been observed by many mycologists in Russula, is the degree to which the cap cuticle can be peeled toward the pileal center before it separates, usually described as: 44 hardly peeling near margin (<20%), separable except at center (20–95%), or completely separable (Buyck 2019). All the major monographs written on the genus also include in-depth descriptions of microscopic characters (Romagnesi 1967; Singer 1986; Sarnari 1998–2005). This includes spore shape and size, spore ornamentation type, and descriptions of the basidia, cystidia, pileocystidia (sterile cells on cap), and pileipellis (cap cuticle). The basidiospores range from globose to elliptic and possess ornamentation that includes spines (acute tips), warts (obtuse tips), ridges (linear elements connecting warts and spines), and/or wings (ridges > 2 µm tall) (Adamčík et al. 2019; Buyck 2019). A small area lacking ornamentation is present on the basidiospore adjacent to the apiculus, which is referred to as the suprahilar plage, its relative size and degree of amyloidity is often described (Buyck 2019). Basidia length and width are measured at the widest and tallest points. Basidia are usually clavate with a pedicellate base (Sarnari 1998–2005), but variation can occur, and basidia can have one to four sterigma. Cystidia length and width are measured as above and the shape is recorded as clavate, fusiform, mucronate (with appendage), or pedicellate; the refractive contents, and their staining in the presence of chemical compounds is noted. Pileocystidia are present in most groups and lacking in a few (Incrustatula). They are sometimes difficult to differentiate from hyphae but are usually more clavate and filled with refractive contents (Sarnari 1998–2005). The staining reaction of the pileocystidia is also noted in sulfovanillin. For the pileipellis, the depth and composition of the suprapellis (upper portion near surface containing pileocystidia and hyphae) and subpellis (portion between the suprapellis and trama) are also described 45 (Adamčík et al. 2019) (FIG. 1A). Details of the pileipellis and pileocystidia are still regarded today as some of the most important characters for differentiation of Russula species (Adamčík et al. 2016b). Kühner (1975) and Ruotsalainen and Vauras (1994) noted the importance of the pileal coating and other characters for differentiating Russula species. Recently, Bart Buyck has recently led the field in regard to examining the microscopic characters of Russula. Buyck (1991) published a paper on how to properly describe and measure the spores and basidia. Buyck (2007) and Buyck et al. (2015) emphasized the importance of proper microscopic examinations and drawings, especially for North American taxa that are not well known. Buyck also offers detailed information on how to describe Russulas on his website Russulales News (Buyck 2019). Bazzicalupo et al. (2017) found a loose connection between morphological character states and species boundaries within Russula. This was done using a multivariate analysis that included molecularly delineated species of Russula and an extensive set of macro- and micro-morphological character information. The results explained some of the complexities that traditional taxonomists have faced. Morphological characters predicted genetic species only about 50% of the time. Their work pointed out that some common characters used in describing Russula did not vary enough to be informative, and this includes surface cap texture when wet or dry, and how far the cap cuticle could be peeled. Currently, Russulologists worldwide are working to produce a globally comprehensible Russula language to tackle some of the taxonomic needs of the genus (Adamčík et al. 2019). In order to produce a globally comprehensible Russula language, Adamčík et al. (2019) stress the importance of proper measurements, 46 drawings, and descriptions of the basidiospores, basidia, cystidia, pileocystidia, and pileipellis. Experts on the genus hope that using a thorough approach when describing Russula species, will help aid in the arduous process of delineating species. Chemical Reactions When working with Russula species, mycologists use various chemicals to produce macroscopic reactions on fresh basidiocarp material and microscopic reactions on basidiospores, basidia, cystidia, hyphae, and the pileipellis. Chemicals are used to aid in the taxonomic sorting of species. While chemicals are used in other fungal groups, chemical reactions can be especially useful in genera containing complex morphology and high species diversity such as Russula. Chemicals used to invoke reactions on fresh basidiocarps include ferrous sulfate (FeSO4), Phenol, and Gum Guaiac. Melzer and Zvara (1928) discovered a pinkish or greenish reaction caused by FeSO4 when it is placed on the fresh context of some Russula species. Singer (1962) reported four different reactions when FeSO4 was applied to the fresh material of Russula. These included a negative reaction in R. cyanoxantha (Schaeff.) Fr., a variable reaction in R. ferrotincta Singer, an olive-green reaction in R. section Xerampelinae, and a pink or salmon reaction in most Russula species. Watling (1971) hypothesized that the olive-green reaction in the R. xerampelina group may be directly related to the volatile compounds that produce the fishy odor. When applied to fresh fungal material, FeSO4 becomes oxidized to a ferric salt that can react with phenolic compounds present. In a laboratory, ferric chloride produces a green color when exposed to catechol; however, this reaction has not been specifically studied in Russula (Watling 47 1971). Phenol is also used to test for various color reactions on fresh basidiocarp material: a chocolate or brownish reaction is present in most species within the Russulaceae (Watling 1971). Lastly, Gum Guaiac is used to test for a greenish to brownish oxidation reaction, which indicates a positive result. The positive reaction is present in most species, so a negative reaction can be taxonomically informative (Moser 1978). There are five main chemicals that are used to test for microscopic reactions on tissues of Russula. These include carbolfuchsin, congo red, cresyl blue, Melzer’s reagent, and sulfovanillin. The purpose of using these chemicals is outlined in Adamčík et al. (2019) and will be briefly reviewed here. Carbolfuschin is used to determine if acid- resistant incrustations are present in primordial hyphae (Romagnesi 1967). Congo red is used to help distinguish various structures in the hymenium and pileipellis (Heilmann- Clausen et al. 1998). Cresyl blue is used to test for a metachromatic reaction in the incrustations found in pileocystidia and hyphae of the pileipellis (Buyck 1989). Melzer’s reagent causes the ornamentation on Russula spores to turn dark blue to black, a reaction referred to as amyloid; a reddish-brown reaction is called dextrinoid. This reaction is thought to be caused by an interaction between iodine in the solution and starch-like polysaccharides on the spore surface, although little research has investigated this (Watling 1971). These reactions are important in separating the Russulaceae from other agarics (Watling 1971). The amyloid reaction produced by Melzer’s reagent was one of the first chemical tests to be widely accepted as having taxonomic value (Melzer and Zvára 1927; Donk 1964; Watling 1971) and helps differentiate species based on spore 48 ornamentation. Sulfovanillin is widely used to observe the microscopic features of Russula (Caboň et al. 2017) and its usefulness was first discovered in the Russulaceae (Donk 1964). Sulfovanillin will turn the contents of the pileocystidia and hymenial cystidia bluish gray to blackish in some species. When this reaction occurs in cystidia, they are referred to as gloeocystidia and this is a synapomorphic trait of the Russulales. Ecology and Hosts Russula species have a cosmopolitan distribution and can be found from the tropics (Buyck et al. 1996; Tedersoo and Nara 2010) to the Arctic (Knudsen et al. 2012; Geml et al. 2009). In tropical regions, the Russula species and their ectomycorrhizal host are vastly different from those in temperate, Arctic, and alpine regions (Buyck et al. 1996; Tedersoo and Nara 2010). This work focuses on Russula in Arctic and alpine habitats, and therefore will focus on the ecological interactions in these regions. Russula species form symbiotic mutualistic ectomycorrhizal associations with all major linages of ectomycorrhizal hosts as well as some less common ectomycorrhizal shrubs and herbaceous species (Romagnesi 1967; Knudsen and Borgen 1982; Singer 1986; Buyck et al. 1996; Sarnari 1998–2005; Knudsen et al. 2012; Miller et al. 2012). Many Russula species are reported as generalists in the literature (Adamčík et al. 2006; Looney et al. 2016). A few species have been reported as host specialists with Salix (Adamčík and Knudsen 2004), Fagus sylvatica L., Quercus L., and Betula (Adamčík et al. 2006). A few studies used phylogenetics and ecological analysis to reveal niche partitioning of Russula species in different habitats and high species diversity in Alaska (Geml et al. 2010; Geml and Taylor 2013). Looney et al. (2016) analyzed metadata from Russula and relatives in 49 GenBank and found host specialization in some plant groups, including Fagaceae (25%) and Pinaceae (24%), while others where angiosperm generalists (12%) or associated with both angiosperms and Pinaceae (9%); the remainder had no host metadata. After phylogenetic analysis, Looney et al. (2016) determined that host association is highly variable in all of the Russula clades studied. Russula species apparently originated in temperate regions with angiosperms and the highest rates of diversification are seen in extratropical taxa (Looney et al. 2016). This is contrary to trends observed in most flora and fauna, which are the most biodiverse in tropical regions (Hillebrand 2004). Looney et al. (2016) hypothesized that numerous abiotic and biotic factors promote Russula diversification in temperate regions, a major factor being the ectomycorrhizal hosts involved. They showed that Russula diversification corresponds with that of major ectomycorrhizal plant lineages like Betulaceae and Salicaceae. In Arctic and alpine habitats, dwarf and shrubby species of Salix are the most common ectomycorrhizal hosts (Knudsen et al. 2012; Cripps and Horak 2008). DNA extracted from soil cores and grouped into OTUs using 97% sequence similarity has shown Russula to be one of the most abundant ectomycorrhizal associates of Arctic and alpine Betula (Deslippe et al. 2011) and to be the fourth most dominant ectomycorrhizal genus in moist tussock and dry tundra in Arctic Alaska (Morgado et al. 2015). Based on morphological data, a particular subset of Russula species is hypothesized to be intercontinentally distributed in Arctic and alpine environments, where they associate with Salix (Ohenoja 1972; Miller 1982a; Miller et al. 1973; Watling 2005; Ryberg et al. 50 2009, 2011), Dryas (Kernaghan and Currah 1998; Ryberg et al. 2009), Bistorta L. Scop. (Watling 2005; Mundra et al. 2016; Thoen et al. 2019), and Betula (Ohenoja 1972; Kernaghan and Currah 1998; Deslippe and Simard 2011; Voitk 2015). For years researchers have been collecting basidiocarps of Russula near Salix in subalpine, alpine, and Arctic habitats. These include collections from Alaska (Miller et al. 1973), Canada (Miller et al. 1973), The Rocky Mountains (Cripps and Horak 2008), Svalbard (Skift 1989), Greenland (Knudsen and Borgen 1982; Lamoure et al. 1982; Adamčík and Knudsen 2004), Iceland (Hallgrimsson 1998), and Europe (Favre 1955; Adamčík and Knudsen 2004; Watling 2005; Ryberg et al. 2009, 2011) to name a few. There is no doubt that Russula species have a strong association with Salix in Arctic and alpine habitats; however, the importance of this host in subalpine or boreal habitats is less clear. Research aimed at understanding these associations with Salix may find that more Russula species are strictly associated with Salix in subalpine or boreal habitats, as was shown with R. subrubens (Adamčík and Knudsen 2004; Adamčík et al. 2016b). Salix is often overlooked in forests where tall trees are often assumed to host ectomycorrhizal fungi. Studies that dig up, sequence, and identify ectomycorrhizal components of Salix root tips will clarify these questions. Much research has focused on understanding the ectomycorrhizal associations between Russula and Betula species in Arctic-alpine regions. Russula species have been reported to form ectomycorrhizal associations with Betula nana L., B. glandulosa, and B. pubescens Ehrh. in Alaska, Canada, Fennoscandia, and Greenland (Miller et al. 1973; Miller 1982b; Elborne and Knudsen 1990; De Groot et al. 1997; Deslippe et al. 2011; 51 Deslippe and Simard 2011). Some Russula species that associate with Betula in Greenland, also occur with the same host in Europe; these species are strictly associated with Betula (Elborne and Knudsen 1990). Fertilization experiments have shown, that as ectomycorrhizal mutualists, Russula species require less carbon from Betula nana and provide better access to labile nitrogen compared to other ectomycorrhizal associates (Deslippe et al. 2011). Other work has shown that in a mycorrhizal community of B. nana dominated by Russula species, below- ground transfer of carbon is facilitated between separate B. nana individuals; however, no transfer of carbon was observed between or within other tundra plant species (Dislippe and Simard 2011). This suggests that Russula species are some of the most important ectomycorrhizal hosts of B. nana. Research is beginning to assess how climate change will alter mycorrhizal fungal communities, but there is no consensus on how Russula species will be affected. Deslippe et al. (2011) reported that following experimental warming of dwarf Betula in the Arctic tundra, some groups of ectomycorrhizal fungi increased in abundance while others, especially Russula with a high affinity for labile nitrogen, decreased in abundance. In another experiment, Deslippe et al. (2012) showed that the genus Russula increased in abundance with warming treatments imposed on Arctic soils. Morgado et al. (2015) studied the effect of increased summer temperature on Russula species in the Alaskan tundra and did not observe any changes in fungal community composition, although there was a shift in extramatrical mycelium morphology. 52 Environments like the Arctic and alpine are seeing some of the largest negative impacts of climate change. As shrub expansion and nitrogen deposition continue in conjunction with changes in climate, the composition of ectomycorrhizal fungal communities will begin to shift. The functioning of individual ectomycorrhizal species is not redundant, with various species, even within the same genus, showing preferences for different forms of nitrogen (Antibus et al. 2018). It is also believed that warming will increase the growth and nutrient demand of ectomycorrhizal hosts like Betula nana, which would increase fungal density and decomposition (Deslippe et al. 2011). Ectomycorrhizal fungi will also play important roles in the spread of shrubby plant species as the climate warms (Deslippe and Simard 2011) and Russula species do have the ability to follow changes in plant host distribution due to climate change (Looney et al. 2019). These changes in ectomycorrhizal communities will alter basic microbial functioning across the landscape, which has important implications for plant ecology and plant processes over time (Cripps and Eddington 2005; Timling et al. 2014). All this underlines the importance of understanding the diversity and function of ectomycorrhizal genus Russula prior to large environmental shifts. Russula in Arctic-alpine habitats of Europe, Asia, and Arctic Islands The study of the genus Russula has been dominated by European mycologists who have published extensively on the genus (Fries 1874; Killerman 1936; Lange 1937; Romagnesi 1967; Singer 1986; Sarnari 1998–2005). Although the study of the genus has been ongoing for around 150 years, European mycologist did not focus heavily on Arctic and alpine Russula until the 1950s. However, Killerman described one of the most well- 53 known and most common Arctic and alpine Russula species, R. nana Killerm. in 1936. The most commonly reported species in Arctic or alpine habitats in Europe, Asia, and Arctic Islands are R. nana, (= R. emetica (Schaeff.) Pers., R. alpina (A. Blytt & Rostr.) F.H. Møller & Jul. Schäff., and R. emetica var. alpina A. Blytt & Rostr.), R. laccata Huijsman (= R. norvegica D.A. Reid), R. pascua (F.H. Møller & Jul. Schäff.) Kühner, and R. delica Fr. TABLE 4 summarizes the reports of Arctic and alpine Russula species. Table 4. Reports of Arctic-alpine Russula species in Eurasia, on Arctic Islands, and in North America. Names as reported, some are now changed or synonymized. Species with an * represent potential subalpine species that were only reported from Greenland. References at end of table1. AK = Alaska, CAN = Canada, DK = Denmark, GNLD = Greenland, ICLD = Iceland, JPN = Japan, EUR = Alps, Pyrenees or Carpathians, FNSC = Fennoscandia, FO = Faroe Islands, NL = Netherlands, PL = Poland, SCT = Scotland, SVBD = Svalbard, RM = Rocky Mountains, U.S.A., RUSS = Russia. Taxa Arctic-Alpine Locations R. aeruginea Lindblad ex Fr. CAN (63), GNLD (51, 65, 79) R. alnetorum Romagn.* GNLD (13) R. amoenipes Romagn. EUR (47), FNSC (47) R. alpina (A. Blytt & Rostr.) F.H. Møller & Jul. FNSC (52), GNLD (45, 50, 51, 65, 79, 80), ICLD (18), SVBD Schäff. (46) R. altaica (Sing.) Sing. CAN (63), GNLD (12, 13, 41, 50), RUSS (75), SVBD (46, 76) R. chamiteae Kühner EUR (27), FNSC (30), GNLD (12, 41, 50) R. citrinochlora Singer* GNLD (12, 13, 42), RUSS (43) R. claroflava Grove* GNLD (13, 41, 50, 51) R. claroflava var. viridis Knudsen & T. Borgen* GNLD (42) R. consobrina (Fr.) Fr.* GNLD (13, 42) R. cupreola Sarnari EUR (24) R. decolorans (Fr.) Fr.* GNLD (13, 51) AK (44), CAN (63), EUR (19, 72, 73), FNSC (28, 52), GNLD R. delica Fr. (12, 13, 41, 45, 50, 51, 65), ICLD (33), RM (20), RUSS (26), SVBD (46, 76) R. delica var. trachyspora Romagn. GNLD (41) R. dryadicola R. Fellner & Landa EUR (17, 23, 24), FNSC (17), GNLD (13), ICLD (33), RUSS (26) R. emetica var. alpestris (Boud.) Singer AK (55), EUR (21) R. felleaecolor Bon & Jamoni EUR (40) R. felleicolor Bon & Jamoni RUSS (26) R. flava Romell FNSC (52) 54 R. fragilis (Pers.) Fr. AK (44), CAN (53) R. gracilis var. altaica Singer GNLD (51), RUSS (75) R. gracillima Jul. Schäff. GNLD (13), ICLD (18, 33) R. groenlandica Ruots. & Vauras GNLD (13, 70) R. heterochroa Kühner EUR (47, 66), FNSC (47) R. laccata Huijsman CAN (63), EUR (24), FNSC (28), NL (36), RM (20) R. laevis Kälviäinen, Ruotsalainen & Taipale FNSC (3) R. lundellii Singer ICLD (33) R. lutea (Huds.) Gray CAN (68), GNLD (51) R. maculata subsp. alpina (Singer) Knudsen & T. Borgen GNLD (42), RUSS (43) R. medullata Romagn.* GNLD (12, 13, 41, 50) AK (54), CAN (62, 63), EUR (1, 5, 6, 7, 8, 10, 15, 16, 19, 21, 22, 24, 32, 35, 38, 39, 47, 48, 49, 56, 64, 66, 71, 72, 73, 74, 77), R. nana Killerm. FNSC (28, 29, 30, 34, 47, 61, 71), FO (78), GNLD (12, 13, 41, 50, 79), ICLD (33), NL (25), PL (67), RM (20, 57), RUSS (26, 43, 60), SCT (81), SVBD (30, 31, 76) R. nana var. alpina (A. Blytt & Rost.) Bon EUR (9) R. nauseosa (Pers.) Fr. EUR (72) R. nitidia (Pers.) Fr. AK (44), GNLD (13, 41), ICLD (33) CAN (37), EUR (11, 14, 27), FNSC (30) GNLD (13, 41, 50, R. norvegica D.A. Reid 79, 80), ICLD (33), JPN (59), RUSS (43), SCT (24, 81), SVBD (76) R. nuoljae Kühner FNSC (4, 69), RUSS (69) R. obscura (Romell) Peck* GNLD (65, 79) R. ochroleuca Fr. CAN (53) R. oreina Singer EUR (9), GNLD (41, 65, 80), ICLD (33), RUSS (58) R. pallidospora J. Blum ex Romagn. EUR (73), RUSS (26) R. paludosa Britzelm. CAN (63) R. pascua (F.H. Møller & Jul. Schäff.) Kühner EUR (2, 4, 16, 24, 27, 67, 72, 73), FNSC (28), FO (2), GNLD (2, 13), PL (2, 4, 67), RM (20), RUSS (2), SCT (81) R. pectinatoides Peck JPN (59) R. persicina Krombh. GNLD (13), SCT (81) R. pubescens A. Blytt CAN (63) R. puellaris Fr. EUR (9, 67), GNLD (13, 41), PL (67) R. pulchella I.G. Borshch.* GNLD (41) R. purpureofusca Kühner EUR (69), FNSC (69) R. queletii Fr. EUR (73), RM (57) R. rivulicola Ruots. & Vauras CAN (63) R. saliceticola (Singer) Kühner EUR (7, 24, 66, 72, 73), FNSC (28, 34), FO (77), GNLD (13, 41, 50), ICLD (33), SVBD (76) R. sororia (Fr.) Romell JPN (59) 55 R. sphagnophila Kauffman FNSC (52) R. sphagnophila var. heterosperma Singer GNLD (51, 65) R. subalpina O.K. Mill. AK (54) R. subrubens (J.E. Lange) Bon DK (4), EUR (4, 67), FNSC (4), GNLD (2, 13), PL (67) Russula Subsection Xerampelinae Singer CAN (63) R. versicolor Jul. Schäff.* GNLD (13, 51) R. violaceoincarnata Knudsen & T. Borgen CAN (63), GNLD (12, 13) R. xerampelina (Schaeff.) Fr.* GNLD (41, 51) R. xerampelina var. pascua F.H. Møller & Jul. Schäff. AK (55), EUR (21) R. xerampelina var. oreina Sing. FNSC (52) 1References: 1. Adamčík 1998; 2. Adamčík and Knudsen 2004; 3. Adamčík et al. 2019; 4. Adamčík et al. 2016b; 5. Bon 1985; 6. Bon 1987; 7. Bon 1991; 8. Bon 2000; 9. Bon and Ballarà 1996; 10. Bon and Cheype 1987; 11. Bon and Noguera 1995; 12. Borgen 1993; 13. Borgen et al. 2006; 14. Bresinsky 1987; 15. Bresinsky et al. 2000; 16. Bresinsky et al. 1980. 17. Caboň et al. 2019; 18. Christiansen 1941; 19. Corriol 2008; 20. Cripps and Horak 2008; 21. Favre 1955; 22. Fellner and Landa 1991; 23. Fellner and Landa 1993a; 24. Fellner and Landa 1993b; 25. Geml et al. 2014; 26. Gorbunova 2014; 27. Graf 1994; 28. Gulden 2005; 29. Gulden and Lange 1971; 30. Gulden et al. 1985; 31. Gulden and Torkelsen 1996; 32. Gyosheva and Dimitrova 2011; 33. Hallgrimsson 1998; 34. Hansen and Knudsen 1992; 35. Horak 1960; 36. Huijsman 1955; 37. Hutchison et al. 1988; 38. Jamoni 1995; 39. Jamoni 2008; 40. Jamoni and Bon 1993; 41. Knudsen and Borgen 1982; 42. Knudsen and Borgen 1992; 43. Knudsen and Mukhin 1998; 44. Kobayasi et al. 1967; 45. Kobayasi et al. 1971; 46. Kobayasi et al. 1968; 47. Kühner 1975; 48. Kühner and Lamoure 1986; 49. Lamoure 1982; 50. Lamoure et al. 1982; 51. Lange 1957; 52. Lange and Skifte 1967; 53. Linder 1947; 54. Miller 1982a; 55. Miller et al. 1973; 56. Moreau 2002; 57. Moser and McKnight 1987; 58. Mueller and Wu 1997; 59. Nara et al. 2003; 60. Niezdoiminogo 2003; 61. Ohenoja 2000; 62. Ohenoja and Ohenoja 1993; 63. Ohenoja and Ohenoja 2010; 64. Peintner 1998; 65. Peterson 1977; 66. Ronikier 2008; 67. Ronikier and Adamćík 2009; 68. Rostrup and Simmons 1906; 69. Ruotsalainen and Huhtinen 2015; 70. Ruotsalainen and Vauras 1994; 71. Sarnari 1998–2005; 72. Schmid-Heckel 1985; 73. Schmid-Heckel 1988; 74. Senn-Irlet 1987; 75. Singer 1938; 76. Skifte 1989; 77. Tondl 1988; 78. Vesterholt 1998; 79. Watling 1977; 80. Watling 1983; 81. Watling 1987. The literature on Arctic and alpine Russula can be grouped into two categories. The first group comprises publications dedicated entirely to the study of Russula in Arctic and alpine environments. These include detailed reviews of species known from the European alpine zone (Kühner 1975; Jamoni 1995; Bon 2000; Knudsen et al. 2012), the Monte Rosa Massif (Bon 1993), Slovakia (Ronikier and Adamčík 2009; Adamčík et al. 2006), Poland (Ronikier and Adamčík 2009), Fennoscandia (Knudsen et al. 2012; Ruotsalainen and Huhtinen 2015), Germany (Bresinsky et al. 1980), Svalbard (Skifte 1989), Greenland (Knudsen and Borgen 1982; Borgen 1993) and Romania (Ronikier 2008). Another detailed review focused solely on Russula sect. Xerampelinae associated 56 with dwarf Salix in Europe and Greenland (Adamčík and Knudsen 2004). The second group includes research that focused broadly on the taxonomy and distribution of basidiomycota in Arctic and alpine habitats and listed one, two, or three Russula species. Examples of these studies are those from Switzerland (Favre 1995), Norway (Lange and Skifte 1967), Slovakia (Fellner and Landa 1993a, 1993b), Svalbard (Kobayasi et al. 1968; Gulden and Torkelsen 1996), and the Pyrenean range (Bon and Ballarà 1996). A majority of this research on the taxonomy and classification of Arctic and alpine Russula species in Europe was based entirely on morphology. Many new species of Russula were described through this research in various Arctic-alpine regions of Europe; however, it was difficult to taxonomically place species based on morphology alone. Taxonomists relied on similarities to subalpine species found in the region. Most of the commonly reported species appeared to have wide ranges across Arctic and alpine habitats, while a few appeared endemic to particular regions. However, many identifications are likely incorrect, and many names may have been published superfluously, which complicates our understanding of Russula in Arctic and alpine regions. Modern molecular diagnostic tools can now provide insight into the diversity and phylogenetic relationships of fungi. Unfortunately, very few researchers have used these tools to understand the diversity and distribution of Russula species in Arctic and alpine habitats. One of the first molecular phylogenies that included Arctic or alpine species of Russula was that of Miller and Buyck (2002); the Arctic-alpine species were R. nana and R. pascua. Another study analyzed DNA from sporocarps and soil samples to determine 57 that Russula was the fifth most species-rich genus in Svalbard (Geml et al. 2012). They proposed that long-distance, transoceanic dispersal occurs in Arctic fungi, in contrast to the more limited dispersal of temperate Russula species (Bazzicalupo et al. 2019). As molecular tools became easier to use and more reliable, researchers began to sort out and clarify certain groups of Russula. Although they did not deal directly with Arctic and alpine species, the work of Adamćík et al. (2016a) and Caboň et al. (2017) produced important insights into understanding species diversity and the evolution of important characters in the genus Russula. Their phylogenies often included Arctic and alpine species, which provided a deeper phylogenetic understanding of Russula. Adamčík et al. (2016a) found that R. maculata Quélet actually consisted of a complex of three closely related species. The following year, Caboň et al. (2017) produced a multi-locus phylogeny and ancestral state reconstruction of Russula subsect. Rubrinae. They hypothesized that mild taste is the ancestral state for both the crown clade and Integrae clade and that acrid taste likely evolved more recently in several lineages within the crown clade. Adamčík et al. (2016b) did focus on Arctic and alpine species from Europe in the R. clavipes Velen. complex using molecular methods. This research determined that some traditional characters for delineating species, including pileus color and habitat preference, may be misleading. The clavipes complex consists of three closely related Russula species and metabarcoding approaches did not resolve ecological or biogeographical differences between two of these species. This indicates that even molecular approaches are not enough to understand Russula diversity unless coupled with 58 morphological, ecological, and biogeographical data. More recently, Caboň et al. (2019) determined that in boreal and Arctic environments of Eurasia the evolution of the R. globispora (J. Blum) Bon lineage has been shaped by glaciation events along with geographic and climactic disjunctions. Their work showed that European and North American collections of this species are nearly identical at the molecular level. Russula laevis Kälviäinen, Ruots. & Taipale was recently described from Arctic habitats of Finland and comparison to environmental sequences in GenBank indicate that this species is present in North America (Adamčík et al. 2019); it shares morphological similarities with the Arctic-alpine species R. delica, which has been reported from the Rocky Mountains (Cripps and Horak 2008) In summary, the study of the genus Russula was initiated by, and has been dominated by, European mycologists. Europeans provided a basis for the study of the complex morphology of the genus and began to explore Arctic and alpine habitats, which expanded knowledge of Russula diversity. Molecular work has uncovered previously unknown clades and helped reshape infrageneric classifications within the genus. However, molecular analyses certainly have not solved all of the taxonomic problems in Russula. The research of Adamčík et al. (2016b) showed that molecular information alone fails to resolve true species diversity. Knowledge of Russula diversity in Arctic and alpine habitats in the northern hemisphere is far from complete, and this is especially true for North America. European names have been applied to North American collections of Arctic and alpine Russula, but the validity of these names has not been confirmed using rigorous approaches, including phylogenetic analysis. 59 North American studies of Russula The study of Russula taxonomy in North America began in the 1870s with Peck, the botanist for the New York State Museum, who described R. sordida Peck (Peck 1873). Peck described 46 new species from the East Coast (Adamčík and Buyck 2014, Peck 1873, 1879, 1898, 1903, 1911) and reported many more (Peck 1906, 1907). W.A. Murrill also contributed significantly to the study of Russula when he described 110 species, mostly from Florida (Looney 2014); others also focused on Florida in the early- to mid-twentieth century (Beardslee 1934; Burlingham 1939; Murrill 1938, 1941, 1943, 1945a, 1945b, 1946). Russula diversity has also been examined in North Carolina (Beardslee 1918), the southern Appalachians (Bills and Miller 1984; Bills 1984, 1985, 1989), and northeastern Canada (Jennings 1936; Homola and Shaffer 1975). The genus has been covered more broadly in the East by (Peck 1906), Burlingham (1921, 1924, 1939, 1944), Singer (1942, 1943), and Fatto (2002). Kibby and Fatto (1990) also produced a key to Russula species found in Northeastern North America. Recently mycologist have begun to use molecular tools to understand the evolution and origins of Russula species in eastern regions, which are considered to have the highest diversity in North America (Looney et al. 2016; Looney et al. 2019). Buyck (2007) summarized the status of Russula research on the East Coast and determined that over 300 species have been described; he also provided a framework for future study of the genus. The first species described from the West Coast was R. nigrodisca Peck from material collected on St. Paul Island in the Bering Sea (Macoun 1899). In the West, regionally-based studies focused on Russula taxonomy covered Arizona (Fatto 2000), California (Earle 1902; Peters 1962; Thiers 1997a, 1997b), Colorado (Burlingham 1915; 60 Shaffer 1975; Fatto 1999), Idaho (Singer 1948; Shaffer 1964), Oregon (Kauffman 1930; Smith and Lebel 2001), Washington (Grund 1965, 1979), Wyoming (Burlingham 1915, 1924, 1936; Singer 1939b), and the Pacific Northwest in general (Burlingham 1913; Woo 1989). Russula in western regions of Canada (Spence 1932; Roberts 2007) and Alaska (Wells and Kempton 1914; Brunner 1989) have also been taxonomically studied. In recent years, Bazzicalupo et al. (2017) examined the morphological variation and value of the characters used to delineate Russula species in the Pacific Northwest. They analyzed one of the largest Russula datasets present in North America, produced by Ben Woo. Their results indicate that there is only a loose connection between morphological character states and species boundaries. Buyck et al. (2015) summarized the current status of Russula in the Western United States in a short paper. Based on all of the above research, only 58 species of Russula have been described from the western United States (Buyck et al. 2015; Phookamsak et al. 2019). Considering that Russula is the second most speciose ectomycorrhizal genus (Kirk et al. 2008), it is likely that many more species are yet to be discovered, especially in under-sampled regions like the Rocky Mountains. Burlingham (1915, 1924, 1936) and Singer (1938, 1939a, 1939b, 1948) provided general information on the genus in North America. Research has also focused on specific infrageneric groups of Russula, including R. subgenus Compactae (Shaffer 1962; Thiers 1994), R. subg. Heterophyllidia (Buyck and Adamčík 2011), R. subsection Emeticinae (Shaffer 1975), R. subsec. Lactarioideae (Shaffer 1964; Buyck and Adamčík 2013), R. subsec. Foetentinae (Shaffer 1972), R. subsec. Nigricantes (Adamčík and Buyck 2014), and R. subsec. Roseinae (Looney et al. 2019, 2020). Bergemann et al. 61 (2005, 2006) is the only researcher who appears to have performed population studies on Russula; she focused on R. brevipes Peck in North America. The R. emetica group (R. montana Shaffer) was breifly examined phylogenetically by Bazzicalupo et al. (2016). Arctic and alpine research on Russula in North America To date there has been little research focused on Russula in Arctic and alpine regions of North America. Researchers studying Russula in these regions have applied the names of taxa originally described from Europe to collections from North America. Russula emetica (likely R. nana) was one of the first names used to describe Russula found in Arctic northern Canada (Spence 1932; Jennings 1936). Since then, researchers have studied Russula species and their host associations in Arctic and alpine regions of Alaska, Canada, Greenland, and the Rocky Mountains. Miller et al. (1973, 1982) studied Russula species and their host associations in Arctic and alpine regions of Alaska. In Canada, Russula species have been collected near Rankin Inlet (Ohenoja 1972), around Schefferville (Kallio 1980), in Québec (Hutchinson et al. 1988), near Hudson Bay, and broadly across Arctic regions (Miller et al. 1973, Ohenoja and Ohenoja 2010). Greenland is also considered part of the North American continent and Russula found in Arctic habitats on the island have been more thoroughly sampled than the rest of North America. Russula species in Greenland have been studied in northern (Watling 1977, 1983), western (Lamoure et al. 1982), and southern regions (Lange 1957; Kobayasi et al. 1971; Elborn and Knudsen 1990) (see TABLE 4 for the specific Russula species reported). 62 Petersen (1977) investigated the ecology and phenology of fungi in Greenland and found six species of Russula distributed throughout various habitats. He determined that frost-free period and temperature are directly related to solar irradiance, making this the most important factor determining fungal fruitification, including that of Russula, in this Arctic region. Although the insular characteristics of Greenland make it a unique Arctic region, solar irradiance is still expected to contribute significantly to fungal fruitification in other Arctic and alpine regions. Borgen (1993) produced detailed descriptions of many fungi including eight species of Russula from Greenland. Knudsen and Borgen (1982) focused an entire paper on the Russulaceae in Greenland, which included excellent descriptions and a key to thirteen Russula species and one variety. Elbone and Knudsen (1990) studied the fungi associated with Betula pubescens in Greenland and confirmed three species of Russula associated with the Arctic shrub. More recently Adamcik and Knudsen (2004) assessed the diversity of R. sect. Xerampelinae associated with dwarf Salix on Greenland. In total, twenty-six species names have been used to describe Russula species on Greenland. Collecting in the Rocky Mountain alpine did not begin until much later when Moser and McKnight (1987) collected agarics at high elevations in Yellowstone National Park and the Beartooth Mountains. They reported R. nana for the first time from the Rocky Mountain alpine in association with Salix. This was the only report of alpine Russula in the Rockies until Cripps and Horak received a grant from the National Science Foundation in 1999 to study alpine Agaricales. Cripps and Horak began intensively surveying alpine fungi in the Rocky Mountains in 1999. Their work uncovered various 63 Russula species (Cripps and Horak 1999; Cripps 2003) and in 2007 they reported R. cf. delica Fr. for the first time from the Rocky Mountains in association with Dryas octopetala (Cripps and Horak 2007). The following year Cripps and Horak (2008) published their survey of agarics from the Rocky Mountain alpine zone focusing on the central and southern regions. In addition to R. cf. delica, they also provided the first formal reports of R. laccata and R. cf. pascua in the region and reported R. nana from the central and southern Rockies. Both R. nana and R. laccata were primarily associated with S. reticulata, and R. cf. pascua with S. glauca. Based on morphological characters Cripps and Horak (2008) reported five species of Russula, including one unidentified species. Overall, studies suggest that at least four taxa of Russula occur in the central and southern Rocky Mountain alpine zone and many more have been reported from Arctic and alpine areas of Canada and Alaska. The Russula species reported in Arctic and alpine areas of North America associate with a variety of hosts including Dryas octopetala, Bistorta vivipara, Betula glandulosa and several species of dwarf and shrubby Salix. All of the aforementioned research on taxonomy and systematics on Russula in Arctic and alpine habitats of North America has been based on morphological characters. Morphology alone suggests that at least some of the Russula species in Arctic and alpine habitats of North America are the same as those found in similar habitats in Europe (Lamoure et al. 1982; Knudsen and Borgen 1982; Borgen 1993; Elbone and Knudsen 1990; Adamčík and Knudsen 2004). However, these species and their potential intercontinental distributions have not been confirmed by modern molecular analyses. One recent paper dealt directly with taxonomy and systematics of the genus Russula in 64 Arctic and boreal habitats of North America and Europe using molecular phylogenetic methods but, this paper only covered the R. globiospora lineage (Caboň et al. 2019). They determined that R. dryadicola Fellner & Landa has an intercontinental distribution based on phylogenetic clustering of European material from the Alps and North American ITS sequences from Alaska. However, R. dryadicola has not yet been collected in North America. Research has indicated that morphological analyses often fail to recognize the true Russula diversity in a region (Buyck 2007). Other studies have found high diversity and species partitioning within Russula in the Arctic using a 97% ITS sequence similarity cutoff (Geml et al. 2010; Geml and Taylor 2013). These results indicate that the true diversity and function of Russula in Arctic and alpine habitats may not be fully understood. It is likely that there are more than the four species of Russula previously reported from the central and southern Rocky Mountain alpine, and that undescribed species are present in this region. To gain an accurate and stable understanding of Russula in the Rocky Mountain alpine, rigorous systematic approaches need to be applied. The goal of this research was to assess the diversity of Russula in the central and southern Rocky Mountains using morphological, ecological, and biogeographical data, along with molecular phylogenetics. This was done by comparing Rocky Mountain alpine Russula collections to type and reference material from Arctic, alpine, and subalpine habitats of North America and Europe. 65 CHAPTER TWO SYSTEMATIC ANALYSIS OF RUSSULA IN THE NORTH AMERICAN ROCKY MOUNTAIN ALPINE ZONE Introduction Arctic and alpine regions make up 5% and 3% of land on earth, respectively (Chapin & Körner 1995; Körner 1999; Körner 2003) and are collectively referred to as the Arctic-alpine biome (Bliss 1962; Billings 1973; Löve & Löve 1974; Bliss 1988; Chapin & Körner 1995; Murray 1995). Arctic and alpine regions are defined as the open, vegetated areas beyond the climatic limit of tree growth (Bliss 1988; Chapin & Körner 1995) or above tree line in mountainous regions worldwide (Körner 2003). The Arctic includes northern areas of Russia, Norway, Svalbard, Iceland, Greenland, Canada, and the United States of America (USA), which together comprise a circumpolar habitat (Callaghan et al. 2005). Alpine zones are found on high elevation mountain tops near the equator and at lower and lower elevations as you approach the poles, eventually merging with the Arctic (Billings 1974; Körner 1995). Research suggests that Arctic and alpine environments are being disproportionately affected by global warming compared to other systems (Serreze and Barry 2011; IPPC 2014). The temperature in Arctic regions is increasing 0.1° C per year (Anisimov et al. 2007; Comiso and Hall 2014), which is altering microbial function (Chapin et al. 1995) and leading to large scale changes in plant distributions (Sturm et al. 2001). As the climate warms, alpine communities will be restricted to ever-decreasing areas on mountain tops or will even disappear completely 66 (Grabherr et al. 1995). Most alpine regions are highly fragmented, disconnected, and contained within small areas of land, and the species present are often adapted to the harsh conditions found at high elevations (Anthelme & Lavergne 2018). Fungi play important ecological roles in Arctic and alpine habitats where they exist as mutualists that enhance nutrient uptake in alpine plants; as decomposers that replenish nutrients in poor alpine soils; and as pathogens that affect alpine plant populations (Haselwandter and Read 1980; Read and Haselwandter 1981; Gardes and Dahlberg 1996; Bjorbækmo et al. 2010; Haselwandter 2007). A knowledge of fungal biodiversity in these systems can be used as a model to understand larger biological trends regarding the distribution and evolutionary history of Arctic and alpine communities (Gardes and Dahlberg 1996). This information can also help scientists and regulators make informed decisions concerning the conservation needs of these sensitive systems which are increasingly threatened by climate change (Grabherr et al. 1995). Therefore, gaining baseline data on Arctic and alpine fungi is essential so that future change can be detected and considered in management plans (Dahlberg and Bültmann 2013). One of the most important fungal guilds in Arctic and alpine habitats are the ectomycorrhizal fungi that form mutualistic relationships with woody plants, which allows plants to survive in these cold-dominated habitats. In Arctic and alpine habitats, woody plants are ecologically important and in combination with grasses comprise the largest biomass in the biome (Körner 2003). In general, ectomycorrhizal fungi increase plant nutrient uptake (primarily nitrogen), ameliorate water relations, protect roots from 67 pathogen invasion, and can prevent heavy metal uptake. In return, the plant provides the fungus with photosynthetically derived sugars (Halling 2001; Smith and Read 2008). The most abundant ectomycorrhizal woody plants in these cold-dominated regions are the shrubs Salix L., dwarf Betula L. (Körner 2003), and Dryas L. (Trappe 1962; Miller et al. 1974; Chapin and Körner 1995; Cripps and Eddington 2005). One herb, Bistorta vivipara (L.) Delarbre, and sedges in the genus Kobresia Willd. are also known to form ectomycorrhizae (Trappe 1962, 1987; Lipson et al. 1999; Schadt 2002; Cripps and Eddington 2005; Ronikier and Mleczko 2006; Bjorbækmo et al. 2010; Thoen et al. 2019). Members of the ectomycorrhizal fungal family Russulaceae play important roles in nutrient cycling across a variety of ecosystems worldwide. The Russulaceae has been proposed as a model family to explore ecosystem function and evolutionary diversification in ectomycorrhizal fungi (Looney et. al. 2018). However, this model cannot be fully realized until species in the family are known and currently there is a knowledge gap. One of the largest ectomycorrhizal genera in the Russulaceae is Russula Pers. which is commonly reported to contain 750 species worldwide; however, this is known to be an underestimate of the true species diversity, which may extend to 3000 species (Buyck 2007; Buyck et al. 2015; Kirk 2017; He et al. 2019). Russula is most diverse in temperate zones where the genus apparently originated (Looney et al. 2016), but its species exist from the tropics to the Arctic. Russula is an important ectomycorrhizal genus in Arctic and alpine habitats. It is reported to be one of the most abundant ectomycorrhizal associates of Betula in Arctic and alpine regions (Deslippe et al. 2011) and is the fourth most dominant ectomycorrhizal 68 genus in moist tussock and dry tundra in Arctic Alaska (Morgado et al. 2015). A study that analyzed the DNA from sporocarps and soil samples, found Russula to be the fifth most species-rich genus in Svalbard, a large Arctic island (Geml et al. 2012). Two studies of Russula in boreal regions of Alaska found high diversity (42 nonsingleton OTUs in one study) and that there was species partitioning by habitat using ITS sequence similarity to environmental and physical collections (Geml et al. 2010; Geml and Taylor 2013). These studies suggest that the diversity of Russula may be high in Arctic and alpine habitats, and that the true number of Russula species in northern latitudes is not fully realized. The genus Russula comprises species with hypogeous, sequestrate, and pileate- stipitate basidiocarps, but only the latter has been reported from Arctic and alpine habitats. Pileate-stipitate Russula species have colorful, convex to funnel shaped pilei, attached lamellae that are not usually decurrent, a white to dark yellow spore print, no clamp connections, no latex, amyloid ornamented basidiospores, and sphaerocysts in a heteromerous trama (Romagnesi 1967; Singer 1986; Sarnari 1998–2005). The taxonomy of Russula is complex and several monographs have attempted to provide usable and coherent classifications for the genus (Romagnesi 1967; Singer 1986; Sarnari 1998– 2005). The type species for the genus is R. emetica (L.) Pers. and eleven subgenera are currently recognized phylogenetically; however, the validity of some is still under debate (Sarnari 1998–2005; Hongsanan et al. 2015; Das et al. 2017; Buyck et al. 2018, 2020). The study of Russula has been dominated by European mycologists who have published extensively on the genus (Fries 1874; Killerman 1936; Lange 1937; Romagnesi 69 1967; Singer 1986; Sarnari 1998–2005). Although these studies have continued for around 150 years, European mycologists did not focus heavily on Arctic and alpine species of Russula until the 1950s. In Arctic and alpine regions of Eurasia, Russula species have been reported from the European Alps (Kühner 1975; Jamoni and Bon 1993; Jamoni 1995, 2008; Bon 2000; Adamčík and Knudsen 2004; Knudsen et al. 2012; Adamčík et al. 2016b), the Monte Rosa Massif (Bon 1993), the Pyrenean range (Bon and Ballarà 1996; Corriol 2008), Slovakia (Fellner and Landa 1993a, 1993b; Ronikier and Adamčík 2009; Adamčík et al. 2006), Poland (Ronikier and Adamčík 2009; Adamčík et al. 2016b), Fennoscandia (Favre 1955; Lange and Skifte 1967; Knudsen et al. 2012; Ruotsalainen and Huhtinen 2015; Adamčík et al. 2016b), Germany (Bresinsky et al. 1980), Svalbard (Kobayasi et al. 1968; Skifte 1989; Gulden and Torkelsen 1996), Romania (Ronikier 2008), Denmark (Adamčík et al. 2016b), the Faroe Islands (Vesterholt 1998; Adamčík and Knudsen 2004), Scotland (Fellner and Landa 1993b; Watling 1987), and Iceland (Christiansen 1941; Hallgrimsson 1998). The majority of the taxonomy and classification of Arctic and alpine Russula species has been based almost entirely on morphology. Around 50 names have been used to describe Russula taxa in Arctic and alpine habitats worldwide, based mostly on basidiocarp surveys (TABLE 4, Chapter 1). However, this is likely an overestimate as some names have been synonymized and others have been incorrectly applied to Arctic and alpine Russula collections. That being said, it is also possible that Russula diversity has not been completely explored in Arctic and alpine regions. Morphological analyses often fail to recognize the true diversity of Russula in a region (Buyck 2007). 70 In North America, Russula diversity has been investigated in cold habitats to a limited extent in Alaska (Miller et al. 1973, 1982), Canada (Ohenoja 1972; Miller et al. 1973; Kallio 1980; Hutchinson et al. 1988; Ohenoja and Ohenoja 2010), and extensively in Greenland (Lange 1957; Kobayasi et al. 1971; Watling 1977; 1983; Knudsen and Borgen 1982; Lamoure et al. 1982; Elborne and Knudsen 1990; Borgen 1993; Adamcik and Knudsen 2004). Virtually nothing was known of Russula species in the central and southern Rocky Mountain alpine region until the last two decades. Russula nana was reported above treeline in Wyoming by Moser and McKnight in 1987, and it remained the only Russula known from the Rocky Mountain alpine until the early 2000s. Four species of alpine Russula were later reported from a large-scale survey of alpine Agaricales focused on the central and southern Rocky Mountains (Cripps and Horak 1999, 2008; Cripps 2003). This includes R. nana along with the first formal reports for Russula cf. delica Fr., R. laccata Huijsman, and R. cf. pascua (F.H. Møller & Jul. Schäff.) Kühner for the Rocky Mountains (Cripps and Horak 2008). These four species were found in association with a variety of hosts, including Dryas octopetala L., Bistorta vivipara, and species of shrubby (S. glauca L. and S. planifolia Pursh) and dwarf willows (S. reticulata L. and S. arctica Pall.) (Moser and Mcknight 1987; Cripps and Horak 2008). Most Russula species reported from the Rocky Mountain alpine appear to be in R. subgenus Russula and R. subgenus Brevipes. All of the taxonomic work on Russula in Arctic and alpine habitats of North America up to this time has been based solely on morphological characters. These species, their distribution, and their phylogenetic placement were not confirmed by molecular analyses. 71 Based on morphology, species in other genera previously reported from the Rocky Mountain alpine (Miller et al. 1973; Moser and McKnight 1987; Cripps and Horak 2008, 2010a, 2010b; Osmundson et al. 2005) are mostly those known from alpine habitats in Europe and the Arctic (Lamoure et al. 1982; Knudsen and Borgen 1982; Borgen 1993; Elbone and Knudsen 1990; Adamčík and Knudsen 2004). More recently, molecular research on Lactarius Pers., Inocybe (Fr.) Fr., and Hebeloma (Fr.) P. Kumm. has confirmed that most species from the Rocky Mountain alpine are also present in the alpine of Europe and the Arctic, although a few appear to be North American endemics (Barge et al. 2016; Cripps et al. 2019, 2020). All this suggests that there may be at least some overlap between Russula species in the Rocky Mountain alpine and those in similar habitats of Europe and the Arctic. If Russula species in the Rocky Mountain alpine are the same as those in Arctic and alpine habitats of Europe, this implies that these species have large, intercontinental distributions. Large, intercontinental distributions have been molecularly confirmed for other ectomycorrhizal genera from the Rocky Mountain alpine, including Hebeloma (Beker et al. 2010; Cripps et al. 2019), Cortinarius (Pers.) Gray (Peintner 2008), Inocybe (Cripps et al. 2010; Larsson et al. 2014; Larsson et al. 2018; Cripps et al. 2019), and Lactarius (Barge et al. 2016; Cripps and Barge 2016). While fungal communities in the Rocky Mountain alpine have certainly been shaped by glaciation and other abiotic factors, ectomycorrhizal fungi also need to follow their plant host. One of the primary hosts for ectomycorrhizal fungi, including Russula, in northern cold-dominated habitats are Arctic and alpine species of Salix (Knudsen et al. 2012; Cripps and Horak 2008); these species are known to have large, circumpolar 72 distributions in Arctic and alpine habitats (Hultén 1968). Some of these willow species are present in the disjunct isolated areas that comprise the Rocky Mountain alpine. As further evidence, Russula distributions have even been shown to shift in response to changes in plant host distribution due to climate change (Looney et al. 2019, 2020). We also know that Russula diversification corresponds with that of major ectomycorrhizal plant lineages such as Betulaceae and Salicaceae (Looney et al. 2016), both of which occur in Arctic and alpine regions. Thus, it is likely that alpine species of Russula in the Rocky Mountains have at least the potential to occur wherever their host exists, which could result in large distributions in the case of Salix-associated species. Large, intercontinental distributions have already been confirmed for two species of Russula. Russula dryadicola Fellner & Landa and R. laevis Kälviäinen, Ruots. & Taipale are reported in boreal and Arctic environments of Europe and confirmed in North American using molecular comparison to environmental sequences (Adamčík et al. 2019; Caboň et al. 2019). Russula laevis was previously reported from the southern Rocky Mountains as R. cf. delica which extends the range of this species (Cripps and Horak 2008). Therefore, we hypothesize that the Russula species that occur in the Rocky Mountain alpine will have large intercontinental distributions. Long distance, transoceanic dispersal has been hypothesized to explain minimal molecular differences in Arctic species of ectomycorrhizal fungi (Geml et al. 2012), in contrast to the more limited dispersal of temperate Russula species (Bazzicalupo et al. 2019). However, long distance dispersal is a problematic explanation for fungal distributions in disjunct alpine areas. The central and southern Rocky Mountains, which is the primary collecting sites for this 73 study, are thousands of miles from the Arctic and an explanation for large intercontinental distributions in Arctic-alpine fungi is outside the scope of this project. There are several possibilities for how Russula species came to occur in relatively young available Arctic and alpine habitats. Did one or a few species arrive and diversify like Darwin’s finches? Or did they originate from various subalpine lineages scattered throughout the Russula phylogeny? Molecular phylogenetics can elucidate the evolutionary history of these species within the entire Russula clade. Based on morphology alone, the few Russula species that have been reported in the Rockies appear to be distantly related (Moser and McKnight 1987; Cripps and Horak 2008), and do not appear to share a recent common ancestor. Thus, given the data so far, it appears most likely that Russula species in the Rocky Mountain alpine do not share a recent common ancestor unique to this group but instead evolved independently from more than one lineage into alpine habitats. Despite being a diverse genus, only about 58 species of Russula have been newly described from the western United States in all habitats (Buyck et al. 2015). This is certainly an under representation. It is also likely that there are more than the four species of Russula previously reported in the Rocky Mountain alpine zone, and that additional species remain to be found. To gain an accurate and stable understanding of Russula taxonomy, rigorous systematic approaches need to be applied. This research integrated morphological, ecological, and biogeographical data along with molecular phylogenetics to elucidate the true diversity of Russula in alpine regions of the Rockies. We hypothesize that 1) the species reported from the Rocky Mountain alpine will be the same 74 as those reported in Arctic and alpine habitats of Europe based on molecular and morphological data, 2) the Russula species present in the Rocky Mountain alpine will have large, intercontinental distributions, similar to the Arctic and boreal species studied molecularly in Europe, and 3) the Russula species in the Rocky Mountains have independently colonized alpine habitats and do not form a monophyletic group. This research analyzed the morphological and molecular characters of over 130 Russula collections (collected from 1997 to 2018) from the Rocky Mountain alpine in Montana, Wyoming, and Colorado and compared them to type and reference material from several herbaria throughout the world. Detailed morphological descriptions were created for each species; scanning electron microscope and compound microscope photos accompany descriptions. The diversity of Russula in the Rocky Mountain alpine was assessed with a multi-locus phylogeny, using the nuclear ribosomal ITS1-5.8S-ITS2 region (ITS barcode) and the second largest subunit of the RNA polymerase II gene (RPB2). Species distributions are addressed, and some Russula species are reported for the first time in the Rocky Mountain alpine and in North America. A key was created to help identify the Russula species present in the Rocky Mountain alpine zone using macro- and micromorphological features. Methods Study Sites The goal of this study was to determine which species of Russula occur in the central and southern Rocky Mountain alpine zone. The Rocky Mountains extend from northern Alaska to northern New Mexico and host a semi-contiguous alpine zone, which 75 consists of open, vegetated areas above treeline (Billings 1988). The vegetation is often variable, but consists of perennial grasses, sedges, forbs, low-growing prostrate shrubs and cushion plants; mosses and lichens are also abundant (Korner 2003). The Rocky Mountains were thrust up through thick sedimentary rocks and erosion exposed ancient granites and metamorphic rock. However, sedimentary rocks including shale, limestone, sandstone, and quartzite still remain, and make up entire subranges of the Rocky Mountains; basalt, andesite, and rhyolite can also be found (Retzer 1956). High elevation alpine environments in the Rockies often experience cold temperatures, high winds, and a majority of their moisture occurs as snow fall (Billings 1973; Körner 1999; Körner 2003); these factors lead to a short growing season. In the Rocky Mountain alpine, the ectomycorrhizal hosts include the shrubby willows Salix glauca and S. planifolia, and the dwarf willows S. reticulata and S. arctica. Other important hosts include Betula glandulosa Michx., Dryas octopetala, and Bistorta vivipara (Cripps and Eddington 2005; Cripps and Horak 2008). These hosts are found at most alpine field sites in the Rocky Mountains with the exception of B. glandulosa, which is more common in the southern Rockies of Colorado and the northern Rockies of Alaska, and less common in the central Rockies. This study was based on specimens collected in the Rocky Mountain alpine as well as reference material collected and obtained (through herbaria) from Arctic and alpine regions of Europe. A majority of the fungi used in this study were collected in the central and southern Rocky Mountains of Montana, Wyoming, and Colorado; a few collections came from Alaska. The main collecting sites for the central Rocky Mountains 76 are located in the Beartooth Mountains, which lie on the border of Montana and Wyoming. The Mountain range is part of the Absaroka-Beartooth Wilderness, which is located within the Custer, Gallatin, and Shoshone National Forests. The Beartooth Mountains are home to Granite Peak (3904 m), which is the highest point in the state of Montana. The mountain range includes multiple high elevation alpine plateaus. The Beartooth Plateau is considered one of the most biologically unique regions in North American and is the highest elevation alpine plateau in the forty-eight conterminous United States. Collecting sites located on the Beartooth Plateau include the Billings Fen (44°58′16.9″ N, 109°26′07.3″ W, 3230 m), Birch Site (45°01′26.64″ N, 109°24′29.164″ W, 2990–3020 m), Highline Trail (45°00′21.36″ N, 109°24′23.22″ W, 3060–3100 m), Frozen Lakes (44°57′55.92″ N, 109°28′59.88″ W, 3190–3200 m), Solifluction Terraces (44°58′22.32″ N, 109°26′48.6″ W, 3258 m), and Wyoming Creek (45°00′10.62″ N, 109°24′30.06″ W, 3200 m). Two other collecting locations in Montana were at Cinnabar Basin (45°07′21.07″ N, 110°48′59.17″ W, 1930 m) in the Absaroka Mountains and Hellroaring Plateau (45°03′31.79″ N, 109°28′36.58″ W, 3166 m) in the Beartooth Mountains. Study sites in the southern Rocky Mountains are in alpine areas of Colorado and include Independence Pass (39°06′36″ N, 106°33′36″ W, 3600–3700 m) and Cottonwood Pass (38°49′12″ N, 106°24′00″ W, 3700 m) in the Sawatch Range; Black Bear Basin (37°24′00″ N, 107°42′00″ W, 3760 m), California Gulch (37°55′30″ N, 107°35′06″ W, 3941 m), Cinnamon Pass (37°55′48″ N, 107°31′48″ W, 3700–3850 m), Engineer Pass (37°58′19.2″ N, 107°35′2.4″ W, 3901 m), Mineral Basin (38°45′38.16″ N, 106°23′57.12″ 77 W, 3900 m), and Stony Pass (37°46′48″ N, 107°31′48″ W, 3700 m) in the San Juan Mountains; Blue Lake area (40°05′24″ N, 105°37′12″ W, 3572 m), Loveland Pass (39°39′36″ N, 105°52′48″ W, 3700m), Mt. Evans (39°35′17.88″ N, 105°38′37.68″ W, 4310 m), and Niwot Ridge (40°03′34.92″ N, 105°37′00.84″ W, 3748 m) in the Front Range. Study sites in Alaska include Crow Pass road (61°04′26″ N, 149°07′28″ W, 457 m) near Girdwood and Alpine Glen (61°05′26.16″ N, 149°39′55.08″ W, 1070 m) on Flattop Mountain outside Anchorage. Taxon Sampling and Processing Basidiocarps of Russula were collected in the Rocky Mountain alpine during July and August from 1997 to 2018. All collections were described when fresh and photographed when possible. Habitat, possible mycorrhizal hosts, location, and date were noted for all collections. Spore prints were obtained when possible. All basidiocarps were dried on a standard dehydrator and deposited in the Montana State University herbarium (MONT), Bozeman, Montana 59717–3150. Other Rocky Mountain alpine collections of Russula were obtained from the Denver Botanic Gardens’ Sam Mitchel Herbarium of Fungi, Denver, Colorado (DBG). Reference collections were obtained from the University of Michigan fungal herbarium, Ann Arbor, Michigan (MICH), the New York Botanical Garden, Bronx, New York (NY), the Botanical Museum, University of Oslo, Norway (O), the Arctic University of Norway, Norway (TROM), and Åbo Akademi University, Finland (TURA). Herbarium Codes are from http://sweetgum.nybg.org/ih/ (Thiers B., continuously updated). 78 Our collections were initially identified based on morphological species concepts when possible, using taxonomic keys (Jamoni 1995; Sarnari 1998–2005; Bon 2000; Moreau 2002; Adamčík et al. 2016b) and personal expertise (Cripps pers. com. 1997– 2020). Detailed morphological descriptions were made for each collection and compiled for each species. Morphological descriptions include most of the characters traditionally described by fungal taxonomists, even if characters like the pileus color and the degree to which the cap cuticle peels have been determined to be less informative for species delineation in Russula (Bazzicalupo et al. 2017). Final species determinations were based on a combination of morphological, molecular, and ecological data, with the goal being concordance between all data types. If there were disagreements between particular datasets, we looked for concordance between as many independent data points as possible while taking the species taxonomic history into account. Morphological Descriptions Macromorphological characters were described using fresh material and micromorphological characters were assessed following observation of dried, rehydrated material. Macromorphological descriptions of the pileus, lamellae, stipe, and context follow Buyck (2019). The odor and taste of fresh material were included when possible. Spore print colors are described and compared to Romagnesi’s (1967) color code for spores. Chemical tests for ferrous sulfate (FeSO4), Phenol, and Gum Guaiac were done by placing one drop of each chemical on a small section of fresh context; reactions were observed for five minutes (Melzer and Zvara 1928; Watling 1971; Moser 1978). Exsiccata were also described. The degree to which the cap cuticle can be peeled was 79 described as: hardly peeling near margin (<20%), separable except at center (20–95%), or completely separable (Buyck 2019). The number of lamellae (L) refers to those that reach the stipe on select basidiocarps. A lower-case n was used to denote the number of basidiocarps from which lamellae were counted. A Leica DMLS research microscope was used for all microscopic observations and measurements. The micromorphology was examined for collections listed in TABLE 5. Micromorphological characters described and measured include basidiospores, basidia, hymenial cystidia, pileipellis, and pileocystidia. Hymenial cystidia refer to the sterile cells on the surface of the lamellae (Romagnesi 1967; Thiers 1994; Sarnari 1998–2005; Adamčík and Knudsen 2004; Adamčík et al. 2016b). Pileocystidia are unique terminal cells in the pileipellis with granular (and possibly gelatinous) contents of usually a larger width than hyphal terminations (Sarnari 1998–2005). For microscopic examination, fungal tissue was first reconstituted in 70% ethanol. Basidiospores were then observed in Melzer’s reagent (0.5 g iodine, 1.5 g potassium iodide, 20ml dH2O, 2 ml chloral hydrate) (Melzer and Zvara 1927) and basidia, hymenial cystidia, and pileocystidia were observed in 3% KOH and sulfovanillin. When observing features in sulfovanillin, a few crystals of vanillin were placed next to a reconstituted scalp section of the pileipellis or a piece of lamella and mixed with one drop of sulfuric acid and observed for five minutes; the reaction was recorded as: none, slightly graying, graying, slightly blackening, or blackening. On average, 20 basidiospores were measured at random from the apex of the stipe or directly from the lamellae. For each basidiospore, the length and width were measured 80 at the widest point under a 100× oil-immersion lens (1000× magnification); the apiculus and ornamentation were not included in measurements. The length to width ratio (Q) was calculated for each basidiospore and the average Q value was used to describe the spore shape: subglobose (Q = 1.05–1.15), broadly ellipsoid (Q = 1.16–1.3), ellipsoid (Q = 1.31–1.45), narrowly ellipsoid (Q = 1.46–1.6), oblong (Q>1.6) (Adamčík et al. 2019). The height of basidiospore ornamentation was recorded from at least five spores and the range recorded. Degree of amyloidity and relative size was recorded for the suprahilar plage. Ornamentation terminology follows Adamčík et al. (2019), and includes: spines (acute tips), warts (obtuse tips), ridges (linear elements connecting warts and spines), and/or wings (ridge > 2µm tall). The general appearance of the ornamentation was described as: isolated warts, clustered warts, individual lines, branched, subreticulate, or reticulate (Adamčík et al. 2019). Ten basidia, hymenial cystidia, and pileocystidia were randomly measured under a 40× lens (400× magnification) to determine length and width at the widest points; the contents were also described. Mucronate appendages on hymenial cystidia were measured, when encountered, and the range recorded. Within the pileipellis, the suprapellis (upper portion near surface containing pileocystidia and hyphae) and subpellis (portion between the suprapellis and context) were measured and the general form of each layer described (FIG. 1A). The general shape and width of the hyphal terminations located in between the supra- and subpellis were also described. On select taxa the length and width of the terminal cells of hyphae in the piliepellis were recorded from ten hyphal ends at the pileus margin and near the pileus center. Photographs of some of the micromorphological features are shown in FIG. 1. After 81 macro- and micromorphological descriptions were complete for each collection, they were combined for each species as determined by morphological, ecological, and molecular phylogenetic analyses. Then, the minimum measurement, standard deviation, average, average − standard deviation, average + standard deviation, and maximum measurement was calculated for basidiospores (length, width, and Q), basidia, hymenial cystidia, pileocystidia, and for hyphal terminations when applicable for each species; average values are italicized. Figure 1. Select microscopic features in Russula. A. Important microscopic features present in the pileipellis in 3% KOH (400×). B. Sphaerocysts in 3% KOH (1000×). C. Pileocystidia showing a blackening reaction in sulfovanillin, indicated by arrow (200×). D. Hymenial cystidia showing a blackening reaction in sulfovanillin, indicated by arrow (400×). 82 Scanning Electron Microscopy Morphological descriptions were complemented with scanning electron microscope (SEM) photographs of the basidiospores for most species found in the Rocky Mountain alpine (FIG. 17). These were included to provide detailed images of the basidiospore ornamentation, which can be useful for taxonomic identification and for understanding intraspecific morphological variation. The scanning electron microscope and associated equipment were provided by the Imaging and Chemical Analysis Laboratory at Montana State University. Dried pieces of lamella about 2 mm2 were attached to glass coverslips using double-stick tape. They were either iridium or gold sputter-coated for 45 seconds at 35 milliamps to achieve a coating thickness of 15 nm. Basidiospores were examined using a field emission scanning electron microscope (Zeiss SUPRA 55VP) at 1.00 KV. Photos were edited in Adobe Lightroom 3 and Photoshop v. 21.0.1. A scale bar was created by modifying the scale produced by the SEM. Compound Microscope Photography Morphological descriptions were complemented with compound microscope photographs of the basidiospores for all species found in the Rocky Mountain alpine (FIG. 18). These were included for the same reasons as the SEM photographs, but also to replicate what is observed under the compound microscope because most researchers won’t have access to SEM images. Basidiospores were photographed using a Sony a7 III camera with a Full Frame 2x DSLR/ML Microscope Camera Adapter on a Leica DMLS research microscope. Three to five separate photographs were taken at various focal depths of the same image to increase depth of field. Photographs were focus-stacked in 83 Helicon Focus 7 and edited in Photoshop v. 21.0.1. Photographs were taken of the eyepiece reticle scale so a 10 µ scale could be added to the basidiospore photographs. Table 5. Collections whose micromorphological features were examined1 or were included in the phylogenetic analyses2. Type specimens are bolded. ITS or RPB2 is bolded if the sequence was generated in this study. Herbarium Codes from http://sweetgum.nybg.org/ih/. Taxon Voucher Date Location Host or Habitat ITS RPB2 Lactarius deceptivus2 AFTOL ID 682 NA MA, U.S.A. NA AY854089 AY803749 Multifurca PC0723654 20 Jun Newton County, TX, ochricompacta2 2014 U.S.A. NA MH063878 MH061175 R. aeruginea2 AT2003017 NA Sweden NA DQ421999 DQ421946 R. albonigra2 AT2002064 NA Sweden NA DQ422029 DQ421966 R. aff. CLC 3822B 22 Aug Crow Pass Rd. Girdwood, alpigenes2 (MONT) 2018 AK, U.S.A. Dryas, Salix Yes Yes R. aff. CLC 3821 22 Aug Crow Pass Rd. Girdwood, alpigenes2 (MONT) 2018 AK, U.S.A. Salix Yes Yes R. altaica1,2 333753 (NY) 5 Aug 1937 Russia Salix, Betula Yes NA R. altaica1,2 CLC 1608 2 Aug Blue Lake Dam, CO, Betula, (MONT) 2001 U.S.A. Dryas Yes Yes R. altaica1,2 CLC 1618 3 Aug Blue Lake Dam, CO, (MONT) 2001 U.S.A. Betula, Salix Yes Yes R. amara2 FH 12213 18 Aug 2012 Hildesheim, Germany NA KT933998 KT933930 R. amoenipes2 309IS77 NA NA NA AY061656 NA R. cf. amoenoides2 Sav_F_1340 NA Italy NA KU205284 NA R. aquoa2 TROM_F_17338 8 Sept 2017 Norway Picea UDB037477 NA R. aquosa2 TU 101719 NA NA NA KX579813 NA R. atrorubens2 TU 101728 9 Sept Varstu vald, Võru 2011 maakond, Estonia Picea UDB011308 NA R. atrorubens2 TU 101718 NA NA NA KX579812 NA R. betularum2 PC BB2004-235 NA Great Smoky Mountains National Park, TN, U.S.A. NA EU598183 NA R. betularum2 SAV:F-20026 6 Oct 2014 Apuseni Mts., Romania Betula, Picea KY582694 KY616687 R. betularum2 JV 30628 17 Aug Tolola, Saukkovaara, Picea, 2014 Paltamo, Kainuu, Finland Betula, Salix, Yes NA Pinus Tsuga R. betularum2 BPL 269 8 Aug Great Smoky Mountains carolinensis, 2012 National Park, TN, U.S.A. Abies, Picea, KT933969 KT933900 Betula R. cf. brevipes2 321965 4 Jul Point Reyes National Pseudotsuga 2018 Seashore, CA, U.S.A. menziesii MH714879 NA R. cf. brevipes2 RK8 NA Canada NA KF007188 NA 84 R. brevipes1,2 CLC 2730 5 Aug (MONT) 2011 Alaska, U.S.A. Dryas Yes Yes R. brevipes1,2 CRN 063 (MONT) 18 Sept Hyalite Canyon, MT, 2017 U.S.A. Picea, Abies Yes NA R. brevipes2 18598 (DBG) 6 Jul 1996 Boulder, CO, U.S.A. Pinus ponderosa Yes Yes R. brevipes2 TENN:070667 16 Jul Great Smoky Mountains 2015 National Park, NC, U.S.A. NA KY848511 NA R. camarophylla2 PAM01081108 NA France NA DQ421982 DQ421938 R. chamiteae1 24741 (DBG) 23 Aug Glacier Lake, Near 2010 Roosevelt N.F., CO, Conifers, U.S.A. Salix Yes NA R. aff. chloroides2 FH 12273 10 Oct 2012 Belgium NA KT934015 KT933947 R. chloroides2 ue68 (TUB) NA Germany Fagus sylvatica AF418604 NA R. chloroides2 205Rus24 NA NA NA AY061663 NA R. chloroides2 RUS-12091401 14 Sept 2012 Ireland NA KF432954 NA R. claroflava2 FH 12212 18 Aug 2012 Hildesheim, Germany NA KT933997 KT933929 Betula, R. clavipes2 FR:652 NA RheinLahnKreis, Germany Populus, KU205304 NA Picea R. clavipes2 SAV:F-1327 NA W. Carpathians, Slovakia Picea, Abies, Sphagnum KU205292 NA R. compacta2 BPL 242 2 Aug Great Smoky Mountains Betula, 2012 National Park, TN, U.S.A. Tsuga KT933960 KT933890 R. compacta2 BPL 227 30 Jul 2012 TN, U.S.A Tsuga KT933952 KT933881 R. cremeirosea2 BPL 289 2 Oct 2012 Knoxville, TN, U.S.A. Quercus KT933983 KT933915 R. cuprea2 SAV:HK12050 17 Aug 2012 Gotland Island, Sweden Quercus KU886591 NA R. cuprea2 FH 12250 31 Aug 2012 Rudigsdorf, Germany NA KT934010 KT933942 14 Sept Ledmore Oakwood, Quercus R. cuprea2 LM-18 2006 Spinningdale, Sutherland, petraea, UDB002420 NA Scotland Betula R. cf. 10 Aug cyanoxantha2 BPL 280 2012 TN, U.S.A Tsuga KT933976 KT933908 R. decolorans2 FH 12196 2 Aug 2012 Ilmenau, Germany NA KT933992 KT933924 R. aff. delica2 FH 12266 3 Oct 2012 Walkenried, Germany NA KT934014 KT933946 R. aff. delica2 UE24.08.2004-20 NA NA NA DQ422005 NA R. cf. delica2 UBC F30260 2 Nov 2014 Canada NA KX812852 NA R. delica2 TU116211 22 Jul 2013 Tartu, Estonia NA UDB025023 NA R. delica2 hue22 (TUB) NA NA Fagus sylvatica AF418605 NA R. delica2 FH 12-272 10 Oct 2012 Belgium NA KF432955 NA R. dryadicola2 PAM95082603 26 Aug Dryas 1995 Svoie, France octopetala MG386715 MG386741 R. earlei2 BPL 245 3 Aug Great Smoky Mountains Quercus, 2012 National Park, TN, U.S.A. Fagus, KT933961 KT957331 Tsuga, Pinus 85 R. emetica2 Iw081 15 Oct Nationalpark Bayerischer, 1996 Germany Picea UDB000300 NA R. emetica2 JV 25569 5 Oct Pinus 2007 Finland sylvestris Yes NA R. emetica2 Buyck 2444 (517IS76) NA Europe NA AY061673 NA R. emetica2 UBC F30056 NA NA NA KX579781 NA R. emeticicolor2 FH 12253 5 Sept Erfurt, Aspenbusch, 2012 Germany NA KT934011 KT933943 R. faginea2 Sav:F-1336 NA Slovakia NA KU205285 NA R. farinipes2 UE28.09.2002.4 NA France NA DQ421983 DQ421939 R. favrei2 IB1995/0098 NA Tirol, Austria Conifers KU205318 NA R. foetens2 FH 12277 27 Aug 2012 Keula, Germany NA KT934016 KT933948 R. font-queri2 FH 12223 30 Aug 2012 Rudigsdorf, Germany NA KT934003 KT933935 R. fragilis2 FH 12197 18 Aug 2012 Hildesheim, Germany NA KT933993 KT933925 R. gigasperma2 438/BB 07.280 NA NA NA NA KU237787 R. globispora2 GENT FH2007 8 Sept BT111 2007 Thuringen, Germany NA KU928144 KY616671 R. gracillima2 JV 32758F 5 Oct 2007 Finland Picea, Betula Yes NA R. gracillima2 TU101698 6 Sept 2011 Krabi, Estonia Picea, Betula UDB011284 NA R. gracillima2 UE23.08.2004-14 NA NA NA DQ422004 NA R. gracillima2 FH 12-264 NA Germany NA KR364094 NA Picea, R. graminea2 KKJV 26121F 16 Aug (MONT) 2008 Finland Betula, Pinus Yes NA sylvestris R. graveolens2 Sav:F-1343 NA Slovakia NA KU205298 NA R. grisescens2 MxM R-0838 26 Jul 2008 Österreich (Tirol), Austria Picea abies UDB031192 NA Betula, R. grisescens2 JV 32642F 30 Aug Tololanmäki, Paltamo, 2018 Kainuu, Finland Alnus, Picea, Yes NA Populus R. CLC 1723 12 Aug Cottonwood Pass, CO, Alpine heterochroa1,2 (MONT) 2001 U.S.A. Dryas Yes Yes Arctic Salix R. CLC 1919 21 Aug Nordensklods Land, heterochroa1,2 (MONT) 2002 Adventdalen, Todalen, polaris, Yes Yes Svalbard Dryas octopetala Arctic Salix R. CLC 1918 21 Aug Nordensklods Land, polaris, heterochroa1,2 (MONT) 2002 Adventdalen, Todalen, Svalbard Dryas Yes Yes octopetala R. heterophylla2 UE20.08.2004-2 NA Sweden NA DQ422006 NA R. integra2 FH 12172 1 Aug Erfurt, Aspenbusch, 2012 Germany NA KT933984 KT933916 R. cf. CLC 2759 9 Aug Near Girdwood, AK, intermedia1,2 (MONT) 2011 U.S.A. Dryas, Salix Yes Yes R. cf. CLC 3784 20 Aug Alpine Glen, Flattop, AK, intermedia1,2 (MONT) 2018 U.S.A. Dryas, Salix Yes Yes R. cf. CLC 3822 22 Aug Crow Pass Rd., Girdwood, intermedia1,2 (MONT) 2018 AK, U.S.A. Dryas Yes Yes 86 R. intermedia2 Sav:F-3093 6 Sept Dovrefjell National Park, Betula 2009 Dovre, Oppland, Norway pendula KU928147 NA R. intermedia2 JV 32189F 27 Aug Melalahti, Paltamo, 2017 Kainuu, Finland Picea, Betula Yes NA R. aff. laccata1 23923 (DBG) 30 Aug Squaw Pass Road, CO, 2007 U.S.A. Salix, Picea Yes Yes R. cf. laccata2 UBC F30302 8 Nov 2014 Canada NA KX812849 NA Oulanka National Park, R. laccata1 JV 23194 26 Aug 2005 Kuusamo, Koillismaa, Salix Yes NA Finland R. laccata1 CLC 1381 19 Aug Birch Site, Beartooth Alpine Salix (MONT) 1999 Plateau, MT, U.S.A. planifolia Yes Yes R. laccata1 CLC 1465 6 Aug Independence Pass, CO, (MONT) 2000 U.S.A. Alpine Yes NA R. laccata1 CLC 1467.1 6 Aug Independence Pass, CO, (MONT) 2000 U.S.A. Alpine Yes NA R. laccata1 CLC 2275 1 Aug (MONT) 2005 Niwot Ridge, CO, U.S.A. Alpine Yes Yes CLC 2371 6 Aug Wyoming Creek, R. laccata1 (MONT) 2008 Beartooth Plateau, WY, Alpine Yes Yes U.S.A. Billings Fen Site, R. laccata1 CLC 3617 23 Aug (MONT) 2017 Beartooth Plateau, MT, Alpine Salix Yes Yes U.S.A. Billings Fen Site, R. laccata1 CRN 133 (MONT) 7 Aug 2018 Beartooth Plateau, MT, Alpine Salix Yes Yes U.S.A. R. laccata1 CRN 157 (MONT) 10 Aug Highline Trailhead, 2018 Beartooth Plateau, MT, Alpine Salix U.S.A. reticulata Yes NA R. laccata1 CRN 166 (MONT) 19 Aug Loveland Pass, CO, Alpine Salix 2018 U.S.A. planifolia Yes NA R. laccata1 CRN 175 (MONT) 21 Aug Niwot Ridge, CO, U.S.A. Alpine, Salix 2018 planifolia Yes NA R. laccata1,2 CLC 1378 19 Aug Birch Site, Beartooth (MONT) 1999 Plateau, MT, U.S.A. Alpine Salix Yes Yes R. laccata1,2 CLC 1487 9 Aug Loveland Pass, CO, (MONT) 2000 U.S.A. Alpine Yes NA R. laccata1,2 21511 (DBG) 21 Aug Loveland Pass, CO, 2003 U.S.A. Alpine Salix Yes Yes R. laccata1,2 21608 (DBG) 21 Aug Loveland Pass Lake, CO, 2003 U.S.A. Alpine Salix Yes Yes 6 Aug Solifluction Site, R. laccata1,2 CRN 114 (MONT) 2018 Beartooth Plateau, WY, Alpine Salix Yes Yes U.S.A. Highline Trailhead, R. laccata1,2 CRN 128 (MONT) 7 Aug 2018 Beartooth Plateau, Alpine Salix MT/WY, U.S.A. reticulata Yes Yes Alpine Salix R. laccata1,2 CRN 168 (MONT) 20 Aug 2018 Niwot Ridge, CO, U.S.A. planifolia, S. Yes Yes reticulata R. laccata1,2 CRN 186 (MONT) 24 Aug Blue Lake, CO, U.S.A. Alpine Salix 2018 planifolia Yes Yes R. laccata2 GC_1b_III Jun 2005 Sweden Salix JQ724003 NA R. laccata2 TU101871 22 Aug 2008 Savukoski, Finland Salix, Betula UDB016024 NA R. laccata2 23.08.2004-6 24 Aug 2004 Abisko, Lappland, Sweden Salix, Betula UDB000915 NA 87 R. laccata2 F-73860 (O) 24 Aug Finse, Ulvik, Hordaland, 2004 Norway NA Yes NA R. laccata2 F-127987 (O) 27 Jul Sør-Varanger, Finmark, 1979 Norway Marsh Yes NA R. laevis Lapponia (Holotype)2 JR4016 21 Aug 1995 Enontekionensis, Betula, Salix MN130091 NA Kilpisjaervi, Finland R. laevis1,2 CLC 1146 17 Aug (MONT) 1997 Niwot Ridge, CO, U.S.A. Alpine Salix Yes Yes Alpine Salix R. laevis1,2 CLC 1642 4 Aug Cottonwood Pass, CO, glauca, S. (MONT) 2001 U.S.A. arctica, Yes Yes Dryas Alpine Salix R. laevis1,2 CLC 1690 9 Aug (MONT) 2001 Stony Pass, CO, U.S.A. reticulata, S. Yes Yes arctica R. laevis1,2 CLC 1740 13 Aug Independence Pass, CO, Alpine (MONT) 2001 U.S.A. Dryas Yes Yes R. laevis1,2 CLC 1883 18 Aug Arctic Salix (MONT) 2002 Longyearbyn, Svalbard polaris Yes Yes R. laevis1,2 CLC 2271 1 Aug (MONT) 2006 Niwot Ridge, CO, U.S.A. Alpine Bistorta Yes Yes R. laevis1,2 CLC 2981 12 Aug Mt. Evans, CO, U.S.A. Salix arctica, (MONT) 2013 Bistorta Yes Yes R. laevis2 F-5917 (TROM) 20 Jul Nordkapp, Finnmark, 1961 Norway Dryas Yes (partial) NA R. laevis2 27685 (DBG) 20 Aug Alpine Salix 2013 Summit Lake, CO, U.S.A. arctica, S. Yes Yes glauca R. cf. maculata2 HJB 10019 NA Europe NA DQ422015 NA R. mairei2 FH 12262 2 Oct 2012 Walkenried, Germany NA KT934013 KT933945 R. montana Pseudotsuga (Holotype)1,2 12231 (MICH) 10 Aug 1972 Perigo, CO, U.S.A. menziesii, Yes NA Picea R. montana1 3596 (DBG) 15 Aug 1974 Perigo, CO, U.S.A. NA Yes NA R. montana1,2 CLC 1624 2 Aug Blue Lake Dam, CO, Betula (MONT) 2001 U.S.A. glandulosa Yes Yes Pinus R. montana1,2 19236 (DBG) 18 Aug 1997 Copper Mtn., CO, U.S.A. contorta, Yes Yes Picea R. montana1,2 19170 (DBG) 15 Aug Shrine Pass, Copper Mtn., Burned 1997 CO, U.S.A. wood, moss Yes NA R. montana1,2 19556 (DBG) 4 Aug 1998 Squaw Pass, CO, U.S.A. Picea, Abies Yes NA R. montana1,2 9654 (MICH) 13 Aug Jefferson Lake, CO, 1972 U.S.A. Picea, Abies Yes NA R. montana2 JV 9788 24 Sept Hammaudda, Jomala, Picea, Pinus, 1994 Åland, Finland Betula Yes NA R. montana2 NL GM15A-018 NA NA NA KX579801 NA R. montana2 UBC F30293 8 Nov 2014 Canada NA KX812853 NA R. nana2 01930708 (NY) 27 Aug 1984 Switzerland Salix retusa Yes NA R. nana1 CLC 1101 28 Jul Birch Site, Beartooth (MONT) 1997 Plateau, MT, U.S.A. Alpine Salix Yes (partial) Yes R. nana1 CLC 1156 16 Aug (MONT) 1997 Niwot Ridge, CO, U.S.A. Alpine Salix Yes Yes R. nana1 CLC 1426 31 Jul Engineer Pass, CO, U.S.A. Alpine Salix (MONT) 2000 planifolia Yes (partial) Yes 88 R. nana1 CLC 1467.2 6 Aug Independence Pass, CO, (MONT) 2000 U.S.A. Alpine Yes Yes (partial) 7 Aug Billings Fen Site, R. nana1 CRN 135 (MONT) 2018 Beartooth Plateau, MT, Alpine Salix U.S.A. planifolia Yes Yes R. nana1 CRN 154 (MONT) 10 Aug Highline Trailhead, Alpine Salix 2018 Beartooth Plateau, MT, planifolia, S. Yes (partial) Yes U.S.A. reticulata R. nana1 CLC 3575B 17 Aug Frozen Lakes, Beartooth Salix (MONT) 2017 Plateau, WY, U.S.A. reticulata Yes NA R. nana1 CLC 1544 20 Aug (MONT) 2000 Sismiut, Greenland Arctic Salix Yes herbacea Yes (partial) R. nana1 CLC 1875 31 Jul California Gulch, CO, Alpine Salix (MONT) 2002 U.S.A. arctica Yes Yes (partial) Alpine Salix R. nana1,2 CLC 1440 1 Aug Cinnamon Pass, CO, (MONT) 2000 U.S.A. planifolia, S. Yes Yes reticulata R. nana1,2 CLC 1450 3 Aug Black Bear Basin, CO, (MONT) 2000 U.S.A. Alpine Yes Yes R. nana1,2 CLC 1812 27 Jul Cinnamon Pass, CO, Alpine Salix (MONT) 2002 U.S.A. arctica Yes Yes R. nana1,2 CLC 2330 14 Aug Hellroaring Plateau, MT, (MONT) 2007 U.S.A. Alpine Salix Yes Yes R. nana1,2 CLC 3619 23 Aug Frozen Lakes, Beartooth Alpine Salix (MONT) 2017 Plateau, MT, U.S.A. reticulata Yes Yes R. nana1,2 21359 (DBG) 6 Aug Independence Pass, CO, 2003 U.S.A. Alpine Salix Yes Yes R. nana2 01943297 (NY) 12 Sept 1978 Scotland Salix herbacea Yes Yes (partial) R. nana2 F-127873 (O) 7 Jul Finse, Ulvik, Hordaland, Salix 1980 Norway herbacea Yes NA Alpine Salix R. nana2 EC 17.08.2011 17 Aug 2011 Belluno, Italy reticulata, S. Yes NA retusa R. nana2 CRN 177 (MONT) 23 Aug 2018 Niwot Ridge, CO, U.S.A. Alpine Salix Yes Yes R. nana2 TU 101701 NA NA NA KX579809 NA R. nana2 TU 101878 18 Sept 2006 Pori, Finland Betula UDB016029 NA R. nauseosa2 FH 12173 1 Aug Erfurt, Aspenbusch, 2012 Germany NA KT933985 KT933917 R. nigricans2 UE20.09.2004-7 NA NA NA DQ422010 NA R. nitida2 2-1148IS79 NA NA NA AY061696 NA R. nitida2 UPS:UE08.07.2004-2 NA Sweden NA KU205269 NA R. nitida2 FH 12218 18 Aug 2012 Hildesheim, Germany NA KT934001 KT933933 R. norvegica2 10183 5 Aug 2000 Italy NA JF908687 NA R. norvegica2 12163 23 Aug 2000 Italy NA JF908690 NA R. nuoljae2 UPS:UE24.08.200 23 Aug 4-09 2000 Lappland, Sweden Betula KU205270 NA 89 R. nuoljae2 UPS:UE23.08.200 23 Aug 4-10 2000 Lappland, Sweden Betula, Salix UDB002430 NA R. nuoljae2 UPS:UE25.08.200 24 Aug 4-06 2000 Lappland, Sweden Betula, Salix, Picea UDB002540 NA R. cf. 31 Jul Great Smoky Mountains Quercus, ochrophylla2 BPL 231 2012 National Park, TN, U.S.A. Tsuga KT933953 KT933883 R. pallescens2 146/2002 (TUR) NA Norway Picea, Pinus sylvestris DQ421987 DQ421941 R. pallidospora2 rw1901 14 Sept Bombergstraat, Turnhout, Quercus 1996 Belgium robur UDB002459 NA R. paludosa2 FH 12216 18 Aug 2012 Hildesheim, Germany NA KT934000 KT933932 CLC 1220 8 Aug Highline Trailhead, R. aff. pascua1 (MONT) 1998 Beartooth Plateau, MT, Alpine Yes Yes U.S.A. R. aff. pascua1,2 CLC 2274 1 Aug (MONT) 2006 Niwot Ridge, CO, U.S.A. Alpine Yes Yes R. aff. pascua1,2 CRN 138 (MONT) 8 Aug Frozen Lakes, Beartooth Alpine Salix 2018 Plateau, MT, U.S.A. reticulata Yes Yes Billings Fen Site, R. aff. pascua1,2 CRN 146 (MONT) 8 Aug Alpine Salix 2018 Beartooth Plateau, MT, U.S.A. reticulata Yes Yes R. pascua2 JV 32526F 14 Aug 2018 Sweden Salix Yes NA R. pascua2 IB:2005/1153 NA Tirol, Austria Salix, Polygonum KU205312 NA R. pascua2 IB:2004/0149 NA Dolomites, Italy Salix, Dryas KU205315 NA R. pascua2 KRAM:F-45052 NA S. Carpathians, Romania Dryas, Salix, Polygonum KU205327 NA Tsuga R. peckii2 BPL 270 8 Aug Great Smoky Mountains caroliniana, 2012 National Park, TN, U.S.A. Abies, Picea, KT933970 KT933901 Betula R. pubescens1,2 JV 31657F 24 Aug Picea, (MONT) 2016 Finland Betula, Pinus Yes NA sylvestris R. pulchra2 BPL 226 30 Jul Great Smoky Mountains Quercus, 2012 National Park, TN, U.S.A. Tsuga KT933951 KT933880 R. CLC 3820 22 Aug Crow Pass Rd. Girdwood, purpureofusca1,2 (MONT) 2018 AK, U.S.A. Salix, Dryas Yes Yes (partial) R. 19 Aug Loveland Pass, CO, Alpine Salix purpureofusca1,2 CRN 164 (MONT) 2018 U.S.A. planifolia Yes Yes R. 19 Aug Loveland Pass, CO, Alpine Salix purpureofusca1,2 CRN 165 (MONT) 2018 U.S.A. planifolia Yes Yes R. 22 Aug purpureofusca2 JV 32876F 2019 Sweden Dryas, Salix Yes NA R. H6042234 30 Aug Tromso, Troms, Norway Salix purpureofusca2 2013 herbacea UDB022579 NA R. 21 Aug Levanger, Nord- purpureofusca2 F-60573 (O) 1998 Trøndelag, Norway NA Yes NA Pinus, R. pusilla2 BPL 267 7 Aug Great Smoky Mountains Tsuga, 2012 National Park, TN, U.S.A. Quercus, KT933968 NA Betula R. aff. queletii1,2 CLC 1616 3 Aug Blue Lake Dam, CO, (MONT) 2001 U.S.A. Betula, Abies Yes NA R. queletii2 FH 12237 30 Aug 2012 Elendstal, Germany NA KT934007 KT933939 90 R. queletii2 CLC 1616 2 Aug 2002 Blue Lake, CO, U.S.A. Betula, Abies Yes NA R. queletii2 hue168 1 Oct Villingen, Baden-1998 Württemberg, Germany Picea UDB000316 NA R. renidens1,2 JV 31542F 15 Aug (MONT) 2016 Sweden Alpine Salix, Betula Yes NA R. renidens1,2 CLC 2871 27 Aug (MONT) 2012 Near Utsjoki, Finland Boreal, Betula Yes NA R. renidens2 UWBM:WTU-F- 24 Sept 038601 1983 Priest Lake, ID, U.S.A. NA KX813140 NA R. rubellipes2 BPL 240 1 Aug Great Smoky Mountains 2012 National Park, TN, U.S.A. Picea, Betula KT933958 KT933888 R. cf. rugulosa2 BPL 237 1 Aug Great Smoky Mountains 2012 National Park, TN, U.S.A. Picea KT933955 KT933885 Highline Trailhead, R. saliceticola1 CRN 125 (MONT) 7 Aug Beartooth Plateau, Alpine Salix 2018 MT/WY, U.S.A. reticulata Yes Yes Billings Fen Site, R. saliceticola1 CRN 134 (MONT) 7 Aug 2018 Beartooth Plateau, MT, Alpine Salix Yes NA U.S.A. R. saliceticola1 CRN 140 (MONT) 8 Aug Billings Fen Site, 2018 Beartooth Plateau, MT, Alpine Salix U.S.A. planifolia Yes NA Billings Fen Site, R. saliceticola1 CRN 144 (MONT) 8 Aug 2018 Beartooth Plateau, MT, Alpine Salix Yes NA U.S.A. planifolia R. saliceticola1 CLC 3575C 20 Aug Birch Site, Beartooth Salix (MONT) 2017 Plateau, MT, U.S.A. planifolia Yes NA R. saliceticola1,2 CLC 1137 29 Jul Birch Site, Beartooth Yes (MONT) 1997 Plateau, MT, U.S.A. Alpine Salix Yes (partial) R. saliceticola1,2 CLC 2370 6 Aug Wyoming Creek, (MONT) 2008 Beartooth Plateau, WY, Alpine Yes Yes U.S.A. Billings Fen Site, R. saliceticola1,2 CLC 3616 23 Aug Alpine Salix (MONT) 2017 Beartooth Plateau, MT, U.S.A. reticulata Yes NA Billings Fen Site, R. saliceticola1,2 CRN 143 (MONT) 8 Aug 2018 Beartooth Plateau, MT, Alpine Salix Yes Yes U.S.A. R. saliceticola1,2 CRN 155 (MONT) 10 Aug Highline Trailhead, 2018 Beartooth Plateau, MT, Alpine Salix U.S.A. planifoila Yes Yes R. saliceticola1,2 CRN 173 (MONT) 20 Aug Niwot Ridge, CO, U.S.A. Alpine Salix 2018 planifolia Yes Yes R. saliceticola1,2 CRN 184 (MONT) 24 Aug Alpine Salix 2018 Blue Lake, CO, U.S.A. planifolia Yes Yes R. saliceticola2 01782831 (NY) 30 Aug 1984 Albula Pass, Switzerland Alpine Yes (partial) NA R. saliceticola2 F-127988 (O) 4 Aug Holtålen, Sør-Trøndelag, Salix 1980 Norway herbacea Yes (partial) NA 27 Aug Finse, Ulvik, Hordaland, Salix R. saliceticola2 F-74342 (O) 2005 Norway herbacea, S. Yes Yes lanata R. saliceticola2 JV 32515F 13 Aug Arjeplog, Lycksele 2018 lappmark, Sweden Salix, Betula Yes NA R. saliceticola2 IB 05 150 NA NA NA NA NA R. sanquinea2 FH 12240 30 Aug 2012 Elendstal, Germany NA KT934008 KT933940 R. sardonia2 FH 12215 18 Aug Kolshorner Moor, 2012 Germany NA KT933999 KT933931 91 R. silvestris2 NL 15.09.08.av01 NA NA NA KX579800 NA R. silvestris2 NL 15.09.06.av04 NA NA NA KX579798 NA R. sphagnophila 256930 (MICH) 1936 St. Petersburg, Russia NA Yes NA R. 6 Sept Bay View, Emmet, MI, Pseudotsuga sphagnophila 12254 (MICH) 1905 U.S.A. menziesii, NA NA (Holotype)1 Abies, Picea R. CLC 3779 19 Aug Crow Pass Rd. Girdwood, Betula nana, sphagnophila1,2 (MONT) 2018 AK, U.S.A. Hemlock, Yes NA Picea R. subalpina 01919735 (NY) 24 Aug Eagle Summit, AK, Subalpine (Holotype)1 1976 U.S.A. tundra, NA NA Betula nana CLC 1218 8 Aug Highline Trailhead, R. subrubens1 (MONT) 1998 Beartooth Plateau, MT, Alpine Salix Yes Yes U.S.A. R. subrubens1 CLC 1219 8 Aug Highline Trailhead, (MONT) 1998 Beartooth Plateau, MT, Alpine Salix Yes Yes U.S.A. R. subrubens1 CLC 1382 19 Aug Birch Site, Beartooth (MONT) 1999 Plateau, MT, U.S.A. Alpine Salix Yes NA R. subrubens1 CLC 1464 6 Aug Independence Pass, CO, Alpine Salix Yes (MONT) 2000 U.S.A. planifolia Yes (partial) R. subrubens1 CLC 3597 22 Aug Frozen Lakes, Beartooth Alpine Salix (MONT) 2017 Plateau, MT, U.S.A. planifolia, S. Yes Yes glauca R. subrubens1 CLC 3601 22 Aug Frozen Lakes, Beartooth Alpine Salix (MONT) 2017 Plateau, MT, U.S.A. planifolia Yes Yes R. subrubens1 20848 (DBG) 3 Aug Independence Pass, CO, 2000 U.S.A. Salix Yes Yes R. subrubens1 CRN 137 (MONT) 7 Aug Billings Fen Site, 2018 Beartooth Plateau, MT, Alpine Salix Yes NA U.S.A. 10 Aug Highline Trailhead, Alpine Salix R. subrubens1 CRN 156 (MONT) 2018 Beartooth Plateau, MT, planifolia, S. Yes NA U.S.A. reticulata R. subrubens1 CRN 185 (MONT) 24 Aug 2018 Blue Lake, CO, U.S.A. Alpine Salix planifolia Yes NA R. subrubens1,2 CLC 1466 6 Aug Independence Pass, CO, Alpine Salix (MONT) 2000 U.S.A. planifolia, S. Yes NA glauca R. subrubens1,2 CLC 1488 9 Aug Loveland Pass, CO, (MONT) 2000 U.S.A. Alpine Yes Yes R. subrubens1,2 CLC 1666 7 Aug Mineral Basin, CO, U.S.A. Alpine Salix (MONT) 2001 arctica Yes Yes R. subrubens1,2 CLC 1719 11 Aug Black Bear Basin, CO, Alpine Salix (MONT) 2001 U.S.A. arctica Yes Yes R. subrubens1,2 CLC 1808 27 Jul Cinnamon Pass, CO, Alpine Salix (MONT) 2002 U.S.A. arctica Yes Yes R. subrubens1,2 CLC 3550 17 Aug Birch Site, Beartooth Alpine Salix (MONT) 2017 Plateau, MT, U.S.A. glauca Yes Yes R. subrubens1,2 CLC 3588 21 Aug Highline Trailhead, (MONT) 2017 Beartooth Plateau, MT, Alpine Salix U.S.A. planifolia Yes Yes 92 R. subrubens1,2 23519 (DBG) 6 Aug Independence Pass, CO, 2006 U.S.A. Alpine Salix Yes Yes 8 Aug Billings Fen Site, R. subrubens1,2 CRN 139 (MONT) Beartooth Plateau, MT, Alpine Salix 2018 U.S.A. reticulata Yes Yes R. subrubens1,2 CRN 174 (MONT) 20 Aug 2018 Niwot Ridge, CO, U.S.A. Alpine Salix planifolia Yes Yes R. subrubens1,2 CRN 187 (MONT) 24 Aug 2018 Blue Lake, CO, U.S.A. Alpine Salix planifolia Yes Yes R. subrubens2 JV 02-624 30 Sept 1998 NW Jutland, Denmark Salix, Alnus UDB002536 NA R. subrubens2 JV 98-294 26 Aug 1994 Central Jutland, Denmark Salix UDB002537 NA Quercus, R. subtilis2 BPL 275 9 Aug Great Smoky Mountains Pinus, 2012 National Park, TN, U.S.A. Fagus, KT933974 KT933906 Tsuga R. suecica1,2 JR 9092 (MONT) 6 Sept Finland Primeval 2013 forest Yes NA R. suecica2 MATC_N24 Jul 2007 Toolik Lake, AK, U.S.A. Betula nana GU997948 NA R. velenovskyi2 NL GM15c-061 NA NA NA KX579803 NA R. cf. versicolor2 FH 12259 2 Oct 2012 Walkenried, Germany NA KT934012 KT933944 R. vesicatoria2 PC0124666 NA TX, U.S.A NA KY800359 NA R. aff. vinosa1,2 CLC 3778 19 Aug Crow Pass Rd. Girdwood, (MONT) 2018 AK, U.S.A. Betula Yes NA R. vinosa2 500RUS26 NA NA NA AY061724 NA R. virescens2 HJB9989 NA Belgium NA DQ422014 DQ421955 R. xerampelina2 24138 (DBG) 4 Aug Independence Pass, CO, 2011 U.S.A. Picea Yes Yes R. xerampelina2 20531 (DBG) 1 Sept Mt. Goliath Nature Trail, 1999 Arapaho N.F., CO, U.S.A. Picea, Abies Yes Yes R. xerampelina2 S. Miller 9772 (2-684RUS28) NA Europe NA AY061734 NA R. zvarae2 FH 12175 1 Aug Erfurt, Aspenbusch, 2012 Germany NA KT933986 KT933918 DNA Extraction, PCR Amplification, DNA Purification, and Sequencing For molecular analyses, we extracted and amplified the ITS and RPB2 gene regions. The ITS region was chosen because it is the universal barcode for species identification in fungi (Schoch et al. 2012). The ITS region is 600–800 base pairs in length and present in multiple copies making it easy to amplify (White et al. 1990). However, because it is a multicopy gene, variation can exist between populations or individuals, which has the potential to complicate phylogenetic analyses (Smith et al. 93 2007). The ITS is a spacer region containing ITS1 and ITS2, which are transcribed but excised prior to translation, the highly conserved 5.8S gene sits between the two spacers (White et al. 1990; Gardes and Bruns 1993). These characteristics allow the ITS to undergo mutations freely, which makes the region variable between species, but conserved within species, making it useful for identification at the species level (Schoch et al. 2012). The RPB2 region was chosen to complement the ITS region because it is a conserved protein coding region; in general, these regions have more species resolving power, but are more difficult to amplify using PCR (Schoch et al. 2012). The RPB2 region encodes the second largest subunit of RNA polymerase II (Liu et al. 1999; Matheny 2005). The ITS-RPB2 combination has also been suggested as a target DNA barcode for Russula because it has sufficient intra- and inter-specific variation for species identification and high rates of successful PCR amplification (Guo-Jie et al. 2019). For the broad dataset described below, the nuclear ribosomal 28s large subunit (LSU) and the gene responsible for encoding the largest subunit of RNA polymerase II (RPB1) were included in the analysis; this data was extracted from Looney et al. (2016). LSU is a large gene region adjacent to the ITS and RPB1 is a slowly diverging, protein coding gene that is universally present (Tanabe et al. 2002). Both are commonly used in fungal phylogenetics (James et al. 2006). The ITS and RPB2 gene regions were amplified from dried basidiocarps, portions of which were placed in a clean 2ml screw cap tube. DNA was extracted following the procedure in the Sigma-Aldrich® Extract-N-Amp™ Plant PCR kit: manufacturer instructions were followed except a 1:2 (25:50 µm) ratio was used for the 94 extraction:dilution solution. A CTAB (Cetyltrimethylammonium bromide) extraction was used for older herbarium material or when extractions produced poor sequences using the previous method. This extraction method followed a slightly modified version of Nguyen et al. (2013) and used CTAB buffer to lyse cells followed by a chloroform extraction and ethanol precipitation. The CTAB extraction protocol with pertinent lab notes can be found in Appendix A. The ITS region was amplified using only the primers ITS1-F and ITS4 or a combination of primers ITS1-F and ITS2 and/or ITS3 and ITS4 to amplify the ITS1 and ITS2 regions separately (White et al. 1990; Gardes and Bruns 1993). When the ITS1 and ITS2 regions were amplified separately from the same collection each region was analyzed using a Standard Nucleotide BLAST® provided by NCBI (https://blast.ncbi.nlm.nih.gov/Blast.cgi) to check for the possible ITS contamination. If no contamination was present, the ITS1 and ITS2 regions were combined into one sequence for phylogenetic analysis and missing portions of the 5.8S region between them was filled with Ns (ambiguous nucleotides). For these collections, the alignment was carefully examined to ensure correct positioning within related sequences. When amplified separately the ITS1 and ITS2 regions will be uploaded separately to GenBank. Primer sequences are in TABLE 6. All primer sequences used were ordered from Integrated DNA technologies (https://www.idtdna.com/pages) and were prepared following the protocol in Appendix B. 95 Table 6. Primer sequences used in this study. Primer Sequence (5´– 3´) Reference ITS1-F CTTGGTCATTTAGAGGAAGTAA White et al. 1990, Gardes and Bruns 1993 ITS2 GCTGCGTTCTTCATCGATGC White et al. 1990, Gardes and Bruns 1993 ITS3 GCATCGATGAAGAACGCAGC White et al. 1990, Gardes and Bruns 1993 ITS4 TCCTCCGCTTATTGATATGC White et al. 1990, Gardes and Bruns 1993 fRPB2-7cR CCCATRGCTTGYTTRCCCAT Liu et al. 1999 bRPB2-6f TGGGGYATGGTNTGYCCYGC Matheny 2005 For the polymerase chain reaction (PCR) amplification (Mullis et al. 1986; Saiki et al. 1988), the reaction mix was combined in 0.2 ml ThermoGrid™ strip PCR tubes following Barge et al. (2016): 7.5 µl of molecular grade water, 12.5 µl of Extract-N-Amp PCR ReadyMix, 1.0 µl forward primer (10 µM), 1.0 µl reverse primer (10 µM), and 3.0 µl of fungal DNA. DNA amplification of the ITS region using PCR was done in a Thermocycler running with the following settings: 94 °C for 2 minutes, followed by 30 cycles of 94 °C for 30 seconds, 55 °C for 1 minute, and 72 °C for 1 minute, followed by a final 5-minute elongation step at 72°C. The RPB2 region was amplified using the primers fRPB2-7cR and bRPB2-6f (Liu et al. 1999; Matheny 2005), (TABLE 6). For PCR amplification of the RPB2 region, the following was combined in 0.2 ml ThermoGrid™ strip PCR tubes following Barge et al. (2016): 5.5 µl of molecular grade water, 12.5 µl of Extract-N-Amp PCR ReadyMix, 3.0 µl forward primer (10 µM), 3.0 µl reverse primer (10 µM), and 1.0 µl of fungal DNA. DNA amplification of the RPB2 region using PCR was done in a Thermocycler running with the following settings: 94 °C for 90 seconds, followed by 40 cycles of 94 °C for 30 seconds, 55 °C for 90 seconds, and 68 °C for 3 minutes, followed by a final 5-minute elongation step at 68°C. 96 PCR products were verified using gel electrophoresis. A 1.5% gel was made using molecular grade agarose and 1XTBE buffer, 0.5 µl of Ethidium Bromide (EtBr) or GelRed was added to bind the DNA so bands could be visualized under UV (ultraviolet) light. Next, 5–10 µl of PCR product was added to the gel, which was run for 1 hour at 96 volts and viewed with a trans-illuminator. Once PCR products were verified, PCR purification was performed using the QlAquick Kit (Qiagen) following the manufactures instructions. Purified PCR product concentrations were quantified using the spectrophotometer NanoDrop™ 2000. DNA for each gene was sent to Genscript® in Piscataway, New Jersey for sequencing. DNA was diluted to 2 ng/µl as specified by the “DNA Sequencing Sample Submission Guidelines – Genscript” and added to two 0.2 ml ThermoGrid™ strip PCR tubes, one containing 2.5 µl (10 µM) of the forward primer and another containing 2.5 µl (10 µM) of the reverse primer. Enough molecular grade water was added to each tube to bring the total volume to 15 µl. DNA was sequenced at Genscript® using Sanger DNA sequencing. Forward and Reverse complemented sequences were loaded into SeqTrace v. 0.8.1 (Stucky 2012); SeqTrace protocol can be found in Appendix C. Chromatograms were examined and contig sequences were constructed. Nucleotide sequences were trimmed at the ends until 8 out of 10 nucleotides were supported with at least a 30% confidence score. Ambiguous nucleotide positions were replaced by N’s. Positions where nucleotides were less ambiguous (could be 1 of 2 overlapping peaks) were coded as follows: W (A or T), S (C or G), M (A or C), K (G or T), R (A or G), or Y (C or T). NCBI BLAST® search was used for tentative identification of fungal isolates and to 97 check for similar sequences; sequences were then subjected to phylogenetic analysis. We searched for similar sequences in BLAST but did not add them to the phylogenies because of the need to examine morphology and take ecological data into account when delineating Russula species. Highly similar sequences in public databases were discussed under species observations when necessary. For particular species in the phylogenetic analyses, a UNITE species hypothesis was included in the phylogenetic analysis. UNITE is an online database that connects vouchered collections to well annotated sequences for ectomycorrhizal fungi (Kõljalg et al. 2005). A species hypothesis is species-level group that shares a certain degree of sequence similarity like an operational taxonomic unit. UNITE species hypotheses are included in groups containing significant taxonomic confusion or lacking reference specimens. Sequence Alignment and Phylogenetic Analyses The goal of the phylogenetic analyses was to determine phylogenetic placement of the species of Russula found in the Rocky Mountain alpine zone based on the genetic loci studied. The ITS and RPB2 sequences produced in this study were aligned using the online or desktop version of MUSCLE (http://www.ebi.ac.uk/Tools/msa/muscle/) (Madeira et al. 2019) using default parameters. For the desktop version of MUSCLE, an example of the code required to run the program can be found in Appendix D. Sequences were then inspected using PhyDE v.0.9971 (http://www.phyde.de/index.html) and misaligned regions were manually adjusted. Four phylogenetic trees were produced using the ITS-RPB2 data from this study. All datasets were analyzed using both the maximum 98 likelihood and Bayesian analysis methods described below. The phylogeny produced using the Bayesian method was selected to represent the tree because there were no significant differences in the topologies regarding the species clades studied here. All of the maximum likelihood phylogenies can be found in Appendix E. The bootstrap support from maximum likelihood and the Bayesian posterior probability was annotated onto each branch in the Bayesian phylogeny. A phylogeny representing a broad overview of the genus Russula, including eight well-recognized clades, was produced using select taxa from the multi-locus dataset (including ITS, LSU, RPB1, and RPB2 gene regions) published by Looney et al. (2016) in addition to ITS and RPB2 sequences generated in this study. The sequences of select taxa from Looney et al. (2016) were chosen to represent the relative diversity of the eight well-recognized clades within the genus Russula. The clade names used here follow Adamčík et al. (2019), and all represent the subgenera for which they are named. The Russula clade is divided into two large clades, the crown clade and core clade, which represent Russula subgenus Russula. The crown clade represents the large, highly diverse, apically resolved clade (FIGS. 2, 3). We realize that the use of crown clade here is inconsistent with the traditional use of crown clade defined as: a clade containing the most distant extant members of a group including the most recent common ancestor of that group and all its decendents (Budd and Jensen 2000). However, we use the term Russula crown clade to be consistent with recent literature (Looney et al. 2016; Adamčík et al. 2019; Buyck et al. 2020). The multi-locus dataset containing eight well-recognized clades within the genus Russula will be referred to as the broad dataset or phylogeny and 99 includes 61 taxa with the maximum number of characters being 4416; this alignment file along with all others will be uploaded to TreeBase. A multi-locus phylogeny that includes the ITS and RPB2 gene regions was also produced for the Russula crown clade (95 taxa and 1656 characters), Russula core clade (81 taxa and 1567 characters), and Brevipes clade (29 taxa and 1568 characters) in order to take a closer look at species relationships. Only the ITS and RPB2 data was used for these detailed datasets due to the lack of LSU and RPB1 sequences available for type and references collections relevant to this study. Collections included in the phylogenetic analyses are indicated in TABLE 5. Bayesian analysis was performed using MrBayes v. 3.2.2 (Ronquist et al. 2012). Throughout this project several multi-locus trees, including time trees, were also produced in Beast v.2.4.8 (Bouckaert et al. 2019); these data are not included in this thesis but the protocol for Beast can be found in Appendix F. For Bayesian, analysis all loci in each dataset were partitioned so that each locus or each codon position in a protein coding gene is allowed different rates of nucleotide evolution, which has been shown to improve tree topology (Kainer and Lanfear 2015). The broad dataset was divided into the following partitions: ITS, LSU, and the 1st, 2nd, and 3rd codon positions of RPB1 and RPB2. For Bayesian analysis, the Russula crown clade, Russula core clade, and Brevipes clade were divided into the following partitions: ITS and the 1st, 2nd, and 3rd codon positions of RPB2. The RPB1 and RPB2 regions were partitioned into the 1st, 2nd, and 3rd codon positions in Mesquite (Maddison and Maddison 2018) with the goal being to minimize stop codons in the beginning and middle of the coding region (Bazzicalupo pers. com. 2/1/19). This was done in Mesquite by selecting characters > list 100 of characters > select all > codon position > set codon position > and minimizing stop codons. The Display tab was selected followed by > color matrix cell > color nucleotide by Amino Acid. In the RPB2 alignment a large ambiguously aligned intron was removed prior to phylogenetic analysis to improve the overall alignment. The final ITS and RPB2 data sets were concatenated manually in Notepad++. Using PartitionFinder v.2.1.1 (Lanfear et al. 2012), the best substitution model was calculated for each partition in each dataset; the selected substitution models can be found in Appendix G. In MrBayes, the command block used the partitions set by PartitionFinder. Two independent runs of 30 million generations were sampled every 100 000 runs with the first 25% of samples discarded. The number of chains was set to four. Both independent runs reached likelihood stationarity and the convergence of runs was visually assessed using the Trace function in Tracer version 1.5 (Rambaut et al. 2014). The maximum likelihood (ML) analyses were performed in RAxML GUI version 1.5 (Silvestro and Michalak 2012). The alignment was loaded into RAxML as a PHYLIP file and analyzed as a partitioned data set. The broad dataset was again partitioned into the ITS, LSU, and each codon position of the RPB1 and RPB2 region; whereas, the detailed datasets were partitioned into the ITS and each codon position of the RBP2 region. Branch lengths were not calculated independently for each partition due to the obviously flawed topologies produced using this setting. The dataset was analyzed using the ML + rapid bootstrap option (default option) using the GTRGAMMAI model with 1000 bootstrap iterations. The BS brL option was not checked. The outgroup was 101 manually selected. The tree files from each analysis were viewed and edited in TreeGraph 2 (Stöver and Müller 2010) and annotated in Adobe Illustrator v 23.0.2. Maps of Russula Species Distributions Maps were created to visualize the observed, geographic distribution of each Russula species studied here. Maps only include collection locations that are confirmed in the phylogenetic analyses (FIGS. 3, 4). The maps were created in R Studio (https://rstudio.com/) using ggmap, which is part of the ggplot2 package (Wickham 2016). An example of the code used to create each map can be found in Appendix H. Results Phylogenetic Analysis In total 138 ITS and 83 RPB2 sequences were generated from 143 collections in this study (TABLE 5). An ITS1 and ITS2 sequence was generated for the R. montana Shaffer holotype (MICH 12231). The R. laveis holotype sequence (JR 4016) was included in the phylogenetic analyses (FIGS. 2, 5) (Adamčík et al. 2019). Sequences were not successfully obtained from the holotypes of R. sphagnophila Kauffman (MICH 12254) or R. subalpina O.K. Mill. (NY 01919735). Other types could not be obtained; however, sequences were generated or received for collections from the type locality or type country from reliable sources for most species. The alignments and multi-locus trees produced in this study will be uploaded to TreeBase. The topology of all the phylogenetic trees produced using the maximum likelihood and Bayesian phylogenetic methods were consistent in regard to the following 102 trends. Ten species of Russula were resolved from the Rocky Mountains. Nine of these species were resolved as monophyletic and eight species clades were strongly-supported. All analyses indicate that alpine species of Russula in the Rocky Mountains are distantly related from each other, and do not share a recent common ancestor that is unique to this group. All species studied were consistently resolved in either R. subgenus Russula or R. subgenus Brevipes. A few small differences in tree topology and support between the phylogenies produced here and other studies are worth noting. The topology leading to the clade containing R. altaica (Singer) Singer is weakly-supported and paraphyletic in the broad phylogeny (FIG. 2) and strongly-supported and monophyletic in the Russula core clade phylogeny (FIG. 4). This variation may be due to the fewer number of sequences included in this clade in the broad phylogeny compared to the Russula core clade phylogeny. The clade containing R. altaica, R. gracillima Jul. Schäff., and R. queletii Fr. forms a polytomy in the broad analyses with the remainder of the Russula core clade and the Russula crown clade (FIG. 2); whereas, in the Russula core clade phylogeny and in previous studies (Looney et al. 2016; Adamčík et al. 2019), this clade is strongly- supported and resolved within the Russula core clade. There was also some variation in the topology and weak-support for deeper nodes at the subgeneric level within the broad phylogeny (FIG. 2) compared to previous studies (Looney et al. 2016; Adamčík et al. 2019). For example, our analysis resolved the Crassotunicata and Heterophyllidia clades as the most basal clades of the genus Russula; whereas, Looney et al. (2016) and Adamčík et al. (2019) resolved the Brevipes clade as the most basal. We believe that the 103 broad phylogeny has varying topology and low-support for deeper nodes at the subgeneric level due to the absence of LSU and RPB1 sequence data for the species studied here and closely related reference sequences. Other studies (Looney et al. 2016; Adamčík et al. 2019; Buyck et al. 2020) produced strong-support for nodes at the subgeneric level using four or five loci including the LSU and RPB1 regions which are highly conserved throughout the genus Russula. A phylogeny representing a broad overview of Russula, including eight well- recognized clades, was produced using select taxa from the multi-locus dataset published by Looney et al. (2016) in addition to ITS and RPB2 sequences generated in this study. Multifurca ochricompacta (Bills & O.K. Mill.) Buyck & V. Hofst. (PC0723654) and Lactarius deceptivus Peck (AFTOL ID 682) were used as outgroups. The Russula crown and core clades in the broad phylogeny (FIG. 2) correspond to Russula subgenus Russula and the remaining six clades correspond to the subgenera for which they are named. The topology of the broad phylogeny is similar to that produced in recent taxonomic studies focused on Russula (Looney et al. 2016; Buyck et al. 2018; Adamčík et al. 2019). The broad phylogeny was used to determine the subgeneric placement of Russula species from the Rocky Mountain alpine zone. The Russula species found to occur in the Rocky Mountain alpine zone in this study occur in two subgenera: nine are in R. subgenus Russula and one is in R. subgenus Brevipes. 104 105 Figure 2. Bayesian phylogeny of the genus Russula produced using select sequences from the multi-locus dataset produced by Looney et al. (2016) in addition to ITS and RPB2 sequences generated in this study. Support values for bootstrap support (BS) and Bayesian posterior (PP) probabilities are indicated above or below branches. Thickened branches lead to clades receiving ≥ 75% bootstrap support (BS) and Bayesian posterior probabilities (PP) ≥ 0.95. Bolded tip labels represent Rocky Mountain alpine collections. AA = Arctic-alpine habitats. Each gray box is labeled with a well-recognized clade in the genus Russula following Adamčík et al. (2019). Bolded tip labels represent Rocky Mountain collections. AA = Arctic-alpine habitats. Subgenus Russula. The subgenus Russula consists of two large clades, the Russula crown clade and the Russula core clade (Looney et al. 2016; Adamčík et al. 2019; Buyck et al. 2020), which is confirmed here. A multi-locus phylogeny that included the ITS and RPB2 gene regions was produced for each clade to take a closer look at species relationships. Five species from the Rocky Mountain alpine (R. subrubens, R. cf. pascua, R. saliceticola, R. heterochroa, and R. purpureofusca) are in the Russula crown clade as represented in Adamčík et al. (2019) and R. fragilis Fr. (FH 12197) was used as the outgroup (FIG. 3). The five Russula species in the crown clade have primarily red and magenta pilei, ocher or yellow spore prints, and isolated or subreticulate spore ornamentation. All species studied in the crown clade are represented by at least two collections observed in this study and are supported in both phylogenetic analyses (FIG. 3). Russula subsection Xerampelinae forms a clade within the Russula crown clade and contains species with reddish-brown pilei, fishy odors, and a greenish reaction in the context when exposed to ferrous sulfate. This group includes the well- known edible R. xerampelina (Schaeff.) Fr., which has been recognized because of these unique characteristics since Singer (1938) (Adamčík and Knudsen 2004). 106 In the Russula crown clade, the analysis produced a weakly-supported clade containing species originally identified as R. saliceticola (Singer) Kühner ex Knudsen & T. Borgen from Switzerland (NYBG 01782831), Norway (F-74342), and Sweden (JV 32515F) along with several collections from the Rocky Mountain alpine zone. The R. saliceticola clade also contains two sequences labeled R. nitida (AY061696 and FH 12218). The R. saliceticola clade forms a polytomy within a strongly-supported clade that also contains R. nitida (Pers.) Fr. and R. sphagnophila Kauffman. Within subsection Xerampelinae, a strongly-supported clade contains collections identified as R. subrubens (J.E. Lange) Bon from the Rocky Mountains as well as a collection from western Jutland, Denmark (C JV02-624), the epitype locality of R. subrubens (Adamčík et al. 2016b). Also, within R. subsection Xerampelinae is a strongly-supported clade containing three collections identified as R. cf. pascua from the Rocky Mountain alpine that form a polytomy with members of the closely related R. clavipes complex (Adamčík et al. 2016b). The R. clavipes complex is known to have little molecular differentiation between three closely related species; our collections in this complex were identified based on the key in Adamčík et al. (2016b), which focuses on characters in the pileipellis and ecology. The Russula crown clade also contains a strongly-supported clade containing collections identified as R. purpureofusca Kühner from the Rocky Mountain alpine zone and reference material from Norway (F-60573) and Sweden (JV 32876F), including a collection (FIPUTO23) examined by Ruotsalainen and Huhtinen (2015) who recently synonymized R. purpurofusca with R. cupreola Sarnari based on examination of type material. The R. purpureofusca clade is sister to a strongly-supported clade 107 containing R. dryadicola R. Fellner & Landa and R. globispora (J. Blum) Bon, which have been referred to as the R. globispora lineage (Adamčík et al. 2019). One of our collections from the Rocky Mountains and two from Svalbard identified as R. heterochroa form a strongly-supported clade sister to a strongly-supported clade containing several sequences of R. cuprea (FH 12250, UDB002420, KU886591) and one sequence of R. gigasperma Romagn. (KU237787). 108 109 Figure 3. Bayesian phylogeny of the Russula crown clade combining ITS and RPB2 data. Support values for bootstrap support (BS) and Bayesian posterior (PP) probabilities are indicated above or below branches. Thickened branches lead to clades receiving ≥ 75% bootstrap support (BS) and Bayesian posterior probabilities (PP) ≥ 0.95. Bolded tip labels represent Rocky Mountain alpine collections. Each gray box indicates a species found in the Rocky Mountains. AA = Arctic-alpine habitats. Collections in quotation marks indicate questionable identifications. Four species from the Rocky Mountain alpine are placed in the Russula core clade as represented in Adamčík et al. (2019) and R. subtilis Burl. (BPL 275) was used as the outgroup (FIG. 4). These species all have red or magenta pilei, white to pale cream spore prints, an acrid or mild taste, and spores with various types of ornamentation. All species studied are represented by at least two collections in the phylogeny. Our Rocky Mountain alpine collections originally identified as R. laccata form a strongly-supported clade with collections from Norway (F-127987, F-73860, type country of R. norvegica D.A. Reid = R. laccata), Finland (JV 23194, UDB016024), Sweden (UDB000915, JQ724003), Italy (10183, 12163), and Canada (UBC:F30302). The R. laccata clade is sister to a strongly- supported clade containing R. atrorubens Quél. and is located within a strongly- supported, larger clade containing R. emetica (Shaeff.) Pers., R. silvestris (Singer) Reumaux, R. fragilis, and R. betularum Hora. The Russula core clade also includes collections originally identified as R. nana, which form a strongly-supported clade that includes specimens from the Rocky Mountain alpine zone as well as reference sequences from Italy (EC 17.08.2011, type country), Scotland (NYBG 1943297), and Switzerland (NYBG 1930708). The R. nana clade forms a polytomy with numerous collections originally identified as R. montana or R. grisescens (Bon & Gaugué) Marti. Within this polytomy, our analysis includes several collections identified as R. montana from 110 Colorado, including the holotype (MICH 12231) as well as several collections identified as R. grisescens from Austria (UDB031192) and Finland (JV 9788, JV32642F). All of the collections identified as R. montana or R. grisescens, except for one (F-127873, identified as R. nana from Norway with Salix) were collected in subalpine habitats, usually with conifers. Last in the Russula core clade is a weakly-supported clade identified as R. altaica, which contains two of our collections from the Rocky Mountain alpine zone along with a reference sequence from Russia identified as R. altaica (NYBG 00333753) by Singer, who described the species. The R. altaica clade contains species associated with Betula and is sister to a strongly-supported clade containing four sequences labeled R. gracillima, from Germany (FH 12264), Finland (JV 32758F), Estonia (UDB011284) and one without locality data (UE23.08.2004-14). The sequence from Estonia (UDB011284) represents the UNITE species hypothesis for R. gracillima. 111 112 Figure 4. Bayesian posterior probability phylogeny of the Russula core clade combining ITS and RPB2 data. Support values for bootstrap support (BS) and Bayesian posterior (PP) probabilities are indicated above or below branches. Thickened branches lead to clades receiving ≥ 75% bootstrap support (BS) and Bayesian posterior probabilities (PP) ≥ 0.95. Bolded tip labels represent Rocky Mountain alpine collections. Each gray box indicates a species found in the Rocky Mountains. AA = Arctic-alpine habitats. Collections in quotation marks indicate questionable identifications. Subgenus Brevipes. Russula subgenus Brevipes contains species with dense flesh, a white pileus that browns in age, a white spore print, and lamellae that are somewhat decurrent. One multi-locus phylogeny of the Brevipes clade, which represents subgenus Brevipes, was produced as represented in Adamčík et al. (2019) and R. nigricans Fr. (UE2009200407) was used as the outgroup (FIG. 5). The Brevipes clade contains sequences representing the UNITE species hypothesis for R. delica Fr. (UDB025023) and R. chloroides (Krombh.) Bres. (AF418605) along with the Rocky Mountain species R. laevis and several sequences with questionable identifications (FIG. 5). Russula subgenus Brevipes is known to contain species that have been difficult for mycologist to sort taxonomically (Buyck and Adamčík 2013). Collections originally identified as R. cf. delica from the Rocky Mountain alpine zone form a strongly-supported clade along with the holotype sequences of R. laevis (JR4106). Russula laevis is newly described (Adamčík et al. 2019) from Finland with Salix and represents an Arctic-alpine species. The sequence representing the UNITE species hypothesis for R. delica forms a clade by itself sister to the clade containing R. laevis and two collections labeled R. cf. brevipes. Within the Brevipes clade is the R. chloroides complex, which is a strongly-supported clade containing the sequence representing the UNITE species hypothesis for R. chloroides. Two sequences from Montana that were morphologically identified as R. 113 brevipes Peck are located within this clade along with several other collections labeled R. chloroides, R. brevipes, and R. delica in Genbank and UNITE. The R. chloroides complex includes sequences from Alaska, Montana, Belgium, North Carolina, Colorado, Canada, Germany, and Ireland, indicating an intercontinental distribution. Two collection labeled R. cf. brevipes are also included in R. subgenus Brevipes (MH714879 and KF007188), which form a strongly-supported clade sister to R. laevis. Figure 5. Bayesian posterior probability phylogeny of the Brevipes clade combining ITS and RPB2 data. Support values for bootstrap support (BS) and Bayesian posterior (PP) probabilities are indicated above or below branches. Thickened branches lead to clades 114 receiving ≥ 75% bootstrap support (BS) and Bayesian posterior probabilities (PP) ≥ 0.95. Bolded tip labels represent Rocky Mountain alpine collections. AA = Arctic-alpine habitats. Ecology of Russula in the Rocky Mountain alpine This study recorded potential ectomycorrhizal hosts near most fungal collections and some broad host patterns emerged. In alpine habitats of the Rocky Mountains the most common ectomycorrhizal hosts are dwarf (Salix reticulata and S. arctica) and shrubby (S. glauca and S. planifolia) Salix species (Cripps and Eddington 2005; Knudsen et al. 2012; Cripps and Horak 2008). In this study, a majority of the species in the alpine zone were associated with Salix. These include R. nana, R. laccata, R. purpureofusca, R. saliceticola, R. cf. pascua, and R. subrubens. In the Rocky Mountain alpine zone, R. laevis was commonly reported with Bistorta vivipara and Dryas octopetala, occasionally it was reported with Salix. Russula heterochroa was collected with Dryas in Colorado and with Salix polaris Wahlenb. and Dryas octopetala in Svalbard. Nine of the ten Russula species found in the Rocky Mountains were conspecific with European collections from Arctic and alpine regions. These European collections were reported in association with species of Salix that are not found in the Rocky Mountains including dwarf (S. polaris and S. herbacea L.) and shrubby (S. retusa L. and S. lanata L.) species. Taxonomy All descriptions are based on collections from the Rocky Mountain alpine zone. 115 Russula altaica (Singer) Singer FIGS. 6, 16D, 18D, 20D Lilloa 22:715 1951 (‘1949’) = Russula gracilis subsp. altaica Singer, Bull. trimest. Soc. mycol. Fr. 54(2):143 1938 = Russula gracillima f. altaica (Singer) Vassilkov, in Novin (Ed.), Ecologiya i Biologiya Rastenii Vo.vtochnoevropeskot Lesotundry, Pt. 1 (Leningrad):60 1970 Macromorphology. Pileus 10–40 mm wide, convex, then dished, occasionally umbonate, mottled olive yellow, olive green, grayish magenta, darker at center to umber or lighter to pinkish, smooth, greasy; margin not striate or indistinctly striate in age, can be lighter in color. Cuticle thick, gelatinous, separable except at center. Lamellae narrowly attached, crowded, L = 100–120 (n = 3), rather thick, white, graying somewhat, occasionally forking. Lamellulae absent. Stipe 20–30 × 5–15 mm, clavate or equal, white, with or without pink blush or strong pink coloring, slightly hoary. Context watery white, slightly graying, vinaceous beneath cuticle; stipe stuffed. Odor indistinct or absent. Taste mild, possibly bitter, and not hot. Exsiccata: Pileus with solid green or grayish ruby colors or mottled ruby, grayish ruby, olive green, greenish black, ocher, ruby under pileus cuticle, darker over disk or at margin. Lamellae deep yellow, yellow orange, browning or graying in age. Stipe pale yellowish, ocher, crème, with pink tints over entire surface, and brown stains near base. Chemical reactions: Not available. Micromorphology. Basidiospores reported as pale cream (IIa-IIb) (Knudsen et al. 2012; Sarnari 1998–2005), (7.1–)–7.4–7.9–8.5(–9.6) × (5.1–)5.8–6.1–6.5(–7.1) µm, broadly ellipsoid to ellipsoid, Q = (1.1–)1.2–1.3–1.4(–1.5); ornamentation of numerous, 116 isolated, conical, amyloid warts (rarely as spines), <0.5–1 um high, occasionally fusing, without thin lines or ridges, not reticulate; apiculus truncate and inamyloid; suprahilar plage medium, weakly amyloid. Basidia (38.1–)41.9–46.1–50(–58.4) × (7.6–)8–8.6– 9.3(–10.2) µm, clavate, 4-spored, some possibly 2-spored; basidiola slightly clavate. Hymenial cystidia (53.3–)61.2–72.4–83.6(–91.4) × (7.6–)8.7–9.7–10.7(–12.7) µm, clavate-pedicellate, apically subacute, often mucronate, with swollen, 5.1–8.9 µm long appendage; contents granular or crystalline, orange-brown, red-brown, brown to black in sulfovanillin. Pileipellis sharply delimited from underlying context; suprapellis weakly gelatinized, ca. 38–76 µm thick, composed of loosely to tightly interwoven, generally erect, hyphae and pileocystidia, well-defined or gradually transitioning to subpellis; subpellis strongly gelatinized, ca. 50–64 µm thick, composed of horizontally oriented, hard to differentiate hyphae. Hyphal terminations clavate or cylindrical, occasionally tapering, ca. 2.5–4 µm wide at apex, apically obtuse, difficult to observe. Pileocystidia near the pileus margin rarely septate, cylindrical to slightly clavate, 57.3–134.5–211.72(– 411.5) × (3.8–)4.7–5.5–6.2(–7.6) µm, apically subacute or obtuse, occasionally forking; contents gelatinous, granular, or crystalline, more granular and orangish red in Congo Red, blackening in sulfovanillin. Pileocysitidia at center slightly longer, otherwise similar, (63.5–)66.3–155.3–244.3(–467.4) × 4.7–5.7–6.7(–8.9) µm, rare in some areas, abundant in others, occasionally forking below apex. 117 Figure 6. Russula altaica (CLC 1618). A. with Betula, Blue Lake Dam, Colorado. B. same collection photographed on a gray card. Scale = 2 cm. 118 North American Ecology. Likely with Betula glandulosa at treeline in Colorado, also with Salix and Dryas species nearby. Aug. Material Examined. USA. COLORADO. Summit County, Blue Lake Dam, with Betula glandulosa and Dryas, 2 Aug 2001, C. Cripps CLC 1608 (MONT); loc. cit., with Betula and Salix species, 3 Aug 2001, C. Cripps CLC 1618 (MONT). RUSSIA. Arkhangelsk, Ostrov Artura, with Betula, Salix, and Carix, 5 Aug 1937, R. Singer and Vasilieva 00333753 (NYBG). Observations. Singer described Russula gracilis subsp. altaica, from the Altai Mountains of central Asia with Betula and Dryas in 1938, and later designated it R. altaica (Singer) Singer (1951 ’1949’). Singer’s species was subsequently reported with Salix and Betula in Arctic and alpine habitats of Greenland (Knudsen and Borgen 1982; Lamoure et al. 1982; Borgen 1993) and Svalbard (Kobayasi et al. 1968; Skifte 1989). Knudsen and Borgen (1982) believed that Kühner’s (1975) description of R. amoenipes Romagn. was synonymous with R. altaica (Sing.) Sing., and if true this would extend this species distribution to the Alps and Swedish Lapland; however, this conclusion has been rejected by others (Singer 1986, Sarnari 1998–2005). Kobayasi et al. (1967) reported R. fragilis in wet tundra in Alaska during August but notes that this species is similar to R. altaica (Sing.) Singer (1951). More recently, Russula altaica has been reported in Arctic and alpine habitats in Greenland (Borgen et al. 2006), Canada (Ohenoya and Ohenoya 2010), and the Alps (Moser and Jülich 1985–2005). A similar species, Russula gracillima, was described by Schäffer in 1931 from Germany and it is well known in Europe from low elevations near Betula. Some consider 119 Russula altaica to be merely a synonym of R. gracillima, although significantly, Sarnari (1998–2005) and Romagnesi (1967) do not. Russula gracillima is associated with Betula (Adamčík et al. 2006) and has been reported based on morphology in Slovakia (Adamčík et al. 2006), Greenland (Borgen et al. 2006), and Iceland (Christiansen 1941, Hallgrimsson 1998). Geml et al. (2010) sequenced soil cores from Alaska and reported R. gracillima based on only 97% sequence similarity to data available in public databases. Both R. gracillima and R. altaica are reported to have a variable pileus color with combinations of red, dark red, purple, olive green, and ocher, a pink-flushed stipe, a weakly acrid taste, indistinct odor, and light-colored spores with isolated warts; this matches our collections except that we reported a mild taste. Our mature specimens are notable for olive-green and purple colors, unlike other Russula species in the Rocky Mountain alpine, however young specimens were dark reddish purple. Moser and Jülich (1985–2005) show a much redder pileus for R. altaica, and a resemblance to the dark red subalpine species R. queletii has been noted (Knudsen and Borgen 1982). However, Greenland collections also have green, pink, and gray coloration (Knudsen and Borgen 1982). Russula altaica may have slightly wider spores (7–8.5(10) × 6–7.8(8.5) µm) in the original latin description compared to R. gracillima (7.2–8.5 × 5.2–6.5 µm) (Romagnesi 1967). Our phylogenetic analysis produced a strongly-supported clade with two clusters: one with sequences labeled as R. gracillima (DQ422004, AY061678, and KR364094), and the second with two collections from Colorado along with a reference sequence from Russia identified as R. altaica by Singer himself (00333753), although in the broad 120 phylogeny our collection (CLC 1618) was sister to the clade containing R. altaica (NYBG 333753) and R. gracillima (DQ422004) (FIG. 2). Singer’s collection was found with Salix and Betula on an Arctic Island near Russia. In the alignment our collections from the Rocky Mountain alpine have eight conserved base pairs in the ITS1 and ITS2 regions that are not shared with the R. altaica sequence, most of which are also different from the R. gracillima sequences. The average spore size of Singer’s collection is 8 × 6.4 µm and the spore size and morphology match that of our collections from the Rockies. Given the data presented above we elect to call our collections R. altaica at this time. There are no sequences of R. altaica in Genbank or UNITE, and the type is in Russia. Our reference collection (00333753, NYBG) was collected and determined by Singer was collected in 1937 and will be the first publicly available sequence of this species. This is the first report of R. altaica from the Rocky Mountains. Interestingly, there is a known floristic relationship between the Altai and the Southern Rocky Mountains (Weber 2003) and we hypothesize that there may be a mycological connection as well. There is a possibility that R. altaica is synonyms with R. gracillima, but the type material of R. gracillima needs to be examined. For now, we elect to call our Rocky Mountain collections R. altaica due to the morphological similarities and phylogenetic clustering with Singer’s R. altaica collection. Our collections and Singer’s were found near Betula. Betula glandulosa is present in the Rockies but rare on most of the sites sampled, except for a few shrubs on the Beartooth Plateau and in a few areas of Colorado such as near Blue Lake Dam where our collections were found at treeline. This is the only Russula we report with Betula from 121 these areas, however, our Alaskan collections of red-capped Russulas are primarily with Betula, and this includes R. cf. alpigenes (Bon) Bon, R. purpureofusca, R. cf. intermedia P. Karst., R. sphagnophila, and R. vinosa Lindblad. These should be considered when collecting in the Rocky Mountains near Betula (see Addendum). Russula heterochroa Kühner FIGS. 7, 17D, 18I, 21B Bull. trimest. Soc. mycol. Fr. 91(3):389 1975 Macromorphology. Pileus 20–40 mm wide, convex to shallow convex with or without slight depression, mottled dark red brown, ruby, dark magenta, ocher and yellow, can be pale but disk remaining dark, viscid, smooth, not striate or indistinctly so; cap cuticle separable except at center. Lamellae narrowly attached, adnate, on av. 100–120 L (n = 3), whitish to medium yellow cream, somewhat thick, occasionally forking. Lamellulae absent. Stipe 25–40 × 10–20 mm, clavate or equal, central, white, with or without faint pink tint. Context white, pink underneath cap cuticle. Odor fruity or very faintly fishy. Taste mild not hot. Exsiccata: Pileus orange brown to dark brown, nearly uniform in color. Lamellae light yellow to darker orange yellow. Stipe white, staining yellowish near base. Chemical reactions: Not available. Micromorphology. Basidiospores, no print obtained, but R. heterochroa reported as cream yellow (IIIc) (Ronikier 2008), (8.6–)9.4–10.5–11.6(–13.6) × (6.6–)7.7–8.4– 9.2(–9.6) µm, broadly ellipsoid, Q = (1.0–)1.2–1.2–1.3(–1.5); ornamentation of numerous isolated amyloid warts (spines present but rare), 0.5–1 µm high, occasionally fusing, with no reticulation; apiculus truncate and inamyloid; suprahilar plage medium to large, 122 amyloid. Basidia (53.3–)55.6–62.1–68.5(–73.66) × (10.2–)11.3–12.1–12.9(–14) µm, clavate-pedicellate, primarily 2-spored, occasionally 1- and 4-spored; basidiola clavate. Hymenial cystidia (58.4–)70.9–89.7–108.6(–152) × (10.2–)10.8–12.1–13.5(–15.2) µm, fusiform, occasionally clavate, pedicellate, often mucronate, with 2.5–5.1 um long appendage; contents granular, gelatinous or absent, no reaction in sulfovanillin. Pileipellis sharply delimited from underlying context; suprapellis ca. 50–115 µm thick, composed of smooth, tightly interwoven, erect hyphae and embedded pileocystidia, well- defined or gradually transitioning to subpellis; subpellis gelatinized, ca. 125–177 µm thick, composed of hard-to-differentiate horizontally oriented hyphae. Hyphal terminations cylindrical to clavate, thin-walled, ca. 2–4.5 µm wide at apex, apically obtuse or subacute; rarely filled with granular contents. Pileocystidia near pileus margin abundant, with 1–2 septa, clavate to cylindrical, (53.3–)53.5–101.1–148.7(–228.6) × (3.8–)3.6–5.5–7.5(–14) µm, rarely forking, occasionally diverticulate (with small lateral protrusions); contents lightly granular and gelatinous, blackening in sulfovanillin. Pileocystidia near center rare, on average longer and thinner, (86.4–)94.7–127.8–161(– 190.5) × (3.8–)3.5–5.1–6.7(–8.9) µm, similar in shape. 123 Figure 7. Russula heterochroa (CLC 1723). Scale bar = 2 cm. North American Ecology. In alpine zone with Dryas octopetala in Colorado. Aug. Material examined. SVALBARD. Todalen, Adventdalen, Nordensklods Land, with Salix polaris and Dryas octopetala, 21 Aug 2002, C. Cripps CLC 1918 (MONT); loc. cit., with Salix polaris and Dryas octopetala, C. Cripps CLC 1919 (MONT). USA. COLORADO. Gunnison/Chaffee Counties, Sawatch Range, Cottonwood Pass, with Dryas, 12 Aug 2001, C. Cripps CLC 1723 (MONT). Observations. Russula heterochroa was originally described by Kühner (1975) as an alpine species from the French Alps and Swedish Lapland. Kühner (1975) described three new species, R. purpureofusca, R. heterochroa, and R. saliceticola, and placed all in section Tenellae Quélet. Russula heterochroa is recognized by a dark purple pileus, white 124 to cream lamellae, white stipe, large yellow cream spores with primarily isolated warts, and 2-spored basidia (Kühner 1975, Ronikier 2008). Our collections from the Rocky Mountains of Colorado and the Arctic island of Svalbard match the morphology of R. heterochroa. No sequences of R. heterochroa are currently available in public databases. This species has been previously reported from the French Alps (Kühner 1975), Swedish Lapland (Kühner 1975), and the Carpathians (Ronikier 2008). This is the first report of this species in North America and in Svalbard. Our collections of this species from Svalbard (CLC 1918 and CLC 1919) have diverticulate pileocystidia, similar to the R. purpureofusca; these are visible but less abundant in the collection from Colorado (CLC 1723). Similar species morphologically in the Rocky Mountain alpine include R. laccata, R. saliceticola, and R. purpureofusca, which all have predominately purple or magenta pilei. However, the 2-spored basidia and the larger spores (av 10.5 × 8.4 µm) are unique to R. heterochroa, and all three species can be separated phylogenetically using the ITS region. Phylogenetically, R. heterochroa is located in the Russula crown clade. Our three collections (CLC 1723, CLC 1918, and CLC 1919) form a strongly-supported clade sister to R. cuprea, which contains one sequence from Germany and another from the United Kingdom. The sequence from Germany (FH12250) lacks host or ecological data; the sequence from the United Kingdom (UDB002420) was found near Quercus petraea and Betula species. 125 Russula laccata Huijsman FIGS. 8, 16C, 18C, 20A Fungus, Wageningen 24:40 1955 = Russula norvegica var. rubromarginata Kühner, Bull. trimest. Soc. mycol. Fr. 91(3):389 1975 = Russula norvegica D.A. Reid, Fungorum Rariorum Icones Colorate 6:36 1972 = Russula atrorubens Quél. sensu J.E. Lange, Fl. Agar. Dan.:63 1940 – pro parte. Macromorphology. Pileus 15–45 mm wide, convex (shallow or broad), almost applanate, or uplifted with slight central depression, occasionally umbonate, deep magenta, dark magenta, and ruby, with a dark, almost black center, but also with a mix of deep magenta, grayish magenta, vivid red, deep red, rarely with yellow brown, typically with dark purple-black center fading to lighter magenta near margin, occasionally darker at margin and lighter over center, aging dark brown, smooth, greasy, viscid, can be shiny, tacky or dry, with occasional debris adhering; margin short or long striate, typically more striate in age, occasionally not striate or indistinctly so, undulating or entire, incurved when young; cuticle separable except at center. Lamellae attached, adnate to adnexed, relatively close to distant, on av. L = 70–110 (n = 8), white, off-white, cream, pale yellow tinge in some, rarely graying, yellow brown near cap margin, rarely deep magenta or ruby near pileus margin, anastomosing or not, occasionally forking near stipe, elastic not brittle, thick to thin; lamellae edge entire or undulate. Lamellulae absent or rare. Stipe 15–50 × 5–15 mm, equal, clavate, sometimes tapering near base, central or slightly eccentric, white, off-white, translucent pale yellowish, grayish cream, base occasionally with pink flush, or red brown or grayish staining, smooth or rarely powdery, fragile, with 126 indistinct vertical ridges. Context white; stipe spongy, firm or brittle, not hollow but with small to medium cavities. Odor absent, fragrant, or fruity. Taste mildly to strongly acrid or burning, immediate or delayed. Exsiccata: Pileus light buff, brown, grayish ruby, deep ruby, grayish magenta, deep magenta, mottled with ocher tones or uniformly one color, usually darker to black in center. Lamellae light golden, ocher, yellow, straw yellow, browning near cap margin, rarely with hints of gray. Stipe white, off-white, yellow, ocher, staining brownish gray, occasionally with magenta or pinkish blush. Chemical reactions: Gum Guaiac = blue green, red brown, brown; ferrous sulfate = pinkish gray, pinkish brown; 2% phenol = pale yellow. Micromorphology. Basidiospores white in print (Ia-Ib), (6.3–)7.4–8.1–8.9(–11.6) × (5.1–)5.7–6.2–6.8(–9.6) µm, broadly ellipsoid to ellipsoid, Q = (1–)1.2–1.3–1.4(–1.7); ornamentation of amyloid ridges forming a complete reticulum connecting raised warts and spines <0.5 – 1 µm high, occasionally with fine line connections, rarely composed of isolated warts or spines that can fuse to form angular elongated structures; ornamentation of abnormally large spores subreticulate with more isolated elements; apiculus truncate or pointed, inamyloid; suprahilar plage small to large, amyloid, occasionally difficult to observe. Basidia (30.5–)38.2–44.9–51.6(–71.1) × (6–)7.5–8.7–10(–11) µm, clavate- pedicellate, occasionally fusiform, 4-spored, some possibly 2-spored; basidiola clavate. Hymenial cystidia (40.6–)50.7–67.7–84.6(–114.4) × (6–)7.5–9.3–11.1(–16) µm, fusiform, typically pedicellate, often mucronate, with cylindrical or swollen, occasionally septate, 2.5–5 µm long appendage (rarely up to 10.2 µm); contents granular or gelatinous, immediately blackening in sulfovanillin. Pileipellis gradually transitioning to underlying 127 context; suprapellis ca. 96–140 µm deep, composed of densely interwoven, erect hyphae, gradually transitioning to subpellis; subpellis gelatinized, ca. 101–152 µm deep, composed of parallel, vertical or horizontally oriented, septate hyphae. Hyphal terminations clavate, thin-walled; ca. 1.5–3.5 µm wide at apex, apically obtuse, and devoid of contents. Pileocystidia near pileus margin abundant, septate, filiform, clavate or cylindrical, pedicellate, (30.6–)43.8–79.6–115.4(–218.4) × (3–)4.7–6–7.2(–9.6) µm, swollen clavate or acute at apex, occasionally forking near base, occasionally swollen between septa; contents granular and refractive gelatinous, slightly blackening in sulfovanillin. Pileocystidia near the pileus center shorter, similar in shape, contents similar but also crystalline, rarely forking, (30.5–)40–49.8–59.6(–63.5) × (3.8–)4.4–5.1– 5.9(–7.6) µm. 128 Figure 8. Russula laccata. A. with Salix reticulata, Beartooth Plateau, Montana (CRN 150). B. with Salix planifolia and S. reticulata, Niwot Ridge, Colorado (CRN 169). C. with Salix species, Beartooth Plateau, Montana (CRN 114). D. with S. reticulata, Beartooth Plateau, Montana (CRN 157). E. with S. reticulata, Beartooth Plateau (CRN 128). F. with S. planifolia from Blue Lake, Colorado. Scale bar = 2 cm. North American Ecology. In alpine areas of Montana, Wyoming, and Colorado with Salix planifolia and S. reticulata, often among mosses. Habit: solitary, scattered, or gregarious. Aug. 129 Material examined. FINLAND. Koillismaa, Kuosamo, Oulanka National Park, with Salix, 27 Aug 2017, J. Vauras JV 23194. USA. COLORADO. Boulder County, Niwot Ridge, 1 Aug 2005, C. Cripps CLC 2275 (MONT); loc. cit., with Salix planifolia and S. reticulata, 20 Aug 2018, C. Noffsinger CRN 168 (MONT); loc. cit., with S. planifolia, 21 Aug 2018, C. Noffsinger CRN 175 (MONT); Blue Lake, with S. planifolia, 24 Aug 2018, C. Noffsinger CRN 186 (MONT); Clear Creek/Summit Counties, Front Range, Loveland Pass, 9 Aug 2000, C. Cripps CLC 1487 (MONT); loc. cit., with dwarf Salix, 21 Aug 2003, V. Evenson DBG-F-021608 (DBG); loc. cit., with Salix, R. Brace DBG-F-021511 (DBG); loc. cit., with S. planifolia, 19 Aug 2018, C. Noffsinger CRN 166 (MONT); Pitkin/Lake Counties, Sawatch Range, Independence Pass, 6 Aug 2000, C. Cripps CLC 1465 (MONT); loc. cit., 6 Aug 2000, C. Cripps CLC 1467.1 (MONT). MONTANA. Carbon County, Beartooth Plateau, Birch Site, with shrubby Salix, 9 Aug 1999, C. Cripps CLC 1378 (MONT); loc. cit., with S. planifolia, 19 Aug 1999, C. Cripps CLC 1381 (MONT); Highline Trailhead, with S. reticulata, 7 Aug 2018, C. Noffsinger CRN 128 (MONT); loc. cit., with S. reticulata, 10 Aug 2018, C. Noffsinger CRN 157 (MONT). WYOMING. Park County, Beartooth Plateau, Solufluction Terraces, with Salix species, 6 Aug 2018, C. Noffsinger CRN 114 (MONT); Wyoming Creek, 6 Aug 2008, C. Cripps CLC 2371 (MONT); Billings Fen site, with dwarf Salix, 23 Aug 2017, C. Cripps CLC 3617 (MONT); loc. cit., with Salix species, 7 Aug 2018, C. Noffsinger CRN 133 (MONT). Observations. Russula laccata was originally described from the Netherlands, often in association with Salix and Betula in swampy habitats with Sphagnum (Huijsman 130 1955). It was thought to be closely related to R. olivaceoviolascens Gillet in section Violaceae Rom. (Huijsman 1955), or merely a color variant of this species (Romagnesi 1967). Later, R. norvegica Reid (and R. norvegica var. norvegica) was described from alpine habitats in Norway with Salix herbacea (Reid 1972), and this name was commonly applied to Arctic and alpine collections fitting this description. Ortega and Esteve- Raventós (2001) examined the type material of R. laccata and R. norvegica and synonymized the two species giving R. laccata priority. The species is recognized by a dark magenta pileus with black center, white lamellae, white stipe, white reticulate spores, mild to strong acrid taste, and association with Salix. In the Rocky Mountain alpine, it can be confused with R. saliceticola which has more red coloration in the pileus, lacks a hot taste, has larger yellow spores with mostly isolated warts, and more pink on the stipe; and R. purpureofusca which has larger ocher spores with mostly isolated warts and a somewhat hot taste. Russula nana and R. montana also have whitish spores and a hot taste but are known for bright red pilei without the dark center. In rare cases, R. laccata specimens can be pale in the center and thus, can be confused with R. nana. Our North American alpine collections of R. laccata fit the morphology of this species and form a strongly-supported phylogenetic clade with collections from Norway (type country of R. norvegica), Finland, Canada, and Sweden. Phylogenetically, R. laccata is located in the Russula core clade and is a sister group of R. atrorubens. Molecularly, R. laccata, R. saliceticola, R. purpureofusca, R. nana, and R. montana can all be separated using the ITS region alone in a phylogenetic analysis. 131 In the literature, R. laccata is well known from Arctic and alpine habitats in Greenland, Iceland, Fennoscandia, the European alps (Watling 1977; 1983; Knudsen and Borgen 1982; Lamoure et al. 1982; Gulden et al. 1985; Graf 1994; Bon and Noguera 1995; Hallgrimsson 1998; Gulden 2005; Borgen et al. 2006), Scotland (Watling 1987), the Netherlands (Huijsman 1955), East Frisian Islands of Germany (Bresinsky 1987), the Pyrenees (Bon and Ballarà 1995), Slovakia (Fellner and Landa 1993b), Japan (Nara et al. 2003), Russia (Knudsen and Muhkin 1998) and Svalbard (Skifte 1989), but these reports have not been molecularly confirmed. Russula laccata was also reported in montane habitats with Salix atrocinerea Brot. in Spain (Ortega and Esteve-Raventós 2001). It was first reported in North America from Schefferville, Northern Québec (Hutchison et al. 1988) and later reported from the Rocky Mountain alpine zone (Cripps and Horak 2008). This is the first molecularly confirmed report of R. laccata in the Rocky Mountain alpine and based on our phylogenetic analysis, this species has an intercontinental distribution in Arctic and alpine habitats including North America and Europe. There is some debate regarding taxa closely related to R. laccata and their synonymy. Kühner (1975) described a variety from alpine habitats, R. laccata var. rubromarginata, with dwarf willows. Sarnari (1998–2005) recognized R. fragilis var. alpestris Boudier, R. norvegica Reid, and R. norvegica var. rubromarginata Kühner as synonyms of R. laccata. However, some sources synonymize R. fragilis var. alpestris Boudier with R. emetica (Schaeff.) Pers. and Ortega and Esteve-Raventós (2001) only recognized R. norvegica D.A. Reid and R. atrorubens sensu Lange as synonyms of R. 132 laccata. The collections of R. atrorubens included in our phylogeny represent R. atrorubens Quél, a valid species that is sister to R. laccata. Russula laevis Kälviäinen, Ruotsalainen & Taipale FIGS. 9, 16E, 18H, 21A Fungal Diversity: vol 99:413–414. 2019 Names misapplied: Russula delica Fr., Epicr. syst. mycol. (Upsaliae):350 1838 [1836–1838] Macromorphology. Pileus 40–80 mm wide, concave, shallow convex, depressed in center, infundibuliform, pure white, dingy white, cream, mottled with brownish or orange brown stains in age or when bruised, smooth, greasy, kidskin, or finely tomentose (use hand lens), often with debris adhering to surface; margin strongly incurved or rolled under at first, then turned down, undulating, entire; cuticle can overhang gills. Lamellae attached, adnexed to narrowly-adnate, subdecurrent, occasionally toothed, close, on av. 60–160 (n = 4), sometimes forking, flaking when handled, white to very pale yellowish white, occasionally with slight yellow green, greenish, or greyish cast; edges concolorous. Lamellulae present or absent. Stipe 10–30 × 10–20 mm, equal, short, squared off at base, white, staining ocher, matt, minutely pubescent at apex (use hand lens) and smoother below, can be slightly bumpy. Context white, firm, solid, compact, slightly ocher under cuticle. Odor fruity or sweet, unpleasant in older specimens. Taste hot, acrid, occasionally slowly acrid. Exsiccata: Pileus mottled mostly with brown and orange brown tones, with a few white, buff, ocher, or gray areas. Lamellae ocher, deep orange brown, yellow brown, and light brown. Stipe off white or buff near apex, brown 133 or dark brown towards base, or mottled white, buff, and brown. Chemical Reactions: Not available. Micromorphology. No print obtained, but reported as crème (IIb-d) (Adamčík et al. 2019). Basidiospores (7.1–)7.9–8.7–9.4(–13.3) × (6.1–)6.6–7.2–7.8(–10.2) µm, subglobose or broadly ellipsoid, Q = (1.1–)1.1–1.2–1.3(–1.4); ornamentation composed of numerous or sparse, isolated to subreticulate, amyloid warts and spines, 0.5–1 µm high, rarely up to 1.5 µm, occasionally fusing into large angular masses, occasionally connected by lines or ridges; apiculus truncate, occasionally obtuse, inamyloid; suprahilar plage small to large, weakly amyloid. Basidia (58.4–)61.4–68.7–75.9(–88.9) × (8.9–)9.5– 10.5–11.4(–12.7) µm, fusiform, clavate, constricted in middle, pedicellate, 4-spored, rarely 2-spored; basidiola clavate. Hymenial cystidia (58.4–)70–82.9–95.8(–106.7) × (5.1–)6–7.1–8.1(–8.9) µm, cylindrical, fusiform, pedicellate, subacute at apex, or swollen, rarely mucronate with swollen or cylindrical 2.5–7.6 µm appendage, occasionally forking; contents granular or crystalline, graying in sulfovanillin. Pileipellis sharply delimited from underlying context, not stratified, gelatinized, ca. 88–140 µm deep, composed of horizontally oriented hyphae, becoming denser near context. Hyphal terminations cylindrical or clavate, flexuous, thin-walled, occasionally swollen, ca. 3–7 µm wide at apex, forking or not, apically obtuse or subacute, devoid of contents. Pileocystida near the pileus margin clavate, fusiform, cylindrical, occasionally forking, with one septa or not, occasionally constricted at septa, (20.3–)21.5–66.7–88.2(–190.5) × (2.5–)3–5–8(–10.2) µm, apically acute, with one or two ca. 2.5 µm long appendages; contents lightly granular, with no reaction in sulfovanillin. Pileocystidia near pileus 134 center more cylindrical, less granular, rarely forking, occasionally tapering, (30.5–)35.1– 67–98.9(–139.7) × (3.7–)3.8–4.3–4.9(–5.1) µm. Figure 9. Russula laevis. A. with Dryas octopetala, Cottonwood Pass, Colorado (CLC 1642). B. with Salix reticulata, Stony Pass, Colorado (CLC 1690). C. with D. octopetala, Cottonwood Pass, Colorado (CLC 1642). D. with S. arctica and S. glauca, Summit Lake, Mount Evans, Colorado (DBG-F-027685), photo Denver Botanical Garden. Scale bar = 2 cm. North American Ecology. In alpine areas, often slightly buried in Salix or Dryas octopetala debris. Aug. Material examined. SVALBARD. Longyearbyen, with Salix polaris, 18 Aug 2002, C. Cripps CLC 1883 (MONT). COLORADO. Boulder County, Niwot Ridge, with dwarf Salix, 17 Aug 1997, C. Cripps CLC 1146 (MONT); loc. cit., with Bistorta vivipara, 1 Aug 2006, C. Cripps CLC 2271 (MONT); San Juan County, San Juan 135 Mountains, Stony Pass, with S. reticulata and S. arctica, 9 Aug 2001, C. Cripps CLC 1690 (MONT); Pitkin County, Sawatch Range, Independence Pass, with Dryas octopetala, 13 Aug 2001, C. Cripps CLC 1740 (MONT); Chaffee/Gunnison Counties, Sawatch Range, Cottonwood Pass, with D. octopetala and S. glauca, 4 Aug 2001, C. Cripps CLC 1642 (MONT); Clear Creek County, Mount Evans, Summit Lake, with S. arctica and B. vivipara, 12 Aug 2013, C. Cripps CLC 2981 (MONT); loc. cit., with S. arctica and S. glauca, 8 Aug 2013, L. Gillman DBG-F-027685 (DBG). Similar taxa examined. USA. ALASKA. White Mountains, Eagle Summit, with Dryas, 5 Aug 2011, R. brevipes, C. Cripps CLC 2730 (MONT). Observations. Russula laevis was first described by Kälväinen, Ruotsalainen, and Taipale from a boreal area in Finland with Betula pubescens, B. czerepanovii, B. nana, and various Salix species on calcareous soils. Based on a phylogenetic analysis, including sequences from GenBank, R. laevis was determined to be a common inhabitant of Arctic and alpine regions in Finland, Norway, Sweden, Europe, Canada, and China (Adamčík et al. 2019). Russula laevis is in subgenus Brevipes along with other whitish, hard-fleshed, concave, and acrid species. These include R. chloroides (Krombh.) Bres., R. brevipes Peck, and R. delica Fr. Previously, reports of Russulas in subgenus Brevipes from Arctic and alpine areas with Salix and Dryas were referred to as R. delica, but this species was originally described from temperate habitats with other hosts. Morphological reports of R. “delica” from Arctic and alpine habitats include those from Europe (Schmid-Heckel 1985, 1988; Vila et al. 1997; Ballarà 1997; Corriol 2008), Fennoscandia (Gulden 2005; 136 Lange and Skifte 1967), Greenland (Lange 1957; Kobayasi et al 1971; Peterson 1977; Knudsen and Borgen 1982; Lamoure et al. 1982; Borgen 1993; Borgen et al. 2006), Iceland (Hallgrimsson 1998), the Rocky Mountains (Cripps and Horak 2008), Alaska (Kobayasi et al. 1967), the Canadian Arctic (Ohenoja and Ohenoja 2010) and Svalbard (Kobayasi et al. 1968; Skifte 1989). This work has confirmed that the report of R. delica from the Rocky Mountains (Cripps and Horak 2008) actually represents the species R. laevis and that this species is also present in Svalbard. It is possible that most other morphological reports of R. delica in Arctic and alpine regions actually represent R. laevis, but this needs to be confirmed with molecular methods. Previous work matched sequences extracted from ectomycorrhiza in Canada to R. laevis (Adamčík et al. 2019), but this is the first work to collect this species in North America. The Rocky Mountain specimens fall in a strongly supported clade with the holotype of R. laevis (only ITS region) along with collections from Norway, Svalbard, and Sweden. This is the first report of R. laevis from the Rocky Mountain alpine based on molecular sequence data, although it was previously reported as R. delica (Cripps and Horak 2008). Morphologically our specimens fit that of R. laevis with a few notable exceptions. The etymology of ‘laevis’ refers to the smooth and shiny pileus cuticle of the species (Adamčík et al. 2019); however, pilei of our collections are typically rough, with a matte appearance, a difference that could be due to xeric conditions in the southern Rockies. In addition, our spores are 1 µm smaller (8.7 × 7.2 µm) on average than those of R. laevis (10 × 8.5 µm). These differences suggest that further investigation is warranted. 137 Similar to R. laevis, Rocky Mountain specimens were near dwarf and shrubby Salix, Bistorta, and Dryas octopetala in calcareous soil. There has been considerable taxonomic confusion surrounding the subalpine species R. chloroides, R. brevipes, and R. delica located in R. subgenus Brevipes (Buyck and Adamčík 2013); however, issues regarding subalpine taxa are beyond the scope of this study. We followed Adamčík et al. (2019) and included a sequence representing the R. delica UNITE species hypothesis (UDB025023) and R. chloroides UNITE species hypothesis (AF418604) in our phylogenetic analysis. Our phylogenetic analysis confirms that these two species are separate from our Arctic and alpine clade represented by R. laevis. Russula montana Shaffer FIGS. 10, 16B, 18B, 20C Beihefte zur Nova Hedwidia 51: 234 (1975) ?= Russula grisescens (Bon & Gaugué) Marti, Docums Mycol. 14(no. 53):57 1984 Macromorphology. Pileus (25)35–70 mm wide, pulvinate when young, becoming convex to plano-convex, with a depression in center, primarily bright red, red, light to dark reddish brown with light brown, grayish red, light yellow, yellowish pink, yellow brown areas, occasionally brownish in center and dark pink near margin, sticky; margin with faint striations, 2–5 mm long, turned down; cuticle thin, smooth, viscid, shiny, separable except at center. Lamellae attached, adnexed to narrowly adnate, sometimes rounded in front, 3–6 mm broad, close, on av. L = 100–140 (n = 4), white, tinged grayish yellow, turning dingy yellow, intervenose, rarely forking near stipe, fragile; lamellae 138 edge entire. Stipe 20–50 × 10–20 mm, flared slightly at apex, otherwise equal or enlarging at base, white, not staining when bruised, dry, smooth, with a few longitudinal grooves, firm, moderately shiny. Context white, occasionally with hints of gray in age, soft or firm, 3–8 mm thick at pileus center, pinkish under cuticle; stipe stuffed. Odor indistinct or absent. Taste moderately to strongly acrid. Exsiccata: Pileus cherry red, dark red, red brown, dark maroon, purple red, purple brown, yellow buff, yellow orange; lamellae light brown or grayish buff; stipe yellow orange, yellow brown, orange brown, brown, occasionally with gray tones. Chemical reactions: Gum guaiac = slowly to quickly moderate green; ferrous sulfate = grayish yellow pink; 2% phenol = grayish red brown. Micromorphology. Basidiospores white with a yellow tinge, but not as dark as Romagnesi Ib (Shaffer 1975), (7.1–)7.4–8.0–8.6(–9.6) × (5–)5.7–6.3–6.8(–7.1) µm, broadly ellipsoid to ellipsoid, Q = (1.1–)1.2–1.3–1.4(–1.5); ornamentation of numerous, dense or sparse, amyloid, cylindrical or obtuse warts, <0.5 µm high, connected by lines and ridges forming a complete reticulum, with warts merging or rarely isolated; apiculus truncate, inamyloid; suprahilar plage small, weakly to moderately amyloid. Basidia (43.2–)46.8–53.2–59.5(–63.5) × (7.6–)8.4–9.7–11(–12.7) µm, subclavate to clavate, usually 4-spored, but 1- and 2-spored possible; basidiola clavate. Hymenial cystidia moderately abundant, (40.8–)51.7–68.8–86(–106.7) × (5.1–)8–9.6–11.3(–15.2) µm, clavate-pedicellate, fusiform, or almost cylindrical, occasionally subacute at apex, mucronate, with occasionally inflated 2.5–5.1 µm appendage; contents granular or crystalline, orange-brown in sulfovanillin. Pileipellis sharply delimited from underlying 139 context; suprapellis ca. 79–190 µm thick, composed of loosely interwoven hyphae, with occasionally swollen tips, and pileocystidia, well-defined from subpellis; subpellis weakly gelatinized, ca. 73–115 µm thick, composed of tightly interwoven, horizontally oriented hyphae, becoming more dense and darker in color near context. Hyphal terminations cylindrical, occasionally slightly clavate, thin-walled, ca. 1.5–4 µm wide at apex, apically obtuse, lightly granular. Pileocystidia near pileus margin numerous, clavate, occasionally cylindrical, rarely tapering, thin-walled, (45.9–)58.5–91.6–124.6(– 180) × (4–)4.8–5.9–6.9(–8) µm; contents granular or crystalline, dark gray to black in sulfovanillin. Pileocystidia near pileus center similar in shape and size, (55.9–)64.9– 96.5–128.2(–160) × (5.1–)5.2–6.3–7.4(–8.9) µm, occasionally forking; contents gelatinous or granular. 140 Figure 10. Russula montana (DBG-F-003596), photo Denver Botanical Garden. North American Ecology. On buried wood or moss in humus under Psudotsuga menziesii, Picea, Abies, and Pinus contorta. Aug. Material examined. FINLAND. Kainuu, Paltamo, Tololanmäki, with Betula, Alnus, Picea, and Populus, 30 Aug 2018, J. Vauras JV 32642F. USA. COLORADO. Clear Creek County, Arapaho National Forest, Squaw Pass, with spruce and fir, 4 Aug 1998, V. Evenson DBG-F-019556 (DBG); Gilpin County, Perigo, with Psudotsuga menziesii and Picea, 10 Aug 1972, R. L. Shaffer 12231 HOLOTYPE (MICH); loc. cit., 15 Aug 1974, A. H. Smith DBG-F-003596 (DBG); Park County, Jefferson Lake, with spruce-fir, 13 Aug 1972, R. L. Shaffer 9654 (MICH) (Shaffer examined this collection when he described the species); Summit County, Blue Lake Dam, with Betula glandulosa, 2 Aug 2001, C. Cripps CLC 1624 (MONT); Sawatch Range, Shrine Pass, in buried wood and moss, 15 Aug 1997, H. Tuttle DBG-F-019170 (DBG); White River National Forest, Copper Mountain, with Lodgepole Pine and spruce, 14 Aug 1997, B. Woo DBG-F-019236 (DBG). Observations. Shaffer (1975) originally described R. montana as a gregarious species on duff or rotten wood below Pseudotsuga menziesii, Abies, and Picea from Perigo Colorado, USA. After examining Shaffer’s holotype we are confident that our collections from the Rocky Mountains match that of R. montana, and all but one were found below treeline with conifers in the vicinity. The closely related R. nana is a smaller species that typically has a 12–35 mm wide pileus and is found in Arctic and alpine 141 habitats with Salix; R. montana typically has a larger pileus (35–70 mm wide) and occurs with conifers in the subalpine zone. There is also a significant difference in the spore length between R. montana and R. nana when analyzed using a two sample t-test (p-value = 0.002, based on 100 measurements from each species), but spore width is similar. Russula montana spores are on average 0.3 µm longer and have an average Q of 1.3, R. nana has an average Q of 1.2. R. nana forms a strongly-supported clade in a polytomy with sequences of R. montana in our phylogenetic analysis. This is not to be confused with some sequences in GenBank and UNITE that are incorrectly labeled R. nana (KX579809, UDB016029, and UDB015077) that are in a clade sister to R. betularum in our phylogenetic analysis. The R. montana/nana clade is sister to R. aquosa. A few distinct molecular differences between R. montana and R. nana are worth noting. There are a few conserved differences in the ITS sequences of R. montana that are not found in the closely related R. nana. There are two insertions present in the ITS1 region of R. montana sequences that are absent in R. nana; one is a G at position 184 in our alignment, the second is an ATTT from positons 265–268. There are also two single nucleotide polymorphisms (SNP’s) in the ITS2 region (TG at location 602–603) that are conserved throughout all our R. montana sequences that differ from that of R. nana (CT at these positions). Therefore, we maintain these as two separate species based on morphological and molecular differences as well as habitat. A species similar to R. montana is R. emetica var. grisescens described by Bon and Guagué in 1975 from France near Betula. Russula grisescens was raised to species 142 rank by Marti in 1984. Voitk (2015) recognized that R. griseascens (note the error in spelling) forms a species complex with R. aquosa and R. nana using molecular data, which is similar to our results. Bazzicalupo et al. (2016) investigated the distribution and occurrence of R. emetica in Newfoundland and Labrador, Canada, and found that R. montana represented the most common species in the region. Their work also indicated that R. grisescens and R. montana are close morphologically and molecularly. If R. montana is the same species as R. grisescens, the former has priority because it was described at the species level first in 1975, which was noted by Bazzicalupo et al. (2016). Numerous sequences in our phylogenetic analysis from Finland (JV 9788) and Austria (UDB031192) labeled R. grisescens cluster with R. montana in the phylogenetic analysis suggesting that these species are synonymous, but this needs to be confirmed by examining the type material of R. grisescens (Bazzicalupo et al. 2016). A proposed synonomy with R. hydrophila (Reumaux et al. 1996) has been questioned (Bazzicalupo et al. 2016) but should be considered when types can be examined. Based on morphology, R. montana has been reported in Colorado and Utah (Shaffer 1975). Molecularly this species has been confirmed in Newfoundland and Labrador, Canada by Bazzicalupo et al. (2016) and the phylogenetic analysis here include sequences from Colorado USA, Canada, Austria, and Finland, indicating that R. montana is intercontinentally disributed. Interestingly, R. montana has not been found in Montana, although subalpine Russulas have not been well-sampled. 143 Russula nana Killerman FIGS. 11, 16A, 18A, 20B Denkschr. Bayer. Botan. Ges. In Regnsb. 20: 38 1939 Macromorphology. Pileus 12–35(65) mm wide, hemispheric when young, broadly convex, to applanate, occasionally uplifted, and then slightly depressed in center, candy apple red, strawberry red, deep red, paling to white with ocher tints towards margin in age, smooth, slightly greasy, viscid when wet, occasionally matt, tacky when dry; margin slightly striate when young, more striate in age or not; cuticle separable except at center. Lamellae narrowly attached, adnate or adnexed, widest near pileus margin, close, on av. L = 75–120 (n = 5), white becoming light yellow brown in age or on drying, not anastomosing, occasionally forking near pileus margin, neither fragile nor tough; lamellae edge entire. Lamellulae present and extending 1/3 of the way from pileus margin to stipe. Stipe 6–26 × 5–13 mm, clavate, equal, central to slightly eccentric, white, with greyish or grey brown tints developing first at base, smooth, fragile or not, with faint vertical ridges. Context white, not staining; in stipe not hollow but with soft areas or small cavities near base. Odor indistinct. Taste moderately acrid. Exsiccata: Pileus red pink, cherry red, bleached ocher, tan, occasionally dark maroon near margin. Lamellae gray, ocher gray, yellow brown near pileus margin. Stipe white or grayish, with brown staining near base. Chemical Reactions: Gum Guaiac = blue green or turquoise; ferrous sulfate = very light yellow brown, 2% phenol = no reaction. Micromorphology. Basidiospores white in print (Ia), (6.1–)7.1–7.7–8.3(–9.6) × (5.1–)5.9–6.3–6.6(–7.6) µm, broadly ellipsoid, Q = (1.1–)1.2–1.2–1.3(–1.6); ornamentation of numerous, amyloid, warts and spines connected by thin lines and ridges 144 forming a full reticulum, <0.5–1 µm high, occasionally subreticulate with isolated cylindrical warts and spines; apiculus truncate or apically tapering, inamyloid; suprahilar plage small to large, amyloid. Basidia (38.1–)45.7–50.6–55.4(–63.5) × (7.62–)8.6–9.8– 11.1(–12.7) µm, clavate-pedicellate, occasionally cylindrical, 4-spored, possibly 2- spored; basidiola clavate. Hymenial cystidia (55.9–)64.9–74.9–85(–96.5) × (7.6–)9– 10.2–11.4(–15.2) µm, clavate-pedicellate, fusiform, rarely forking near apex, mucronate with swollen or cylindrical 2.5–5.1 µm long appendage or not; contents granular, lipid drops present, blackening in sulfovanillin. Pileipellis sharply delimited from underlying context; suprapellis ca. 100–150 µm thick, composed of loosely interwoven hyphae and pileocystidia with no direction; subpellis ca. 85–140 µm thick, composed of tightly interwoven, horizontally oriented hyphae, becoming more dense near context. Hyphal terminations cylindrical, thin walled; terminal cells 16.3–53 × 1.5–4.5 µm, apically obtuse, devoid of contents or rarely granular at apex. Pileocystida near pileus margin abundant, occasionally septate, clavate-pedicellate, becoming thinner as they increase in length, (30.5–)54.2–92.3–130.4(–203.2) × (3.8–)4.7–5.5–6.3(–7.6) µm, occasionally forking below septa; contents granular or crystalline, blackening in sulfovanillin. Pileocystidia near pileus center scattered to abundant, similar in shape but shorter, occasionally fusiform or swollen, tapering slightly at apex (subacute), (38.1–)48.6–71.6– 94.6(–137.2) × (3.8–)4.7–5.7–6.6(–7.6) µm, occasionally forking; contents similar. 145 Figure 11. Russula nana. A. with Salix reticulata, Beartooth Plateau, Montana (CLC 3619). B. with S. arctica, Cinnamon Pass, Colorado (CLC 1812). Scale bar = 2 cm. 146 North American Ecology. In alpine areas of Colorado, Montana, and Wyoming with Salix reticulata and S. planifolia in wet soil with moss. Jul and Aug. Material examined. ITALY. Belluno, with Salix reticulata and S. retusa, 7 Aug 2011, E. Campo EC 17.08.2011. USA. COLORADO. Boulder County, Niwot Ridge, with Salix species, 23 Aug 2018, C. Noffsinger CRN 177 (MONT); Pitkin County, Sawatch Range, Independence Pass, with dwarf Salix, 6 Aug 2003, V. Evenson DBG-F- 021359 (DBG); San Juan County, San Juan Range, Cinnamon Pass, with S. planifolia and S. reticulata, 1 Aug 2000, C. Cripps CLC 1440 (MONT); loc. cit., Salix arctica, 27 Jul 2002, C. Cripps CLC 1812 (MONT); Black Bear Pass, 3 Aug 2000, C. Cripps CLC 1450 (MONT). MONTANA. Carbon County, Hellroaring Plateau, 14 Aug 2008, C. Cripps CLC 2330 (MONT). WYOMING. Park County, Beartooth Plateau, Frozen Lake, with Salix reticulata, 23 Aug 2017, C. Cripps CLC 3619 (MONT); loc. cit., S. reticulata, 17 Aug 2017, C. Cripps CLC 3575B (MONT); Billings Fen site, with S. planifolia, 7 Aug 2018, C. Noffsinger CRN 135 (MONT). Similar taxa examined. GREENLAND. Sismiut, with Salix herbacea, 20 Aug 2000, C. Cripps CLC 1544 (MONT). Observations. Killerman described R. nana in 1936 based on a collection he found at 2700 meters in Tirol, Austria. The original description mentions a small specimen with a blood-red, flat pileus, white lamellae, and very hot taste; he did not specify the date or where the type was deposited. Kühner (1975) then reported R. nana as a common species in Arctic and alpine habitats of Europe near dwarf willow and occasionally Polygonum 147 viviparum (Bistorta vivpara). Bresinsky et al. (1980) examined the type material of Killerman’s and confirmed that R. nana represented a common Arctic-alpine species as Kühner (1975) had recognized. They also synonymized the name R. alpina (Blytt) Møller and Schäffer with R. nana Killerman. Sarnari (1998–2005) claimed that the type material examined by Bresinsky et al. (1980) was unusable because it lacks a collection date. Killerman’s original illustration consists of only a small, black and white drawing of two fruiting bodies and a single spore, which was also not sufficient. Therefore, Sarnari designated an epitype from the alpine region of Monta Rosa, Italy near Dryas (n˚ 97/800, in Herb. IB). Russula nana is one of the most iconic Arctic-alpine species of Russula and is considered common in regions of Europe were it has been reported in the Alps, Pyrenees, or Carpathians (Favre 1955; Horak 1960; Kühner 1975; Lamoure 1982; Bon 1985; Kühner and Lamoure 1986; Bon 1987; Bon and Cheype 1987; Senn-Irlet 1987; Schmid- Heckel 1988; Tondl 1988; Bon 1991; Fellner and Landa 1991; Fellner and Landa 1993b; Jamoni 1995; Adamčík 1998; Peintner 1998; Sarnari 1998–2005; Bon 2000; Bresinksy et al. 2000; Moreau 2002; Corriol 2008; Jamoni 2008; Ronikier 2008; Gyosheva and Dimitrova 2011), Faroe Islands (Vesterholt 1998), Fennoscandia (Gulden and Lange 1971; Kühner 1975; Gulden et al. 1985; Graf 1994; Sarnari 1998–2005; Ohenoja 2000; Gulden 2005), Netherlands (Geml et al. 2014), Poland (Ronikier and Adamčík 2009), Russia (Knudsen and Mukhin 1998; Niezdoiminogo 2003), Scotland (Watling 1987), Iceland (Hallgrimsson 1998), and Svalbard (Gulden et al. 1985; Skifte 1989; Gulden and Torkelsen 1996). In North America, R. nana has been reported from Alaska (Miller 148 1982a), Canada (Ohenoja and Ohenoja 2010), Greenland (Watling 1977; Knudsen and Borgen 1982; Lamoure et al. 1982; Borgen 1993; Borgen et al. 2006), and the Rocky Mountains (Moser and McKnight 1987; Cripps and Horak 2008). The species has also likely been reported under the names R. nana var. alpina in the European Alps (Bon and Ballarà 1996) and as R. alpina in Fennoscandia (Lange and Skifte 1967), Greenland (Lange 1957; Kobayasi et al. 1971; Peterson 1977; Watling 1977; Lamoure et al. 1982; Watling 1983), Iceland (Christiansen 1941), and Svalbard (Kobayasi et al. 1968). None of these reports have been molecularly confirmed. This is the first research to perform and in-depth systematic analysis of R. nana that includes molecular data. Our phylogenetic analysis includes Rocky Mountain alpine collections from Colorado, Wyoming, and Montana as well as reference sequences from Italy (near epitype locality), Scotland, and Switzerland. Russula nana forms a strongly- supported clade that forms a polytomy with several collections of R. montana (FIG. 4), including the R. montana holotype. Russula nana and R. montana are morphologically similar, but differ in a few important characteristics. Russula nana is a small species (pileus usually 12–35 mm wide) found in Arctic and alpine habitats with Salix. Whereas, R. montana has a larger pileus (usually 35–70 mm wide) and occurs with conifers in the subalpine zone. This ecological distinction is true with the exception of one collection originally identified as R. nana (F-127873); this collection was found near Salix herbacea in an alpine region of Norway and was resolved with R. montana. Sarnari (1998–2005) also noted a few small morphological differences between R. nana and the closely related R. grisescens, the latter being a possible synonym of R. montana (see observations under 149 R. montana). Russula montana spores are significantly longer than those of R. nana when analyzed using a two-sample t-test (p-value = 0.002, based on 100 measurements from each species). There are also few conserved differences in the ITS sequences of R. montana that are not shared with the closely related R. nana (see observations under R. montana). Based on this evidence we maintain R. nana and R. montana as two separate species. This is the first molecularly confirmed report of R. nana in the Rocky Mountain alpine zone; however, Geml et al. (2010) reported it in Alaska base on 97% sequence similarity, which may not be sufficient for Russula species delineation. This is the first work to molecularly confirm that this common Arctic and alpine species is indeed intercontinentally distributed, confirming earlier morphological studies in Alaska and the Rocky Mountain alpine (Miller et al. 1973; Miller 1982a; Moser and McKnight 1987; Cripps and Horak 2008). Several species and varieties have been synonymized with R. nana; however, none of these names have been verified using molecular methods and there are inconsistencies between sources. Kühner (1975) synonymized only R. emetica var. alpina Blytt & Rostr. with R. nana. Whereas, Sarnari (1998–2005) synonymized eight names with R. nana. He stated that two names, Russula emetica var. alpina A. Blytt & Rostr. and R. fragilis var. alpestris Bound., were generally believed to be synonymous with R. nana Killerman because R. nana was first described at the species level. He also synonymized R. alpina (A. Blytt & Rostr.) F.H. Møller & Jul. Schäff., R. alpestris (Boudier) Singer, R. emetica ssp. alpestris (Boudier) Singer, R. emetica var. alpestris Singer, R. nana var. alpina (Blytt & Rostrup) Bon and cited some concerns about the 150 synonymy of R. nigrodisca Peck (Singer 1943; Shaffer 1975) with R. nana. However, many of these names have now been synonymized under R. emetica (Shaeff.) Pers. by others. Therefore, at this time we have not included these proposed synonyms of R. nana until further analysis of type specimens and the corresponding molecular work can be completed. There is some confusion in the literature regarding the correct phylogenetic placement of R. nana. Several studies have resolved R. nana as a closely related species to R. betularum (Miller and Buyck 2002; Li et al. 2015; Bazzicalupo et al. 2016). This work that used morphological and molecular data from European and North America collections determined that R. nana is closely related to R. montana. Multiple sequences in Genbank and UNITE labeled R. nana form a sister clade to R. betularum in our analysis (UDB015077, UDB016029, and KX579809) that are distantly related to our R. nana clade. We also have collections of Russula from Alaska that fall into this imposter clade (CLC 3821 and CLC 3822B), which we are calling R. cf. alpigenes based on the description produced by Bon (1993). Based on our collections this species does not represent the common Arctic and alpine fungus described by Killerman. Russula cf. alpigenes and closely related taxa can be distinguished from R. nana in the field, by the pileus color that is carmine red with a dark center (not cherry red fading to white as in R. nana) and by the longer stipe that can have pink tints, (absent in R. nana). Of particular note is that the stipe dried bright pink in most of our collections tentatively identified as R. cf. alpigenes, this character has not been reported in R. nana. Also, the odor can be slightly sweet, and spores are smaller in R. nana (av 7.7 × 6.3 µm) compared to R. cf. 151 alpigenes (8.9 × 7 µm). There have been sequences generated prior to this study of the true R. nana; however, they are not publicly available. Caution is advised when using R. nana sequences available in public databases. Russula cf. pascua (F.H. Møller & Jul. Schäff.) Kühner FIGS. 12, 17C, 18G, 19A Bulletin de la Société Mycologique de France 91(3):331 1975 Macromorphology. Pileus 25–35 mm wide, convex, irregular convex, becoming applanate, with or without slight depression in center, deep magenta, red, dark red, candy apple red, ocher, rarely with small hints of green, lighter or darker at margin, smooth, dry, coarsely cracking or not, debris adhering to cap; margin striate for short distance or not, undulating in some; pileus cuticle separable except at center. Lamellae attached, adnate to adnexed, relatively close, on av 100–104 L (n = 2), light straw yellow, yellow brown, browning near pileus margin in age, rarely anastomosing or forking, elastic not brittle. Lamellula absent or rare. Stipe 15–20 × 8–20 mm, clavate or strongly clavate, central, white or off-white, with brownish yellow stains near base or covering stipe, slightly pink on drying, smooth with indistinct vertical ridges. Context in stipe hard or soft, not hollow but with small cavities near base. Odor fishy. Taste radish hot, weakly acrid, or mild. Exsiccata: Pileus dark maroon, red, or ocher. Lamellae ocher, browning at pileus margin. Stipe white to ocher, usually darker near base. Chemical reactions: Gum Guaiac = brownish green; Ferrous sulfate = grayish green; 2% phenol = light brown to dark brown with a green hue. 152 Micromorphology. Basidiospores white to light cream in mass (Ib-IIa), (7.6–)8.4– 9.1–9.8(–10.6) × (5.5–)6.5–7–7.5(–8.2) µm, broadly ellipsoid to ellipsoid, Q = (1.1–)1.2– 1.3–1.4(–1.5); ornamentation of isolated, amyloid warts and spines, 0.5–1 µm high that are typically round or elongated at base and occasionally connected by thin lines, clustered to subreticulate; apiculus truncate, inamyloid; suprahilar plage large, amyloid. Basidia (45.7–)50.2–56.2–62.3(–66.3) × (7.6–)9.3–10.4–11.5(–12.2) µm, clavate or fusiform, 4-spored; basidiola clavate. Hymenial cystidia (53.9–)58.6–66.8–75.1(–81.3) × (6–)7.5–8.7–10(–10.2) µm, fusiform, pedicellate, often mucronate, with swollen ca. 3.8– 12.7 µm long appendage; contents granular or refractive, possibly banded, slightly browning in sulfovanillin. Pileipellis sharply delimited from underlying context; not stratified, ca. 114–331 µm thick, composed of highly variable, parallel, horizontally or vertically oriented, loosely or tightly interwoven, septate hyphae. Hyphal terminations near the pileus margin often attenuated, occasionally cylindrical, rarely inflated, thin- walled, (19.4–)24.7–30.7–36.7(–41.8) × (2.5–)3–3.5–4.1(–4.5) µm; contents lightly granular. Hyphal terminations near the pileus center cylindrical or attenuated, occasionally inflated, thin-walled, (24.8–)26.7–31.5–36.4(–40.8) × (3.1–)3.2–3.7–4.2(– 5.1) µm; contents lightly granular. Pileocystidia near the pileus margin rare, clavate, septate, (25.5–)32.9–58.2–83.5(–106.7) × (4.1–)4.5–5.8–7.1(–8.9) µm; contents lightly granular, no reaction in sulfovanillin. Pileocystidia near pileus center similar in shape and size, tapering at apex (subacute), or fusiform, (27.9–)37.2–53.6–70(–88.9) × 4.7–5.8– 6.9(–7.6) µm; contents lightly granular or refractive gelatinous. 153 Figure 12. Russula cf. pascua (CRN 138) growing with Salix reticulata, Beartooth Plateau, Montana. Scale bar = 2 cm. North American Ecology. Occurring in the alpine with Salix reticulata in Montana, Wyoming, and Colorado. Aug. Material examined. USA. COLORADO. Boulder County, Niwot Ridge, 1 Aug 2006, C. Cripps CLC 2274 (MONT). WYOMING. Park County, Beartooth Plateau, Frozen Lakes, with Salix reticulata, 8 Aug 2018, C. Noffsinger CRN 138 (MONT); loc. cit., with S. reticulata, 8 Aug 2018, C. Noffsinger CRN 146 (MONT). Similar taxa examined. USA. MONTANA. Boulder County, Niwot Ridge, 8 Aug 1998, C. Cripps CLC 1220 (MONT). 154 Observations. One species from the Rocky Mountain alpine is phylogenetically located in R. subsection Xerampelinae, whose members are known for having red-brown colors in the pileus, a strong fishy odor, and a green reaction in the context when exposed to ferrous sulfate (Adamčík and Knudsen 2004); the well-known R. xerampelina is in this subsection. When encountering this species in the field, a few collections were initially identified as R. nana because of the bright red color, until a fishy odor was noticed. In the phylogenetic analysis, collections from the Rocky Mountain alpine zone form a polytomy with members of the closely related R. clavipes complex (Adamčík et al. 2016b). The R. clavipes complex contains R. pascua (F.H. Møller & Jul. Schäff.) Kühner, R. clavipes Velen., and R. nuoljae Kühner. Russula nuoljae is morphologically and molecularly distinct; however, R. pascua and R. clavipes are belived to be similar morphologically and do not separate with strong support in phylogenetic analyses (Adamčík et al. 2016b). However, others think of R. clavipes as a bigger subalpine species with Betula (Læssoe and Peterson 2019). Adamčík et al. (2016b) separated the three species in the R. clavipes based on morphological characters, mainly those present in the pileipellis and the habitat. When considering the important characters for species delineation in Adamčík et al.’s (2016b) key, our species has pileocystidia near the pileus margin that are barely less than 6 µm wide on average, attenuated terminal cells that are barely longer than 30 µm wide on average, and is reported with dwarf and shrubby Salix from Arctic-alpine habitats. When using these characters to delineate the European taxa studied in Adamčík et al. (2016b) our taxon from the Rocky Mountain alpine does not clearly fit the description of R. pascua. However, our species matches many other morphological characters of R. 155 pascua (Adamčík and Knudsen 2004) and is found in the same habitat. Another similar species based on morphology and ecology is R. amoenipes Romagn. f. kuehneriana ad int. (Bon 2000). Russula amoenipes f. kuehneriana is characterized by a pileus around 30–50 mm in diameter with a matte red color, yellow basidiospores 9 × 7 µm with warts 0.6–1 µm high, and pileocystidia 4–7 µm wide that do not blacken in sulfovanillin. All of the above characteristics closely match that of our species. Of the three species reported by Adamčík et al. (2016b), Russula pascua is the one most commonly reported in Arctic and alpine areas, and these include Europe (Adamčík and Knudsen 2004; Adamčík et al. 2016b; Fellner and Landa 1993b; Graf 1994; Ronikier and Adamčík 2009; Schmid-Heckel 1988), Fennoscandia (Gulden 2005), Faroe Islands (Adamčík and Knudsen 2004), Greenland (Adamčík and Knudsen 2004; Borgen et al. 2006), Poland (Adamčík and Knudsen 2004; Adamčík et al. 2016b; 59), Russia, and Scotland (Watling 1987). The reports in Europe and Poland have been molecularly confirmed (Adamčík et al. 2016b). Due to the similarity of our species to R. pascua, the large distribution reported for R. pascua in Arctic and alpine habitats, and the lack of reference material available for Russula amoenipes f. kuehneriana we elect to call our species R. cf. pascua at this time. However, our species shared similarities with Russula amoenipes f. kuehneriana and this will continue to be investigated. Russula pascua was originally described as R. xerampelina var. pascua F.H. Møller & Jul. Schäff. and its species status was confirmed by Kühner (1975). Sarnari (1998–2005) incorrectly designated a neotype for R. pascua that also fell into the 156 subrubens clade (Adamčík et al. 2016b). We used sequences from Adamčík et al. (2016b) to build our phylogeny to investigate Rocky Mountain alpine members of the R. clavipes complex. Based on their work, R. pascua has been molecularly confirmed in Italy, Poland, Slovakia, Austria, and Romania. A BLAST search in the UNITE database revealed numerous highly similar sequences. However, species in the R. clavipes complex are so molecularly similar that even BLAST matches that share 99.6% sequence similarity do not fall into our Rocky Mountain clade. If Adamčík et al.’s (2016) circumscription of the R. clavipes complex reflects reality, speciation in this group may occur with as little as 0.3% variation in the ITS region. Russula purpureofusca Kühner FIGS. 13, 18H, 19D Bull. Trimmest. Soc. Mycol. Fr. 91(3):389. 1975 = Russula cupreola Sarnari, Boll. Assoc. Micol. Ecol. Romana 7 (20–21):64. 1990 Macromorphology. Pileus 20–45 mm wide, convex when young, becoming applanate with a slight depression, fully maturing to a funnel-shape, deep red, reddish brown, deep magenta, light yellowish green, yellow brown, darker over disk in age, smooth, viscid, with debris adhering; margin not striate or striate in age, occasionally undulating; cuticle separable except at center. Lamellae attached, adnate, close, on av. 70–104 L (n = 3), light yellow, cream, dark yellow cream, brownish yellow, darker near margin, anastomosis rare, no forking, entire, elastic not brittle. Lamellulae rare. Stipe 10– 35 x 5–15 mm, clavate or equal, central, white, off-white, light yellow, graying, with 157 yellow-brown stains near base, yellow-brown when bruised, smooth. Context hard or soft, white or watery gray, with large cavity near base in some collections. Odor absent, fungoid, or sweet. Taste strongly acrid (after 20–30 seconds), mild in one collection. Exsiccata: Pileus dark violet brown, darker brown to black over disk, sometimes olive green near margin. Lamellae brownish orange, ocher, dark yellow, dull. Stipe white, yellow brown, with hints of gray. Chemical reactions: Gum Guaiac = dark olive brown; ferrous sulfate = faintly pink or pinkish gray brown; 2% phenol = no reaction. Micromorphology. Basidiospores dark yellow in print (IIIb – IIIc), (7.5–)8.0–8.5– 9.0(–9.6) × (6.1–)6.6–7.0–7.4(–8.2) µm, subglobose to broadly ellipsoid, Q = (1.1–)1.1– 1.2–1.3(–1.4); ornamentation of numerous, isolated, amyloid, warts, 0.5–1 µm high and ca. 1 µm in diameter at base, with some warts fusing or occasionally connected by fine lines or ridges, not forming a reticulum; apiculus truncate or pointed, inamyloid; suprahilar plage medium to large, amyloid. Basidia (43.2–)48.9–54.2–59.4(–63.5) × (10.2–)10.3–11.1–11.8(–12.7) µm, clavate-pedicellate, 4-spored; basidiola clavate. Hymenial cystidia (71.1–)74.2–85.5–96.8(–119.4) × (7.6–)8.9–11.2–13.5(–15.2) µm, fusiform or clavate, often mucronate, with 2.5–7.6 µm long appendage; contents granular, gelatinous, or crystalline, graying to slightly blackening in sulfovanillin. Pileipellis sharply delimited from underlying context; suprapellis ca. 114–152 µm thick, composed of erect, forking, septate, hyphae, well-defined or gradually transitioning to subpellis; subpellis gelatinized, ca. 127–203 µm thick, composed of difficult to differentiate horizontal hyphae. Hyphal terminations cylindrical or clavate, flexuous, thin-walled, ca. 5–6 µm wide at apex, apically obtuse or occasionally subacute, 158 occasionally with granular contents. Pileocystidia near pileus margin 1–2 septate, clavate or cylindrical, thin-walled, (53.3–)70.6–102.9–135.2(–177.8) × (5–)5–6–7(–7.6) µm, apically obtuse or subacute, occasionally forking, with small lateral protrusions present (lateral diverticulate); contents granular or gelatinous, occasionally banded, dark black in sulfovanillin. Pileocystidia near pileus center often shorter, strongly clavate, and not forking, (50.8–)60.1–83.1–106(–127) × (3.8–)5–6.4–7.8(–8.9) µm; contents similar. 159 Figure 13. Russula purpureofusca. A. with Salix and Dryas, near Girdwood, Alaska (CLC 3820), photo Noah Siegel. B. with S. planifolia, Loveland Pass, Colorado (CRN 165). Scale bar = 2 cm. North American Ecology. In the alpine zone with Salix planifolia and other Salix species in Alaska and Colorado. Aug. 160 Material examined. NORWAY. Nord-Trøndelag, Levanger, 21 Aug 1998, G. Gulden F-60573 (O). SWEDEN. with Salix and Dryas, 22 Aug 2019, J. Vauras JV 32876F. USA. ALASKA. Anchorage County, Girdwood, with Salix and Dryas species, 22 Aug 2018, C. Cripps CLC 3820 (MONT). COLORADO. Summit County, Loveland Pass, with Salix planifolia, 19 Aug 2018, C. Noffsinger CRN 164 (MONT); loc. cit., with S. planifolia, 19 Aug 2018, C. Noffsinger CRN 165 (MONT). Observations. Russula purpureofusca was originally described by Kühner in 1973 as a small brownish-red alpine species with a light-yellow spore print (IIc), an acrid taste, and 8–8.7 × 6.7–7.5 µm basidiospores, occurring near Salix reticulata (Ruotsalainen and Huhtinen 2015). In 1990, Sarnari described a similar species from the alpine zone of Europe that he called R. cupreola. This species was also collected with S. reticulata but differed from Kühner’s species by the darker yellow spore print (IVc). After examining the holotypes of both R. purpureofusca and R. cupreola, Ruotsalainen and Huhtinen (2015) synonymized the two species and gave the older name, R. purpureofusca, priority. They determined that differences in spore color were due to the maturity of the specimens collected by Kühner compared to the specimens examined by Sarnari. They also found that the holotypes of R. purpureofusca and R. cupreola had identical spore morphology and pileocystidia with abundant lateral diverticulae, which appears to be a distinguishing feature. A BLAST search for sequences similar to CLC 3820 in the UNITE database reveled 98.2% sequence similarity of the ITS region with R. cupreola type (UDB032344), but this sequence is locked and unavailable. 161 This is the first report of R. purpureofusca from North America. Our phylogenetic analysis produced a strongly-supported clade containing North American collections and reference material from Norway and Sweden, including a collection examined by Ruotsalainen and Huhtinen (2015) labeled R. purpurofusca (H6042234). Our North American collections from Colorado have a yellow-green pileus; whereas, the pileus in our Alaskan collection is reddish-brown to magenta, colors normally associated with this species. However, Sarnari (1998–2005) mentioned an olive-ocher coloration in his description of R. cupreola. Significant intraspecific color variation has been observed in the pileus of other species of Russula (Bazzicalupo et al. 2017). All our collections matched R. purpureofusca in spore dimensions and all had lateral protrusions (lateral diverticulae) on the pileocystidia. There are only three base pairs difference between the Alaska collection and the two collections from Colorado. Morphologically and to some degree molecularly, Ruotsalainen and Huhtinen (2015) confirmed R. purpureofusca to occur in Finland, Norway, and Switzerland, although their molecular data were not published. Morphologically, Sarnari (1998–2005) reported R. cupreola from Finland and Fellner and Landa (1993) reported it in alpine areas of Slovakia. Therefore, it is likely that R. purpureofusca has an intercontinental distribution in Arctic and alpine habitats of North America, Fennoscandia, and the European Alps. Russula purpureofusca can be confused with R. laccata and R. saliceticola, which also have a deep magenta to ruby pileus and occur in alpine and Arctic areas with Salix. However, R. laccata has white lamellae and white spores that are smaller (8.1 × 6.2 µm) 162 and reticulate. Russula saliceticola is mild tasting, often with pink tones on the stipe, and the spore print is usually a lighter yellow color. Of the three species, only R. purpureofusca has lateral protrusions on the pileocystidia and all can be separated phylogenetically using the ITS region (FIG. 3). Russula saliceticola (Singer) Kühner ex Knudsen & T. Borgen FIGS. 14, 17A, 18E, 19C Arctic and Alpine Mycology, 1980 (Seattle):224 1982 = Russula saliceticola var. gaviae Bincol., Giac. & Ostellari, Riv. Micol. 54(1):30 2011 = Russula sphagnophila subsp. saliceticola Singer, Annls mycol. 34(6):425 1936 Macromorphology. Pileus 10–55 mm in wide, convex, conic, applanate, occasionally depressed in center, deep red, grayish ruby, almost black in center, red to dark red closer to margin, occasionally with ocher or tan areas, occasionally lighter in center or uniform in color, glabrous, greasy or dry, matte or shiny, with debris adhering; margin smooth to faintly sulcate-striate, more striate in age, inrolled or not, entire or undulate; cap cuticle separable except at center. Lamellae attached, adnate to adnexed, subdistant or close, on av. L = 64–84 (n = 5), white, light yellow, cream, warm yellow, occasionally white near stipe and more straw yellow near pileus margin, occasionally magenta near cap margin, anastomosing or not, forking near margin or not, elastic not brittle; browning near margin in age, entire or undulate. Lamellulae present or absent. Stipe 15–60 × 5–25 mm, clavate or equal, central, pure white, off-white, buff yellow, cream, occasionally becoming tan brown in age, smooth, pruinose near base (maybe due to frost), with or without purple or pinkish tint, that becomes more pronounced after 163 collecting, with greyish brown and brownish red stains near base, with indistinct vertical ridges. Context watery white, flesh tinted red under cuticle; in stipe spongy but firm, neither hard nor fragile, not hollow but occasionally with a few cavities. Odor usually absent, rarely sweet. Taste mild, occasionally weakly acrid. Exsiccata: Pileus grayish ruby to dark ruby, almost black near center, with red, brown, ocher, or buff. Lamellae light buff or brown, ocher gray yellow, grayish orange, egg yolk yellow, staining darker brown in areas. Stipe off-white, yellow brown, light tan brown, brown black, with pinkish purple stains that became more pronounced on drying, gray stains also present. Chemical reactions: Gum Guaiac = bluish green, grayish brown, green brown, brown; ferrous sulfate = ranging from yellow brown, dark pink, to red brown, rarely with green hue; 2% phenol = no reaction or occasionally light brown or brown. Micromorphology. Basidiospores white, light yellow cream, or warm yellow cream in print (Ia-IId), (7.1–)8.1–8.9–9.8(–11.6) × (5.1–)6.3–7–7.6(–9.6) µm, broadly ellipsoid to ellipsoid, Q = (1.1–)1.2–1.3–1.4(–1.5); ornamentation of dense, amyloid, isolated warts, 0.5–1 µm high, with cylindrical, oval, or angular bases, that fuse, and are occasionally connected by thin lines and/or ridges forming a subreticulum; apiculus truncate or pointed, inamyloid; suprahilar plage small to large, amyloid, occasionally difficult to observe. Basidia (38.1–)43.9–51–58.1(–66) × (7.6–)9.5–11–12.5(–14.3) µm, clavate-pedicellate, subacute at apex, 4-spored, occasionally 2-spored; basidiola clavate. Hymenial cystidia rare, (55.9–)62.9–78.8–94.7(–147.3) × (6–)7.8–9.6–11.4(–12.7) µm, fusiform, cylindrical, occasionally pedicellate, often mucronate, with 1–4 µm long appendage; contents granular, crystalline, amorphous, or absent, occasionally with lipid 164 drops, slightly graying, with orangish-brown reaction in sulfovanillin. Pileipellis sharply delimited from underlying context; suprapellis weakly gelatinized, ca. 63–100 µm deep, composed of tightly or loosely interwoven, erect, septate hyphae, hard to differentiate, gradually passing to subpellis; subpellis, strongly gelatinized ca. 63–102 µm deep, composed of hard to differentiate, densely interwoven, septate, horizontally oriented hyphae. Hyphal terminations slightly clavate, thin-walled, ca. 2–4 µm wide at apex, apically obtuse and swollen; rarely with lightly granular contents, difficult to observe. Pileocystidia near the pileus margin usually with one septa, cylindrical, pedicellate, hyphal like, (58.4–)81–110.4–139.7(–221) × (3.8–)4.2–5.1–6(–7.6) µm; contents granular or crystalline, occasionally with small lipid drops, blackening in sulfovanillin. Pileocystidia near the pileus center similar in shape but shorter and with 1–2 septa, occasionally deformed; heavily filled with granular contents (48.3–)51.6–77.2–102.9(– 152.4) × (3.8–)4.7–5.9–7.1(–7.6) µm. 165 Figure 14. Russula saliceticola. A. with Salix planifolia, Niwot Ridge, Colorado (CRN 173). B. with S. planifolia, Beartooth Plateau, Montana (CRN 155). Scale bar = 2 cm. 166 North American Ecology. In alpine areas of Montana, Wyoming, and Colorado with Salix reticulata and Salix planifolia. Jul and Aug. Material examined. SWEDEN. Lycksele lappland, Arjeplog, with Salix and Betula, 13 Aug 2018, J. Vauras JV 32515F. USA. COLORADO. Boulder County, Blue Lake, with Salix planifolia, 24 Aug 2018, C. Noffsinger CRN 184 (MONT); Niwot Ridge, with S. planifolia, 20 Aug 2018, C. Noffsinger CRN 173 (MONT). MONTANA. Carbon County, Beartooth Plateau, Birch site, with S. planifolia, 20 Aug 2017, C. Cripps CLC 3575C (MONT); borrow pit, with dwarf Salix, 29 Jul 1997, C. Cripps CLC 1137 (MONT); Highline Trailhead, with S. planifolia, 10 Aug 2018, C. Noffsinger CRN 155 (MONT). WYOMING. Park County, Beartooth Plateau, Billings Fen site, with S. reticulata, 23 Aug 2017, C. Cripps CLC 3616 (MONT); loc. cit., with Salix species, 7 Aug 2018, C. Noffsinger CRN 134 (MONT); Frozen Lake, S. planifolia, 8 Aug 2018, C. Noffsinger CRN 140 (MONT); loc. cit., with Salix species, 8 Aug 2018, C. Noffsinger CRN 143 (MONT); loc. cit., with S. planifolia, 8 Aug 2018, C. Noffsinger CRN 144 (MONT). Wyoming creek, 6 Aug 2008, C. Cripps CLC 2370 (MONT). Observations. Singer originally described R. nitida var. saliceticola Sing. in 1936 from the Altai Mountains near Salix herbacea (Romagnesi 1967; Schmid-Heckel 1985) and Kühner (1975) raised the variety to species rank. However, Kühner did not validly publish the species and Knudsen and Borgen (1982) re-described R. saliceticola in 1982. This species is recognized by a deep-red to ruby pileus that is almost black in the center, light yellow to cream lamellae, whitish stipe often with a pink tint, yellow spores with primarily isolated warts, and a mild taste. In the Rocky Mountain alpine, R. saliceticola 167 can be confused with R. laccata, which has more magenta in the pileus, a hotter taste, white reticulate spores, and a completely white stipe; and with R. purpureofusca, which has deeper magenta and yellow tones in the pileus, and darker yellow spores. Molecularly, R. saliceticola, R. laccata, and R. purpureofusca can all be separated using the ITS region. In the Rocky Mountains, this species is commonly found with Salix planifolia and S. reticulata in alpine habitats during July and August. Our description matches the morphology of R. saliceticola as described by Kühner (1975) and our Rocky Mountain alpine collections form a moderately-supported clade that includes reference sequences from Norway, Sweden, and Switzerland. The R. saliceticola clade forms a polytomy with the the species R. nitida and R. sphagnophila. Russula nitida, R. sphagnophila, and R. saliceticola are all morphologically similar with magenta in the pileus, cream-yellowish lamellae, whitish stipe, often with a pink tint, weak odor, and mild taste, although R. saliceticola has a typically darker, less striate pileus. Russula nitida and R. sphagnophila are both associated with Betula (Knudsen and Borgen 1982), and R. saliceticola occurs with Salix, primarily in Arctic-alpine habitats. Russula sphagnophila is often in sphagnum moss and appears to be comparatively more fragile, with more of a slight lavender tint at the apex of the stipe and more reticulated spores. Russula nitida has spores with primarily isolated warts. Morphologically, R. saliceticola has been reported in Arctic and alpine areas of the European alps (Favre 1955; Kühner 1975; Schmid-Heckel 1985, 1988; Bon 1991), Fennoscandia (Kühner 1975; Hansen and Knudsen 1992; Gulden 2005), Western 168 Carpathians in Slovakia (Fellner and Landa 1993; Knudsen and Ronikier 2003), Romania (Ronikier 2008), Faroe Islands (Vesterhot 1998), Greenland (Knudsen and Borgen 1982; Lamoure et al. 1982; Borgen et al. 2006), Iceland (Hallgrimsson 1998), and Svalbard (Skifte 1989). Knudsen and Borgen (1982) also claim that this species was described from the alps as R. brunneoviolacea Crawshay (Singer 1936; Favre 1955). This is the first report of R. saliceticola from the Rocky Mountains. Russula subrubens (J.E. Lange) Bon FIGS. 15, 17B, 18F, 19B Docums. Mycol. 2(5):33 1972 = Russula chamiteae Kühner, Bull. trimest. Soc. mycol. Fr. 91(3):389 1975 = Russula chamiteae var. microsperma Kühner, Bull. trimest. Soc. mycol. Fr. 91(3):389 1975 = Russula graveolens var. subrubens J.E. Lange, Fl. Agaric. Danic. 5 (Taxon. Consp.):Vlll 1940 Macromorphology. Pileus 10–80 mm wide, convex (shallow or broad), becoming applanate, or slightly dished, rarely umbonate, primarily mottled with some combination of reddish, reddish brown, reddish orange, orange brown, yellowish brown, ocher, or light brown, also with brownish violet, pale maroon, olive , pale orange, pinkish ocher, and cream tones, lighter in age, smooth or roughened, matt, dry, greasy, or sticky, cracking in age; margin not striate, indistinctly striate, or slightly tuberculate-striate, turned under at first, entire, undulating or not; cap cuticle separable except at center. Lamellae narrowly attached, adnexed, adnate, slightly sinuate or dished, crowded, becoming broad, on av. L = 90–200 (n = 8), white, cream, pale ocher, dark cream, 169 yellowish, pale golden yellow, straw yellow, staining brownish near pileus margin, rarely anastomosing, forking or not, fragile or elastic; edges concolorous, occasionally dark brown. Lamellulae rare. Stipe 10–60 × 10–30 mm, equal to strongly clavate, central to slightly eccentric, white, cream, yellow, ocher, dark cream, orange, yellow, yellow brown, or pinkish ocher, occasionally lighter near apex and darker near base, grooved, rarely with bumps, hoary or smooth, greasy, matt; not staining when bruised. Context white, cream, dark cream, buff, pinkish ocher under cuticle; in stipe hard, spongy, or fragile, slightly watery grey in base, staining slightly brown, not hollow but a few cavities near base. Odor slightly to strongly fishy, especially in older specimens or when dried, but can be absent. Taste mild, bitter, slightly astringent (acidic or bitter), or absent. Exsiccata: Pileus mottled with light and dark brown, grayish magenta, red brown, ocher, shiny. Lamellae light straw yellow to yellowish brown, yellow orange, yellow gray, gray brown, more brownish near pileus margin. Stipe whitish, off-white, yellow brown, or gray brown, with yellow brown and brown stains. Chemical reactions: Gum guaiac = blue green, brown green, brown, or brown gray; ferrous sulfate = strongly to moderately gray green; 2% phenol = slowly pinkish or brownish. Micromorphology. Basidiospores yellow to yellow cream in print (IId-IIIa), (7.1– )7.8–8.5–9.2(–12.2) × (5.6–)6.2–6.7–7.3(–9.6) µm, broadly ellipsoid to ellipsoid, Q = (1.1–)1.2–1.3–1.3(–1.6); ornamentation of dense, isolated, amyloid, warts and spines, 0.5–1 µm high, with circular, angular bases ca. 1–1.5 µm in diameter, with some warts and spines fusing but individually recognizable, occasionally connected by fine lines and ridges, occasionally forming a subreticulum, rarely forming a complete reticulum; 170 apiculus truncate, rounded, or pointed, inamyloid; suprahilar plage medium to large, amyloid. Basidia (43.2–)49.9–59.9–70(–101.6) × (7.6–)9.1–10–10.9(–12) µm, clavate, subacute, occasionally asymmetric, 4-spored, some possibly 2-spored; basidiola clavate. Hymenial cystidia scattered, (61–)70.9–82.4–93.9(–114.3) × (7.4–)8.5–9.9–11.3(–12.7) µm, turbinate (spindle shaped), fusiform, rarely cylindrical, pedicellate, often mucronate with occasionally swollen or irregular 1–7 µm long appendage, thin-walled; contents granular, amorphous, crystalline, or absent, orange-brown in sulfovanillin. Pileipellis sharply delimited from underlying context; suprapellis weakly gelatinized, ca. 68–78 µm thick, composed of loosely to tightly interwoven, erect, septate hyphae, gradually transitioning to subpellis; subpellis, strongly gelatinized, ca. 50–76 µm thick, composed of densely interwoven, horizontally oriented hyphae with slightly swollen tips. Hyphal terminations cylindrical, clavate, or swollen, ca. 2.5–4.5 µm wide at apex, apically obtuse, occasionally forking, difficult to differentiate; contents lightly granular or absent. Pileocystidia near the pileus margin abundant 1–2 septate, clavate, cylindrical, fusiform, occasionally irregular, (81.3–)97.2–117.9–138.6(–165.1) × (2.3–)4.1–5.2–6.4(–9) µm, occasionally forking, appearing to arise from deep within the suprapellis; contents granular, gelatinous, or crystalline, no reaction or slightly graying in sulfovanillin. Pileocystidia near the pileus center sparse, shorter, usually lightly granular or devoid of contents, (53.3–)58.6–73.8–88.9(–101.6) × (3.8–)4.4–5.4–6.5(–7.6) µm. 171 Figure 15. Russula subrubens. A. with dwarf Salix, Loveland Pass, Colorado (DBG-F - 020848), photo Denver Botanical Garden. B. with S. planifolia and S. glauca, Beartooth Plateau, Wyoming (CLC 3597). C. with S. reticulata and S. planifolia, Beartooth Plateau, Montana (CLC 3588). D. with S. reticulata, Beartooth Plateau, Montana (CRN 139). E. with S. planifolia, Blue Lake, Colorado (CRN 187). F. with S. glauca on the Beartooth Plateau, Montana (CLC 3550). Scale bar = 2 cm. North American Ecology. In the alpine zone in open, mesic, meadows with shrubby and dwarf willows including Salix glauca, S. arctica, and S. planifolia. Jul and Aug. 172 Material examined. USA. COLORADO. Boulder County, Blue Lake, with Salix planifolia, 24 Aug 2018, C. Noffsinger CRN 185 (MONT); loc. cit., with Salix planifolia, 24 Aug 2018, C. Noffsinger CRN 187 (MONT); Niwot Ridge, with S. planifolia, 20 Aug 2018, C. Noffsinger CRN 174 (MONT); Clear Creek/Summit Counties, Front Range, Loveland Pass, 9 Aug 2000, C. Cripps CLC 1488 (MONT); Pitkin/Lake Counties, Sawatch Range, Independence Pass, with dwarf Salix, 3 Aug 2000, V. Evenson DBG-F- 020848 (DBG); loc. cit., with S. planifolia, 6 Aug 2000, C. Cripps CLC 1464 (MONT); loc cit., with S. planifolia and S. glauca, 6 Aug 2000, C. Cripps CLC 1466 (MONT); loc. cit., with shrubby and dwarf Salix, 6 Aug 2006, V. Evenson DBG-F-023519 (DBG); San Juan County, San Juan Range, Black Bear Pass, with S. arctica, 11 Aug 2001, C. Cripps CLC 1719 (MONT); Cinnamon Pass, with S. arctica, 27 July 2002, C. Cripps CLC 1808 (MONT); Mineral Basin, with S. arctica, 7 Aug 2001, C. Cripps CLC 1666 (MONT). MONTANA. Carbon County, Beartooth Plateau, Highline Trailhead, with S. reticulata and S. planifolia, 8 Aug 2017, C. Cripps CLC 3588 (MONT); Birch Site, with S. glauca, 17 Aug 2017, C. Cripps CLC 3550 (MONT); loc. cit., with shrubby Salix, 19 Aug 1999, C. Cripps CLC 1382 (MONT); WYOMING. Park County, Beartooth Plateau, Billings Fen site, with Salix species, 7 Aug 2018, C. Noffsinger CRN 137 (MONT); Highline trailhead, with dwarf Salix, 8 Aug 1998, C. Cripps CLC 1218 (MONT); loc. cit., 8 Aug 1998, C. Cripps CLC 1219 (MONT); loc. cit., with S. planifolia and S. reticulata, 10 Aug 2018, C. Noffsinger CRN 156 (MONT); Frozen lakes, with S. planifolia and S. glauca, 22 Aug 2017, C. Cripps CLC 3597 (MONT); loc. cit., with S. planifolia, 22 Aug 2017, C. 173 Cripps CLC 3601 (MONT); loc. cit., with S. reticulata, 8 Aug 2018, C. Noffsinger CRN 139 (MONT). Observations. Russula subrubens is located in subsection Xerampelina, along with the well-known edible, R. xerampelina; species in this group are recognized by a red-brown pileus, strong fishy odor, and green reaction of the context to ferrous sulfate (Adamčík and Knudsen 2004). Russula subrubens was originally described as Russula graveolens var. subrubens from Elsehoved, Denmark and validated by Lange (1938, 1940); the variety was later raised to the species rank (Bon 1972). It was later reported from subalpine habitats with Alnus and Salix (Bon 1972, 1988). The similar R. chamiteae, was described from alpine regions of the Western European Alps (Kühner 1975), and this name became common in Arctic and alpine literature. The two species were synonymized (Knudsen and Stordal 1992) and R. subrubens is now considered to occur in both temperate and alpine habitats. Adamčík and Knudsen (2004) agreed with the synonymy based on morphology and noted that the association with Salix species appears to override habitat or elevation. They designated an epitype for R. subrubens from Western Jutland (Denmark) because the holotype consisted of only a spore print. Later, Adamčík et al. (2016b) examined subsection Xerampelinae and confirmed earlier conclusions using ITS sequence data because the holotype of R. chamiteae Kühner fell into the R. subrubens clade (ITS2 only). The presence of R. subrubens in Italy, Norway, France, Denmark, and Austria was molecularly confirmed. Our phylogenetic analysis produced a strongly supported clade that included sequences from western Jutland, 174 Denmark, the epitype locality of R. subrubens (Adamčík et al. 2016b), and several sequences from the Rocky Mountain alpine zone. In the central and southern Rocky Mountains, R. subrubens is recognized by the features described for the subsection, plus an association primarily with shrubby Salix in open mesic meadows. However, the pileus color and diameter are highly variable in our collections; large specimens were adjacent to small ones (FIG. 15). Occasionally, the fishy odor is absent in fresh collections, but usually becomes apparent on drying. All our collections are from alpine habitats. The similar, R. cf. pascua also occurs in alpine habitats of the central and southern Rockies. The pileus of this species is dominated more by red and magenta colors, the spores are significantly longer and wider (av 9.1 × 7 µm) according to a two sample t-test (length p-value = < 0.001, width p-value = < 0.001 based on 80 measurements from each species), the pileus is usually smaller, and it occurs more with dwarf willow, specifically S. reticulata. Russula subrubens can be separated from the closely related R. pasuca, which has inflated terminal cells of hyphae in the pileipellis near the pileus center that are lacking in R. subrubens (Adamčík et al. 2016b). Inflated terminal cells of the pileipellis near the pileus center were also absent in our collections. Phylogenetically R. subrubens is closely related to, but distinct from, our R. cf. pascua in the Xerampelinae clade (FIG. 3). The two species appear to overlap in habitat, but their host associations indicate that R. subrubens may be found more in open mesic meadows with S. planifolia; whereas, R. cf. pascua is usually found with S. reticulata. Kühner (1975) originally separated R. chamiteae (now synonomous with R. subrubens) from R. 175 pascua based on a weak reaction in sulfovanillin for the former and no reaction in the latter (Kühner 1975), which is consistent with our observations of these species. Based on morphology, R. subrubens has been reported in Arctic and alpine areas of Denmark (Adamčík et al. 2016b), the Alps, Pyrenees, Carpathians (Adamčík and Knudsen 2004; Ronikier and Adamčík 2009; Adamčík et al. 2016b), Fennoscandia (Adamčík and Knudsen 2004; Adamčík et al. 2016b), Greenland (Adamčík and Knudsen 2004; Borgen et al. 2006), Slovakia (Adamčík 2001), and Poland (Adamčík and Knudsen 2004; Ronikier and Adamčík 2009), this does not include reports of R. chamiteae. This is the first report of R. subrubens under this name in the Rocky Mountains where it appears to be quite common in alpine regions of Colorado, Montana, and Wyoming. It has been reported under the name R. cf. pascua in the Rocky Mountains (Cripps and Horak 2008) and has likely been reported under other names (i.e. R. chamiteae). It is now confirmed to have an intercontinental distribution in Arctic and alpine habitats of North America and Europe. 176 Figure 16. Scanning electron microscope photographs of basidiospores from species in the Russula core clade and Brevipes clade. A. Russula nana (CLC 3619). B. R. montana holotype (MICH 12231). C. R. laccata (CLC 3617). D. R. altaica (CLC 1618). E. R. laevis (CLC 1690). Scale bar = 10 µm. 177 Figure 17. Scanning electron microscope photographs of basidiospores from species in the Russula crown clade. A. Russula saliceticola (CLC 3616). B. R. subrubens (CLC 3550). C. R. cf. pascua (CLC 2274). D. R. heterochroa (CLC 1723). Scale bar = 10 µm. 178 Figure 18. Compound microscope photographs of basidiospores. A. Russula nana (CLC 3619). B. R. montana (MICH 12231). C. R. laccata (CLC 3617). D. R. altaica (CLC 1618). E. R. saliceticola (CRN 184). F. R. subrubens (CRN 156). G. R. cf. pascua (CRN 146). H. R. purpureofusca (CLC 3820). I. R. heterochroa (CLC 1723). H. R. laevis (CLC 1883). Scale bar = 10 µm. 179 Figure 19. Maps of Russula species distributions Set 1. Collection locations confirmed in the phylogenetic analyses are indicated by a red dot. Longitude (lon) and latitude (lat) are indicated on the X and Y axes, respectively. A. Russula cf. pascua. B. R. subrubens. C. R. saliceticola. D. R. purpureofusca. 180 Figure 20. Maps of Russula species distributions Set 2. Collection locations confirmed in the phylogenetic analyses are indicated by a red dot. Longitude (lon) and latitude (lat) are indicated on the X and Y axes, respectively. A. Russula laccata. B. R. nana. C. R. montana. D. R. altaica. 181 Figure 21. Maps of Russula species distributions Set 3. Collection locations confirmed in the phylogenetic analyses are indicated by a red dot. Longitude (lon) and latitude (lat) are indicated on the X and Y axes, respectively. A. Russula laevis. B. R. heterochroa. 182 Key 1. Russula in the central and southern Rocky Mountain alpine zone with Salix, Dryas, and Bistorta (one subalpine with conifers) 1 Flesh dense. Pileus 40–80 mm, depressed with inrolled margin at first, dingy cream- colored or whitish, buff, rough; lamellae decurrent, dark cream at maturity; stipe short, dingy cream, solid; odor fruity; taste acrid; spores av 8.7 × 7.2 µm; primarily with Dryas………………………………………………………………………..…..…R. laevis 1’ Flesh fragile. Pileus convex, applanate, or indented, more highly colored; lamellae attached not decurrent…………………………………..…………………..………….….2 2 Spore print white (possibly tinged yellowish). Lamellae primarily whitish at maturity or slightly yellow; odor faint, not distinctive; taste slightly to distinctly hot; spores fully reticulate or subreticulate………..………………………………………………………...3 2’ Not with the above combination of features; spore print yellow; lamellae yellow at maturity; taste mild or slightly hot; spores with isolated warts, occasionally subreticulate……………………………………………………………………………….5 3 Pileus 15–45 mm, dark magenta, maroon, purplish with almost blackish center; lamellae white at maturity; stipe white; odor faint; taste strongly acrid; spores av 8.1 × 6.2 µm; with dwarf Salix or other willows …………………………..……....….R. laccata 3’ Pileus more red without dark center, fading to white or ocher in areas; with Salix or conifers.…………………………………………………... …………………………....…4 4 With dwarf Salix in alpine areas. Pileus 12–35 mm, cherry red fading to white in areas; lamellae white at maturity; stipe white, 6–26 × 5–13 mm; odor indistinct; taste slightly to distinctly hot; spores av 7.7 × 6.3 µm………………………..………..……......….R. nana 4’ With conifers near treeline. Pileus larger, 35–70 mm, cherry red with ocher areas; lamellae white or with slight yellow tinge at maturity; stipe white, 20–50 × 10–20 mm; odor indistinct; taste hot; spores av 8.0 × 6.3 µm …………………………..…R. montana 5 Odor fishy; taste mild, rarely hot; bluish green reaction in FeSO4; spores subreticulate, rarely fully reticulate…….……………………….....………………….…..6 5’ Odor not fishy; taste mild to slightly hot; lacking a reaction to FeSO4; spores mostly with isolated warts that can cluster or form branches, rarely subreticulate.………………7 6 Pileus 10–80 mm, reddish brown, mottled with brown, red, copper, ocher, and hints of green possible, matt; lamellae cream to ocher; stipe white, cream, or yellow, occasionally with brown or pinkish areas, 10–60 × 10–30 mm; odor fishy; taste mild; spores av 8.5 × 6.7 µm; usually near Salix planifolia, reported elsewhere at or below treeline with Salix…………………………………………………………………….….....R. subrubens 6’ Pileus 25–35 mm, with more red, dark red, and magenta colors, occasionally with hints of green; lamellae yellow to yellow brown; stipe white or off-white, with brownish yellow stains near base, slightly pink upon drying, 15–20 × 8–20 mm; odor fishy; taste 183 mild or hot; spores av 9.1 × 7 µm; in alpine habitats, usually near Salix reticulata…………………………………………………………...………....R. cf. pascua 7 Primarily with Betula, near treeline; pileus 10–40 mm, primarily magenta, mottled with green, yellow, and pink; lamellae white, occasionally graying; stipe white, with or without pink blush, 20–30 × 5–15 mm; odor indistinct; taste mild or bitter; spores composed completely of isolated warts, av 7.9 × 6.1 µm......……………………R. altaica 7’ Primarily with Salix or Dryas in alpine habitats; taste mild or slowly acrid; pileus deep magenta, dark reddish, occasionally yellow brown…………………………...…….8 8 Odor lacking or indistinct. Stipe often with pink tint. Spores primarily with isolated warts, but fine lines and ridges possible. Pileus 10–55 mm, deep red, grayish ruby, almost back in center; lamellae cream to pale yellowish, edge occasionally reddish near cap margin; stipe white, often with pink tint, 15–60 × 5–25 mm; taste mild; spores av 8.9 × 7 µm; pileocystidia lacking lateral protrusions; with Salix species…………………………………………..…………………………...R. saliceticola 8’ Odor fruity or of geraniums. Stipe lacking pink tints. Spores with isolated warts or spines; pileocystidia with abundant lateral protrusions …………………………….…9 9 Spores on average 8.5 × 7 µm; pileus 20–45 mm, reddish brown, deep magenta, or occasionally yellow-brown; lamellae cream to dark yellow-cream; stipe white, off-white, or yellow; odor indistinct or of geraniums; taste slowly hot; with Salix species…………………………………………………………………...R. purpureofusca 9’ Spores larger, on average 10.5 × 8.4 µm; pileus 20–40 mm, variable, magenta, ruby, reddish brown, ocher brown; lamellae white to yellow cream, thick; stipe white; odor fruity or faintly fishy; taste mild; with Dryas octopetala…………..………R. heterochroa Discussion This research confirmed eight species of Russula in the alpine zone, one from the alpine-subalpine transition zone and one from subalpine habitats in the Rocky Mountains from Montana to Colorado. Five of the Russula species from the Rocky Mountain alpine (R. nana, R. cf. pascua, R. saliceticola, R. heterochroa, and R. purpureofusca) can be referred to as primarily Arctic and alpine species because they have been collected only in these habitats near shrubby and dwarf Salix species or Dryas in the northern hemisphere. Three species, R. laccata, R. laevis, and R. subrubens were collected only in 184 alpine areas of the Rocky Mountains, mostly with Salix, in this study; however, all have been reported above and below treeline in Europe (Ortega and Esteve-Raventós 2001; Adamčík and Knudsen 2004; Adamčík et al. 2016b, 2019). Even in subalpine habitats, R. subrubens is believed to be strictly associated with Salix (Adamčík et al. 2016b); this may also be the case for R. laccata, which has been reported with S. atrocinerea below treeline (Ortega and Esteve-Raventós 2001). It is possible that R. laccata, R. laevis, and R. subrubens are also present in subalpine habitats in the Rockies, but none have been reported below treeline. Russula cf. pascua is a potential North American endemic, but more research is necessary to confirm this. Russula altaica is reported from the transition zone between alpine and subalpine habitats where it was associated with Betula; it was only collected at Blue Lake, Colorado. Russula montana was the only species identified that was first described from North America (Perigo Colorado, USA) (Shaffer 1975). In our analysis, R. montana was shown to have an intercontinental distribution in subalpine habitats where it associates with conifers. Most matching collections from Europe were originally identified as R. grisescens, which may be a synonym. This research adds first reports of six species (R. altaica, R. heterochroa, R. purpureofusca, R. saliceticola, and R. subrubens) for the Rocky Mountain alpine and two species (R. heterochroa and R. purpureofusca) for North America. Our first hypothesis stated that the Russula species reported from the Rocky Mountain alpine will be the same as those reported in Arctic and alpine habitats of Europe, Asia, and Arctic Islands, based on molecular and morphological data, even though these would be disjunct populations. Our results indicate that all the species we 185 report from the Rocky Mountain alpine were originally described from Europe or Asia with one exception. Russula cf. pascua formed a strongly-supported monophyletic clade containing only collections from the Rocky Mountain alpine; therefore, it may be endemic to this region. However, if this species is found to be conspecific with R. pascua, then it too would have been originally described from Europe. In addition, we report Russula montana, which was originally described from Colorado where it is a known subalpine species with conifers (Shaffer 1975). The second hypothesis stated that the Russula species present in the Rocky Mountain alpine will have large, intercontinental distributions similar to the Arctic and boreal species studied molecularly in Europe (Adamčík et al. 2019; Caboň et al. 2019). Our results confirmed that seven species of Russula have intercontinental distributions in Arctic and alpine habitats of North America and Europe or Asia, which confirms conclusions from earlier studies based on morphology (Lamoure et al. 1982; Knudsen and Borgen 1982; Borgen 1993; Elborne and Knudsen 1990; Adamčík and Knudsen 2004). Species molecularly confirmed to have large, intercontinental distributions in Arctic and alpine habitats are R. nana, R. laccata, R. subrubens, R. saliceticola, R. purpureofusca, and R. laevis. Russula heterochroa was identified based on morphological evidence alone due to the lack of availability of type or reference sequences for this species; but the collections studied formed a strongly-supported, monophyletic clade containing collections from Colorado, USA and Svalbard, indicating that it too has an intercontinental distribution. The collections identified as R. cf. pascua form a strongly-supported clade containing only collections from the Rocky Mountain 186 alpine; if this species is found to be conspecific with R. pascua it would also have an intercontinental distribution in Arctic and alpine habitats; if not it may be endemic to the Rocky Mountain alpine. The subalpine species R. montana/grisescens also appears to have an intercontinental distribution in subalpine and boreal habitats of North America and Europe. Collections were molecularly matched to the type for R. montana which has precedence (Shaffer 1975; Bazzicalupo et al. 2016). However, the type for R. grisescens was not available, so synonymy cannot be absolutely confirmed. North American collections of R. altaica, a transitional zone species that occurs with Betula, were matched to Singer’s collection from the Altai Mountains (Singer originally described the species) (Singer 1951). Intercontinental distributions have also been confirmed for 44 other alpine species, in four ectomycorrhizal genera, found in the central and southern Rockies, even though these are disjunct populations. This includes 15 species of Hebeloma (Beker et al. 2010; Cripps et al. 2019), one Cortinarius (Peintner 2008), 23 species of Inocybe (Cripps et al. 2010; Larsson et al. 2014; Larsson et al. 2018; Cripps et al. 2019), and five species Lactarius (Barge et al. 2016; Cripps and Barge 2016). Therefore, large intercontinental distributions appear to be a trend in Arctic and alpine ectomycorrhizal genera and implies that species maintain large, intercontinental populations. This is in contrast with data from other habitats that show most ectomycorrhizal fungi have small geographic distributions (Bazzicalupo et al. 2019) and that spore dispersal from basidiocarps appears limited (Fischer et al. 2010). Although we do not have data to explain the large, intercontinental distributions found here for alpine species of Russula in the Rocky Mountains, we believe that one of the most parsimonious 187 explanation is that Arctic and alpine fungi are following their primary host, dwarf and shrubby species of Salix. This seems plausible considering that Russula species have the ability to follow their host in response to changes in climate (Looney et al. 2019, 2020). Other possible factors that could help explain intercontinental distributions of Arctic and alpine fungi include historical changes in glaciation that influence ectomycorrhizal communities and their host (Timling et al. 2012; Barge et al. 2016) or long-distance, transoceanic dispersal as maintaining large populations (Geml et al. 2012; Timling et al. 2014). Our third hypothesis stated that the Russula species in the Rocky Mountains have independently colonized alpine habitats and do not form a monophyletic group. This research has confirmed that the Russula species in the Rocky Mountain alpine are in two distinct clades, seven in the Russula clade (R. subgenus Russula) and one in the Brevipes clade (R. subgenus Brevipes). Within R. subgenus Russula, no two species form a sister group. Therefore, the data suggest that Russula species in the Rocky Mountain alpine do not share a most recent common ancestor that is unique to this group. The molecular, morphological, and ecological data presented here indicate that the Russula species present in the Rocky Mountain alpine have distinct evolutionary backgrounds and colonization of alpine habitats appears to have happened independently in multiple Russula lineages. This result implies that the potential of Russula species to colonize alpine habitats is present in multiple linages. Most of the Rocky Mountain alpine species have phylogenetically closely related relatives found in subalpine or boreal habitats. It is possible that the transition from subalpine to alpine habitats involved a host switch from 188 conifers to the hosts commonly found in alpine habitats (Salix, Betula, and Dryas) and the ability to survive in the harsh abiotic environmental conditions present in the alpine (colder temperatures, poor soil composition, etc.). Research has shown that host switching from Pinaceae to angiosperms is about 15 times more likely than in the opposite direction for Russula species and that host switching may drive species diversification (Looney et al. 2016), which indicates that certain Russula species could overcome this barrier. Looney et al. (2016) proposed orogenesis events as a possible mechanism to explain host switching from Pinaceae to angiosperms, which could lead to species diversification. This conclusion is fitting in the context of this study, which is concerned with alpine fungi whose evolutionary history was also likely shaped by orogenesis events. Another possibility is that alpine fungi and their close relatives were already associated with angiosperm hosts like Salix in subalpine and boreal zones and did not need to switch hosts to colonize alpine habitats, which may be the case for R. subrubens and R. laccata (Ortega and Esteve-Raventós 2001; Adamčík and Knudsen 2004; Adamčík et al. 2016b, 2019). Subalpine fungi are often reported to be associated with conifers even when Salix is present. Researchers need to report ectomycorrhizal plant hosts as accurately as possible and find creative ways to confirm host associations; however, this will be difficult in diverse temperate zones where many potential hosts exist. Russula species are known to be one of the most abundant ectomycorrhizal associates of Arctic and alpine Betula (Deslippe and Simard 2011). Russula species have been reported to form ectomycorrhizal associations with Betula nana L., B. glandulosa, 189 and B. pubescens Ehrh. in Alaska, Canada, Fennoscandia, and Greenland (Miller et al. 1973; Miller et al. 1982b; De Groot et al. 1973; Elborne and Knudsen 1990; Deslippe et al. 2011; Deslippe and Simard 2011). Betula is rare in the Rocky Mountain alpine zone, and this study found only R. altaica with Betula shrubs at treeline. It is likely that there are more species of Russula that occur with Betula in the Rocky Mountain alpine zone that await discovery. Several species of Russula from treeline habitats in Alaska were found in association with Betula, including R. cf. alpigenes, R. cf. sphagnophila, R. intermedia, and R. aff. vinosa. It is likely that these species extend into the alpine where Betula occurs. These species were only collected a few times and are included in the phylogenetic trees produced in this study (FIGS. 2, 3, 4). A key to these red-capped Russula from Alaska, mostly with Betula at treeline, is included in the Addendum along with descriptions of each species. Even though identification of Russulas to species can be difficult without molecular data, this work has confirmed some morphological trends in the genus that aid in identification. Russula subrubens and R. cf. pascua are in R. subsection Xerampelinae and have features consistent with this group (Adamčík and Knudsen 2004), including yellow spores, a fishy odor, a green reaction of the context to ferrous sulfate, and a mild taste. This combination of features is unique to the subsection and can be used for identification at this level; however, identification to species can be complex (see Adamčík et al. 2016b). Rocky Mountain alpine species R. nana and R. laccata have reddish to maroon pilei, white reticulate spores, and an acrid taste, which is specific to these taxa when only 190 considering the species examined in this study. However, many other species with these features occur in the subalpine zone that are not fully understood. One, R. montana, was confirmed in this study; this species also has a reddish to maroon pileus, white reticulate spores, and an acrid taste. The well-known R. emetica that can also occur near treeline shares these features, and care is needed to delineate this species and other look-alikes in the transition zone (see Bazzicalupo et al. 2016). Two species in the Rocky Mountain alpine zone, R. saliceticola and R. purpureofusca, typically have deep magenta pilei and a yellow spore print (although some collections of R. purpureofusca can have an olive-yellow pileus), and a non-fishy odor; these characters can be used to aid in identification, and these two species are separated based on microscopic characteristics (see species observations or key for identification to species). One species, R. laevis in R. subg. Brevipes, occurs in the Rocky Mountain alpine and has a unique morphology compared to the other species studied. Russula laevis has a white to brown concave pileus, compact flesh, a white spore print, and an acrid taste, which together are diagnostic for R. laevis in the Rocky Mountain alpine. However, in the subalpine zone, many species in R. subgenus Brevipes including R. brevipes, R. chloroides, and R. delica also share these characteristics, although fruiting bodies are typically larger. However, one collection of small fruiting bodies from a treeline habitat in Alaska, turned out not to be R. laevis, highlighting that size is not always reliable. Russula subsection Lactarioideae, which contains these species, has been morphologically examined but has not been subjected to rigorous molecular phylogenetic 191 analysis (Buyck and Adamčík 2013). In our analysis of R. subg. Brevipes (FIG. 5) sequences representing the UNITE species hypotheses for R. chloroides (AF418605) and R. delica (UDB025023) were distinct from the Rocky Mountain alpine species R. laevis, as has been previously shown (Adamčík et al. 2019). The sequence representing the UNITE species hypothesis R. delica forms a clade by itself and BLAST search for similar sequences found only uncultured Russulaceae isolates that matched with a high Max Score. This is concerning, and warrants investigation due to the prevalence of sequences in public databases labeled R. delica that do not match the UNITE species hypothesis. Russula delica is known to encompass a “wide species concept” because 11 varieties have been described and there is some confusion regarding the interpretation of Fries’s (1838) original description of the species (Buyck and Adamčík 2013). The R. chloroides complex contains two strongly-supported clades with numerous collections labeled R. chloroides, R. brevipes, and R. delica from public databases (FIG. 5). Buyck and Adamčík (2013) indicated that the American R. brevipes is similar to the European R. chloroides based on morphology; however, any potential synonymy needs to be verified using type material. This research indicates that the R. chloroides complex, as defined here, contains two strongly-supported clades, both of which show intercontinental distributions. The distribution of these Russula species needs to be re-evaluated and results could cast doubt on the hypothesis suggesting that subalpine ectomycorrhizal species lack large, transatlantic distributions (Buyck and Hofstetter 2011). There were a few trends that emerged in the spore morphology of Russula species in the Rocky Mountains. The three closely related species R. montana, R. nana, and R. 192 laccata all have spores with fully reticulated ornamentation (FIGS. 16, 18). Russula subrubens and R. cf. pascua in Russula subsection Xerampelinae both have partially reticulated spores with some isolated warts (FIGS. 17, 18). The same spore morphology is also present in R. saliceticola, which is found in the clade sister to R. subsection Xerampelinae. The remaining species (R. altaica, R. heterochroa, R. laevis, and R. purpureofusca) all have spores with primarily isolated warts (FIGS. 16, 17, 18) and these species are found in various other clades. The observed trends in spore ornamentation for closely related species indicate that spore ornamentation may be connected to species evolutionary history, at least at small scales. However, connecting the two would require ancestral character state reconstruction of a detailed, multi-locus, phylogenetic dataset including species from all major clades and habitats along with careful observations of spore morphology. Identification of Russula species in western North America is challenging due to a lack of reliable information and expertise (Buyck 2007; Buyck et al. 2015). This research approached the genus Russula from an ecological perspective by focusing on species occurring above treeline with alpine host plants such as Salix and Dryas. This approach helped to mitigate some of the broader taxonomic issues, but other problems arose that are inherent to alpine research in the Rockies. The Rocky Mountain alpine is remote and difficult to access, making sampling challenging. In addition, the fruiting season is short, and basidiocarps are extremely small, making them difficult to find. Also, a dry, continental climate means potential for droughts and smoke from lower elevation fires, which can make collecting difficult. In some years, alpine fungi are virtually absent. The 193 high elevations are also prone to lightning storms and snow at any time of year. These issues make thorough sampling of alpine fungi in the Rocky Mountains an arduous task. Fortunately, this study had access to collections of Russula and other alpine fungi collected in the Rockies over the last 20 years. Another problem is that much of the literature published on Arctic and alpine Russula is written in a variety of languages including French (Favre 1955; Romagnesi 1967; Kühner 1975; Bon 1993, 2000), Catalan (Bon and Ballarà 1996), Italian (Jamoni 1995; Sarnari 1998–2005), and Danish (Borgen 1993). This literature can be easily translated using online resources, but these iterations are not perfect, and taxonomists rely on small differences among taxa that can be lost in translation. On a broader scale, the diversity of Russula has not been adequately addressed in western North America. This research adds first reports of six species (R. altaica, R. heterochroa, R. purpureofusca, R. saliceticola, and R. subrubens) for the Rocky Mountain alpine, two of which (R. heterochroa and R. purpureofusca) are also first reports for North America. Much remains to be discovered for Russula in the conifer and deciduous forests at lower elevations. The genus Russula has been found to be most diverse in temperate regions (Looney et al. 2016), and these regions have not been adequately sampled, particularly in western North America. We produced the broad phylogeny (FIG. 2) in order to understand the placement of Rocky Mountain alpine species in the larger infrageneric classification of Russula. However, a more thorough sampling of Russula species diversity in temperate regions will influence our understanding of ecological, biogeographical, and evolutionary trends in the genus, 194 which would likely extend to alpine species. It is worth noting that five of the species we found in the Rocky Mountain alpine are located in the Russula crown clade, which also contains the highest species diversity in the genus, including many tropical taxa (Looney et al. 2016). Research has indicated that morphological studies alone often fail to recognize the true diversity of Russula in a region (Buyck 2007), which also appears to be the case for Russula in the Rocky Mountain alpine. Previous morphological research discovered four species of Russula in the Rocky Mountain alpine (Moser and McKnight 1987; Cripps and Horak 2008), while this research doubled the number of species known in the region. Results emphasize the importance of detailed systematic studies that combine morphological, molecular, and ecological data. Fortunately, this approach is already being used for Russula (Adamčík et al. 2016b, 2019; Buyck et al. 2018; Caboň et al. 2019; Buyck et al. 2020) and is further supported by recent research that found a loose connection between morphological character states and species boundaries within Russula (Bazzicalupo et al. 2017). Some general trends in this study also emerged when comparing our molecular and phylogenetic data to other studies and public databases. Based on our phylogenetic analyses the ITS region alone was sufficient at separating and supporting the species studied here and no major changes in species clades were observed when adding the RPB2 gene region to the analysis, although the support and topology of internal nodes varied slightly. The RBP2 gene region was also more difficult to successfully sequence in the Russula studied here. All collections for which the ITS region were successfully 195 sequenced, the RPB2 region was also attempted, but 55 RPB2 sequences could not be obtained (40% less than for the ITS). The ITS-RPB2 gene combination was not sufficient for resolving deep phylogenetic relationships in the broad phylogenetic analysis of Russula (FIG. 2), mainly at the subgeneric level, even when several reference sequences contained additional loci (LSU and RPB1). This result is also evident in earlier studies using only one or a few gene regions where support is lacking for deep nodes (Miller and Buyck 2002; Buyck et al. 2008), stressing the importance of complete, multi-locus phylogenies when studying the evolutionary relationships within Russula, as was done in Looney et al. (2016). Our work uncovered some misidentifications in public databases such as GenBank and UNITE, most notably with R. nana (see observations for R. nana), but many other errors were found that could not be included here. Given that the morphology in Russula is complex (Romagnesi 1967; Singer 1986; Sarnari 1998–2005), some types are difficult or impossible to obtain, and sequence-based identification is known to be insufficient and difficult (Hofstetter et al. 2019), identification based on sequences in public databases is unlikely to provide confident results until the quality of these databases can be improved. Therefore, we stress the recommendations outlined in (Hofstetter et al. 2019), which focus on the importance of proper sequence annotation during submission, to increase the value of sequence-based identification, especially within the genus Russula. Many of the species of Russula found in the Rocky Mountain alpine are morphologically and molecularly similar, with many closely related species being more than 97% similar in ITS sequences. For example, when comparing ITS sequence 196 similarity for the species included in our phylogenetic analysis, R. nana and R. montana are approximately 98.6% similar, but have consistent molecular differences; R. laccata and R. atrorubens are approximately 98.3% similar; R. altaica is approximately 98.2% similar to R. gracillima; R. saliceticola is approximately 98.2% similar to R. nitida; R. laevis is approximately 98.9% similar to R. cf. brevipes; and species in the R. clavipes complex range from 98.9–99.4% in similarity. The other three species studied here (R. heterochroa, R. purpureofusca, and R. subrubens) are all 97% similar or less in the ITS region when compared to closely related species included in our phylogenetic analyses. The low genetic variation found here is similar to what was observed in closely related alpine species of Lactarius from the Rocky Mountain alpine (Barge et al. 2016) and casts doubt on the effectiveness of 97% sequence similarity cutoffs for species delineation in environmental studies (e.g. Geml et al. 2012; Timling et al. 2012, 2014), especially for alpine members of the Russulaceae. Environments like the Arctic and alpine are seeing some of the most severe negative impacts of climate change. As shrub expansion into these cold-dominated habitats continues in conjunction with changes in climate, the composition of ectomycorrhizal fungal communities will begin to shift. The functioning of individual ectomycorrhizal species is not redundant, with various species, even within the same genus, showing preferences for different forms of nitrogen (Antibus et al. 2018), indicating that the shift of individual species could have impacts on overall ecosystem function. These conclusions emphasize the importance of continuing research aimed at understanding the diversity and function of Arctic and alpine ectomycorrhizal fungi prior 197 to large environmental shifts. A deeper taxonomic understanding of Arctic and alpine fungi will increase the value of models aimed at understanding the evolution of Arctic and alpine communities (Gardes and Dahlberg 1996). This research increased our knowledge of Russula diversity in alpine habitats of the Rocky Mountains, adding value to models that use members of the Russulaceae to understand ecosystem function and diversification of ectomycorrhizal fungi in all habitats (Looney et. al. 2018). 198 Addendum – Alaskan species of Russula Several collections in the phylogenetic analyses are from subalpine, treeline, and alpine regions of Alaska (TABLE 5; FIGS 2, 3, 4). Initially, we believed that our Russula collections from Alaska would extend the distribution of some species described from the central and southern Rocky Mountains, and thus would be included in the main study. Other research has reported R. xerampelina var. pascua (R. pascua) and Russula emetica var. alpestris (possibly synonymous with R. nana) based on morphology in Arctic areas of Alaska with Salix (Miller et al. 1973). These are species we report from Colorado, Montana, and Wyoming. It seems highly likely that some of the Russula species from alpine areas of the central and southern Rockies would also occur in alpine areas of Alaska. However, our research confirmed only R. purpureofusca in alpine areas of both Colorado and Alaska where it occurs with Salix and Dryas. Our other collections from Alaska were morphologically and molecularly distinct from those in the central and southern Rocky Mountains. This is likely due to minimal and biased sampling in Alaska. Russula cf. alpigenes, which is found near Salix and Dryas, is reported from Alaska but not Colorado, Montana, or Wyoming. If confirmed, this would be the first report of this rare species for North America. We report two collections of R. intermedia, and one each of R. cf. sphagnophila and R. aff. vinosa with Betula at treeline in Alaska. Betula is rare in our main collecting areas, and only R. altaica is reported with Betula from one area of Colorado. Thus, Russula associated with Betula are under-sampled, but they are included here, because they should be considered when collecting near Betula in the central and southern Rockies. 199 All of these species’ clades were resolved as monophyletic and with strong- support in both phylogenetic analyses (FIGS. 2, 3, 4). There is some taxonomic confusion regarding R. cf. alpigenes, which clusters phylogenetically with sequences previously identified by others as R. nana and is sister to R. betularum (FIG. 4); we believe these sequences are incorrectly identified (see observations under R. nana). All of the species from Alaska were consistently resolved in R. subgenus Russula in the phylogenetic analyses (FIG. 2). Four species were consistently resolved in the Russula crown clade (FIG. 3) and one species, R. cf. alpigenes, was consistently resolved in the Russula core clade (FIG. 4). Previous research on Russula suggests the genus is diverse in Alaska (Geml et al. 2010; Geml and Taylor 2013) and is dominant in moist tussock and dry tundra in Arctic Alaska (Morgado et al. 2015). However, Russula is seriously under-sampled and its diversity is poorly known in Alaska. The species described here provide morphological and molecular information regarding just a few of the undoubtedly numerous Russula in the region. A key is provided for their identification, mainly including Russula with Betula near treeline, and descriptions are provided for each species. Key 2. Red-capped Russula from Alaska, mostly with Betula at treeline 1 Spore print white; Pileus ruby red, depressed in center or not; lamellae white at maturity; stipe white with pinkish tints, especially after drying; odor fruity; taste mild to acrid; spores on average 8.9 × 7 µm, reticulate; with Salix and Dryas in the low alpine…...……………………………………………………………….…R. cf. alpigenes 1’ Not with the above combination of features; spore print light to dark yellow; lamellae yellow at maturity, rarely white; taste mild or slightly hot; spores with isolated warts; with Betula or other hosts, one species with Salix…………………………………2 200 2 Taste strongly acrid, occasionally delayed; with Salix; pileus reddish brown, deep magenta, or occasionally yellow-brown; lamellae cream to dark yellow-cream; stipe white, off-white, or yellow; odor indistinct or of geraniums; taste slowly hot; spores on average 8.5 × 7 µm…………………..........................…R. purpureofusca (see Chapter 2) 2’ Taste usually mild…………………………………………………………………..…3 3 Pileus reddish maroon, edges paling to whitish, dished in center, striate almost to center, fragile, thin-fleshed; lamellae white to cream with a pink tinge; stipe clavate, white, with pinkish rosy tint at apex, especially in age; odor absent; spores on average 8.7 × 6.8 µm, with Betula nana, hemlock, and spruce……….………...…R. cf. sphagnophila 3’ Pileus more brown, orange, or deep red, more thick –fleshed, robust, less striate.…….4 4 Spores usually < 8.5 µm long, on average 8.0 × 6.9 µm; Pileus red, reddish orange, salmon, mottled; lamellae white then cream or golden; stipe clavate, robust; odor absent or fruity; taste bitter; primarily with Betula but occasionally near Dryas and Salix in subalpine habitats……………………...………………………………....….R. intermedia 4’ Spores usually > 8.5 µm long; primarily with deciduous hosts including Betula nana, also with hemlock and spruce; Pileus bright red with center fading to ocher then gray; lamellae white, a bit cream in age; stipe long (50–60 × 15–20 mm), white and gray; spores on average 8.7 × 7.1 µm ……………………………………………..R. aff. vinosa Taxonomy of Alaskan Species Russula cf. alpigenes (Bon) Bon FIG. 22 Bull. trimest. Féd. Mycol. Dauphiné-Savoie 32(no. 128):20 1993 Macromorphology. Pileus 20–60 mm wide, shallow convex almost flat, some dished in center, deep ruby red to almost black in center, smooth, greasy (viscid), not striate; margin turned down, entire. Lamellae narrow adnate, narrow to broad, crowded, L = 100 (n = 1), white, then cream, edges concolorus, staining pink near pileus margin or not. Lamellulae absent. Stipe 25–60 × 8–18 mm, long clavate, clavate, but narrower in middle, white, but with a few pinkish stains in age, smooth. Context white, solid; pink under cap cuticle. Odor fruity-sweet or absent. Taste slightly acrid or not, bitter, or mild. Chemical reactions: not available. 201 Micromorphology. Basidiospores no spore print obtained, but reported as white in print (Bon 1993), (7.6–)8.3–8.9–9.5(–10.6) × (6.1–)6.6–7–7.4(–8.2) µm, broadly ellipsoid to ellipsoid, Q = (1.1–)1.2–1.3–1.4(–1.5); ornamentation of amyloid warts and spines, <0.5–0.5 µm high, occasionally fusing, connected by thin line connections and occasionally ridges forming a reticulum, rarely isolated, clustered, branched or subreticulate; apiculus truncate, inamyloid; suprahilar plage small to medium, weakly amyloid, difficult to observe. Basidia (45.7–)49.2–53.7–58.3(–63.5) × (8.9–)9.8–10.5– 11.2(–11.4) µm, clavate-pedicellate, 4-spored, sterigma swollen and finger-like; contents gelatinous and granular, lipid drops occasionally present, no reaction in sulfovanillin; basidiola clavate, granular, often filled with lipid drops. Hymenial cystidia (48.3–)53.7– 66.5–79.4(–101.6) × (7.6–)7.6–8.5–9.5(–10.2) µm, fusiform, cylindrical, pedicellate, rarely lanceolate, often mucronate, with 1.3–7.6 µm long appendage; contents gelatinous, granular, and crystalline, staining dark red-brown to black in sulfovanillin. Pilepellis sharply delimited from underlying context; weakly gelatinized, ca. 63.5–114 µm deep suprapellis composed of loosely intertangled, erect, hyphae and pileocystidia, well differentiated from subpellis; strongly gelatinized ca. 38.1–101.6 µm deep subpellis composed of densely intertangled, horizontally oriented, hard to differentiate hyphae. Hyphal terminations cylindrical, thin-walled, septate, occasionally tapering at apex, occasionally forking, terminal cells ca. 2–7.6 µm wide, apically obtuse; devoid of contents or lightly granular. Pileocystidia near the pileus margin abundant, cylindrical or clavate, (53.3–)68–108.6–149.1(–203.2) × (5.1–)5.1–5.9–6.8(–7.6) µm, occasionally forking; contents granular, blackening in sulfovanillin. Pileocystidia near the pileus 202 center shorter, occasionally moniliform, more septate, occasionally tapering near apex but still apically obtuse, (63.5–)74.9–91.9–109(–114.3) × (5.1–)5.4–6.6–7.8(–8.9) µm; contents similar. 203 Figure 22. Russula cf. alpigenes. A. with Salix and Dryas, Girdwood, Alaska (CLC 3822B). B. with Salix, Girdwood, Alaska (CLC 3821). Photos by Noah Siegel. Scale bars = 2 cm. 204 North American Ecology. Found in the transition zone between subalpine and alpine habitats in Alaska with Dryas and mixed Salix. Aug. Material examined. USA. ALASKA. Anchorage County, Girdwood, with Salix, 22 Aug 2018, C. Cripps CLC 3821 (MONT); loc. cit., with Salix and Dryas, 22 Aug 2018, C. Cripps CLC 3822B (MONT). Observations. Russula fragilis var. alpigenes was originally described by Bon in 1990 from the European Alps with Salix herbacea, shortly after he raised R. alpigenes to the species rank (Bon 1993). Our species matches Bon’s (1993) description. Our collections form a strongly-supported clade with three accessions incorrectly labeled R. nana (UDB015077, UDB016029, and KX579809, FIG. 4) see observations under R. nana and discussion in Chapter 2. This clade is sister to R. betularum. Russula intermedia P. Karst. FIG. 23 Meddelanden af Societas pro Fauna et Flora Fennica 16:38 1888 = Russula intermedia f. mesospora (Singer) Bidaud, Moënne-Locc. & Reumaux, in Reumaux, Bidaud & Moënne-Loccoz, Russules Rares ou Méconnues (Marlioz):285 1996 = Russula lundellii Singer, Lilloa 22:719 1951 (1949) = Russula mesospora Singer, Bull. trimest. Soc. mycol. Fr. 54:161 1938 = Russula pulcherrima S. Lundell & Jul. Schäff., Annls mycol. 36(4):31 1938 = Russula aurantiolutea Kauffman Rep. Michigan Acad. Sci. 11:81. 1909 205 Macromorphology. Pileus 30–65 mm wide, convex to shallow convex, deep red, reddish brown, orange brown, but also mottled with pink, ocher, cream, and yellow tones, smooth, greasy, viscid, or sticky; margin not striate, turning down, entire; cuticle thick, hardly peeling near margin. Lamellae narrowly attached, adnate, crowded, L=100 (n = 2), white then cream, golden yellow, with mostly concolorous edges, but slightly grayish in one collection, occasionally forking. Stipe 20–50 × 10–20 mm at apex, 10–25 mm at base, clavate, white, smooth, matt, without graying. Context white and solid. Odor absent or fruity. Taste mild, but bitter. Exsiccata: Pileus uniformly violet brown or mottled with violet brown, yellow, orange, and brown, ocher in center. Lamellae light yellow brown, ocher-yellow, lighter yellow near gill margin with occasional hints of gray. Stipe white, yellow brown, light brown, brown, with more brown stains near base. Chemical reactions: Not available. Micromorphology. Basidiospores no print obtained (see observations), (7.1–)7.5– 8.0–8.5(–9.2) × (6.1–)6.4–6.9–7.3(–7.6) µm, subglobose to broadly ellipsoid, Q = (1.1– )1.1–1.2–1.2(–1.3); ornamentation of medium to high density, primarily of isolated, amyloid warts, <0.5 µm high, warts often fusing but still individually recognized, occasionally branched or subreticulate; apiculus inamyloid, truncate; suprhilar plage small to large, amyloid. Basidia (45.7–)51.5–59.4–67.4(–81.3) × (8.9–)9.6–10.4–11.2(– 12.2) µm, clavate-pedicellate, 4-spored, some possibly 2-spored; contents granular, gelatinous, or absent, lipid drops present, vibrant light yellow orange to bright red in sulfovanillin, contents not staining; basidiola clavate, usually occasionally filled with lipid drops and granular contents. Hymenial cystidia (58.4–)66.7–84.3–102(–122) × (6.4– 206 )8.2–10.1–12.0(–12.7) µm, fusiform, clavate, or irregular, rarely with tiers gradually decreasing in size near apex, typically pedicellate, often mucronate, with 2.5–15.2 µm long appendage; contents granular, gelatinous, and crystalline, staining black in sulfovanillin. Pileipellis sharply delimited from the underlying context; suprapellis well- defined, weakly gelatinized, ca. 50–76 µm deep, composed of interwoven and erect hyphae and pileocystidia; subpellis gelatinized, ca. 127–500 µm deep, composed of tightly interwoven, hard to differentiate, horizontally oriented hyphae. Hyphal terminations swollen, forking, or deformed, flexuous, septate; terminal cells ca. 3 µm wide, apically obtuse or subacute. Pileocystidia near the pileus margin with 0–3 septa, cylindrical, slightly or strongly clavate, or tapering, occasionally with visible cell wall 1– 2 µm thick, rarely forking, similar to hyphae but containing contents, (68.6–)85.6–139– 192.4(–254) × (2.5–)4–5.1–6.3(–8.9) µm; contents amorphous, granular, irregular, and gelatinous, or absent, difficult to measure, granular contents staining red-brown in sulfovanillin. Pileocystidia near pileus center similar in shape to those at margin, but thinner, with more septa (2–4), and possibly less abundant, (73.7–)103.3–130.1–157(– 170.2) × (3.8–)3.9–4.7–5.4(–6.4). 207 Figure 23. Russula cf. intermedia. A. with Dryas, Girdwood, Alaska (CLC 3822). B. with Dryas and Salix, Girdwood, Alaska (CLC 3784). Photos Noah Siegel. Scale bar = 2 cm. 208 North American Ecology. Found in the transition zone between subalpine and alpine habitats in Alaska with Betula, Dryas, and Salix. Aug. Material examined. USA. ALASKA. Anchorage County, Girdwood, Crow Pass, with Dryas and Salix, 9 Aug 2011, C. Cripps CLC 2759 (MONT); with Dryas, 22 Aug 2018, C. Cripps CLC 3822 (MONT); Municipality of Anchorage, Flattop Mountain, with Dryas and Salix, 20 Aug 2018, C. Cripps CLC 3784 (MONT). Observations. Russula intermedia Karsten was originally described from Finland (Karsten 1888); Karsten’s second collection, from a forested area near birch, was later designated as a neotype (Ruotsalainen and Vauras 1994). The species is common in forests of Finland where it occurs with Betula and Picea abies but is rare in northern areas. It is also reported from Denmark, Sweden, and Norway with Betula. Ruotsalainen and Vauras (1994) found the well-established species R. lundellii Singer (Singer 1951) to be the same taxon and synonymized the names, although Sarnari (1998–2005) does not agree with this synonymy. Under the name R. lundellii, the species also has been reported from Germany, France, Spain, Belgium, Estonia and Iceland. Ruotsalainen and Vauras (1994) also synonymized the North American species R. aurantiolutea Kauffman (Shaffer 1970) with R. intermedia, after examining MICH collections. The species has been reported from Eastern and Western U.S. and Canada under this name. Thus, R. intermedia in the broad sense appears to have a wide intercontinental range, primarily in montane to subalpine habitats with Betula. Our R. intermedia collections are all from Alaska, and are characterized by a red, red-brown, or orange-brown pileus mottled with pink, ocher, and yellow tones, a hardly 209 peeling cuticle, yellow lamellae and spores, a white stipe, fruity odor, mild but bitter taste, subglobose to broadly ellipsoid spores with isolated warts (8.0 × 6.9 µm), commonly with thin, cylindrical pileocystidia, and in association with Betula or Dryas. Our description matches that for R. intermedia (Ruotsalainen and Vauras 1994) and that for R. lundellii (Sarnari 1998–2005). The pileus color is noted as variable, ranging from red, reddish orange, orange yellow, to yellow brown in various descriptions that could explain some of the taxonomic confusion. A pink tint on the stipe is sometimes noted but was absent in our collections. Also, an acrid taste was not reported in our specimens, but they were bitter and perhaps not chewed long enough for acrid flavors to develop. The Alaskan collections were from just below or just above treeline with Betula, Dryas, and Salix in the vicinity. Phylogenetically our three collections form a strongly-supported clade with an R. intermedia collection from Norway (KU928147, FIG. 3), the type country, and one from Finland (JV 32189F) morphologically identified as R. intermedia by Jukka Vauras. Phylogenetic analyses produced by Adamčík et al. (2016a) also grouped this reference collection with two other collections from Estonia (UDB011296 and UDB015997). Russula cf. sphagnophila Kauffman FIG. 24 Report Mich. Acad. Sci. 11:86 1909 Macromorphology. Pileus 20–35 mm wide, applanate uplifted or slightly infundibuliform, reddish maroon with blackish center, much lighter red toward margin, bit viscid or just wet, striate almost to center, evenly pleated, edge paling to whitish; margin a bit crenate. Lamellae narrow attached, a bit separated, L = 30–40 (n = 1), white 210 to cream, tinged pinkish in age. Stipe 35–50 × 4–5 mm at apex to 13 mm at base, clavate, white, smooth with pinkish rosy tinge at apex, especially in age. Odor absent. Exsiccata: Pileus dark magenta, almost black in center, much lighter toward margin; lamellae light yellow browning in age; stipe ranging from light brown going toward dark brown in age, with tints of magenta. Chemical reactions: Not available. Micromorphology. Basidiospores no spore print obtained, reported as cream (Woo 1989), (7.6–)7.9–8.7–9.5(–11.2) × (6.1–)6.4–6.8–7.2(–7.6) µm, broadly ellipsoid to ellipsoid, Q = (1.2–)1.2–1.3–1.4(–1.5); ornamentation of prominent, cylindrical, amyloid warts, ca. 1 µm high, some warts merging forming larger obscure warts, not forming a reticulum, but with a few thin lines connecting elements; apiculus tapering to a truncate end, inamyloid; suprahilar plage relatively large, weakly to moderately amyloid. Basidia (35.6–)35.9–41.7–47.4(–55.9) × (10.2–)10.2–11.1–11.9(–12.7) µm, clavate, some very swollen, 4-spored; basidiola clavate. Hymenial cystidia (53.3–)60.1–67.6–75.1(–76.2) × (5.1–)7–8.9–10.8(–11.4) µm, fusiform, pedicellate, mucronate, with long, ca. 10.2–12.7 µm appendage with raised granular or crystalline structures; filled with lipid drops, contents granular or crystalline. Pileocystidia near pileus margin clavate, occasionally septate, forking below a septa, (43.2–)47.9–72.9–98(–114.3) × (3.8–)4.8–5.8–6.7 µm; contents granular, crystalline, or amorphous. 211 Figure 24. Russula cf. sphagnophila with Betula nana, Girdwood, Alaska (CLC 3779), photo Noah Siegel. Scale bar = 2 cm. North American Ecology. Found in the transition zone between subalpine and alpine habitats in Alaska with Betula nana, near Tsuga and Picea. Aug. Material examined. USA. ALASKA. Anchorage County, Girdwood, Crow Pass Road, with Betula nana, Tsuga, and Picea, 19 Aug 2018, C. Cripps CLC 3779 (MONT). Observations. Russula sphagnophila was originally described by Kauffman in 1909 from Cold Spring Harbor, Michigan. Kauffman described it as a fragile species with a 20–45 mm wide, rosy-purple to olive-brown pileus, with a raised margin; white to buff lamellae; and a 40–50 × 7–12 mm, fragile, stipe. Phylogenetically our collection forms a strongly-supported clade with a collection identified as R. sphagnophila by Singer from 212 Russia (MICH 256930, FIG. 3). This clade forms a polytomy with a collection of R. nitida (KU205269) and the weakly-supported R. saliceticola clade. At this time, we elect to call our collection R. cf. sphagnophila because it clusters phylogenetically with Singers collection and its morphology matches that of Kauffman’s original description, with the exception of olive-brown coloration in the pileus. Russula aff. vinosa Lindblad FIG. 25 Svampbok: 67 1901 Macromorphology. Pileus 50–60 mm wide, shallow convex, almost flat, sunken in center, bright cherry red, center paling first ocher then grayish, white, smooth, bit viscid, sticky; margin down, faintly striate. Lamellae narrowly attached, narrowly crowded, L = 120 (n = 1), white, maybe a bit cream in age. Lamellulae absent. Stipe 50– 60 × 15–20 mm, slightly enlarged at base, white, bit graying, smooth, greasy-dry. Odor a bit fruity. Taste not hot or with a tiny tingle. Exsiccata: Pileus maroon, red brown, red, orange brown, occasionally ocher in center. Lamellae cream, ocher, with grayish and brown tints. Stipe white with slight yellow, brown, or gray tints. Chemical reactions: Not available. Micromorphology. Basidiospores no spore print obtained, (8.2–)8.3–8.7–9.1(– 9.2) × (6.6–)6.9–7.1–7.4(–7.6) µm, broadly ellipsoid, Q = (1.1–)1.2–1.2–1.3(–1.3); ornamentation of dense, isolated warts, occasionally merging, with no reticulation; apiculus truncate, inamyloid; suprahilar plage small to large, amyloid. Basidia (48.3– )49.7–52.8–56(–56) × (10.2–)10.5–11.5–12.5(–12.7) µm, clavate, pedicellate, 4-spored; 213 basidiola clavate. Hymenial cystidia (71.1–)75–81–87.2(–88.9) × (8.9–)9–9.7–10.3 µm, clavate, weakly fusiform, rarely mucronate, with small, ca. 2.5 µm long appendage; contents granular and crystalline. Pileocystidia near pileus margin abundant, cylindrical, flexuous, septate, forking often, sometimes multiple times, (53.3–)59.2–89.4–119.6(– 152.4) × (2.5–)2.6–3.4–4.3(–5) µm; devoid of contents. Figure 25. Russula aff. vinosa with Betula nana, Girdwood, Alaska (CLC 3778), photo Noah Siegel. Scale bar = 2 cm. North American Ecology. Found in the transition zone between subalpine and alpine habitats in Alaska with Betula nana, near Tsuga and Picea. Aug. Material examined. USA. ALASKA. Anchorage County, Girdwood, Crow Pass Road, with Betula nana, Tsuga, and Picea, 19 Aug 2018, C. Cripps CLC 3778 (MONT). 214 Observations. Russula vinosa was originally described by Lindblad in 1901. Our collection forms a strongly-supported clade with another collection identified as R. vinosa (AY061724, FIG. 3) by Miller and Buyck (2002). Our collections match the habitat and general morphology of this species; therefore, we elect to call our species R. aff. vinosa. 215 REFERENCES CITED 216 Adamčík S. 1998. Dva typické alpínske druhy – pavučinovec drobný Cortinarius pauperculus a plávka vysokohorská Russula nana. Spravodajca slovenských Mykológov 6:11–14. Adamčík S, Buyck B. 2014. Type studies in Russula subsection nigricantes from the Eastern United States. Cryptogamie, Mycologie 35(3):293–309. Adamčík S, Caboň M, Eberhardt U, Saba M, Hampe F, Slovak M, Kleine J, Marxmueller H, Jančovičová S, Pfister DH, Khalid AN. 2016a. A molecular analysis reveals hidden species diversity within the current concept of Russula maculata (Russulaceae, Basidiomycota). Phytotaxa 270(2):71–88. Adamčík S, Knudsen H. 2004. Red-capped species of Russula sect. Xerampelinae associated with dwarf scrub. Mycological Research 108(12):1463–1475. Adamčík S, Looney B, Caboň M, Jančovičová S, Adamčíková K, Avis PG, et al. 2019. The quest for a globally comprehensible Russula language. Fungal Diversity:1– 81. Adamčík S, Ripková S, Zaliberová M. 2006. Diversity of Russulaceae in the Vihorlatské vrchy Mts. (Slovakia). Czech Mycology 58(1–2):43–66. Adamčík S, Slovák M, Eberhardt U, Ronikier A, Jairus T, Hampe F, Verbeken A. 2016b. Molecular inference, multivariate morphometrics and ecological assessment are applied in concert to delimit species in the Russula clavipes complex. Mycologia 108(4):716–730. Ahl ED. 2016. A geographical analysis of alpine lichen in Rocky Mountain National Park. University of Colorado Master’s Thesis, Denver, Colorado. Ammirati JF, Laursen GA. 1982. Cortinarii in Alaskan Arctic tundra. In: Laursen GA, Ammirati JF, eds. Arctic and Alpine Mycology, The First International Symposium on Arcto-Alpine Mycology. University of Washington Press, Seattle. p. 282–315. Anderson JP. 1940. Notes on Alaskan rust fungi. Bulletin of the Torrey Botanical Club 67(5):413–416. Anisimov OA, Vaughan DG, Callaghan TV, Furgal C, Marchant H, Prowse TD, Vilhjálmsson H, Walsh JE. 2007. Polar regions (Arctic and Antarctic). In: Parry ML, Canziani OF, Palutikof JP, Van Der Linden PJ, Hanson CE, eds. Climate Change 2007: Impacts, Adaptation and Vulnerability. Contribution of Working Group II to the Fourth Assessment Report of the Intergovernmental Panel on Climate Change. p. 653–685. 217 Anthelme F, Lavergne S. 2018. Alpine and arctic plant communities: a worldwide perspective. Perspectives in Plant Ecology Evolution and Systematics 30(SI):1–5. Antibus RK. 2010. Temperature acclimation effects on growth, respiration and enzyme activities in an arctic and a temperate isolate of Cenococcum geophilum Fr. North American Fungi 5(5):187–204. Antibus RK, Croxdale JG, Miller Jr. OK, Linkins AE. 1981. Ectomycorrhizal fungi of Salix rotundifolia III. Resynthesized mycorrhizal complexes and their surface phosphatase activities. Canadian Journal of Botany 59(12):2458–2465. Antibus RK, Hobbie EA, Cripps CL. 2018. Sporocarp δ15N and use of inorganic and organic nitrogen in vitro differ among host-specific suilloid fungi associated with high elevation five-needle pines. Mycoscience 59(4):294–302. Antibus RK, Linkins AE. 1978. Ectomycorrhizal fungi of Salix rotundifolia Trautv. I. Impact of surface applied Prudhoe Bay crude oil on mycorrhizal structure and composition. Arctic:366–380. Arthur JC. 1911. Some Alaskan and Yukon rusts. The Plant World 14(10):233–236. Ballarà J. 1997. Nou estudi d’espècies fúngiques interessants dels estatges alpí I subalpí dels Pirineus Catalans. Revista Catalana micol. 20:1–24. Barge EG, Cripps CL. 2016. New reports, phylogenetic analysis, and a key to Lactarius Pers. in the Greater Yellowstone Ecosystem informed by molecular data. MycoKeys 15:1–58. Barge EG, Cripps CL, Osmundson TW. 2016. Systematics of the ectomycorrhizal genus Lactarius in the Rocky Mountain alpine zone. Mycologia 108(2):414–440. Barr ME. 1959. Northern Pyrenomycetes I. Canadian eastern Arctic. Contributions de I’Institut Botanique de I’Universite de Montréal 73:1–101. Bazzicalupo AL, Berbee M, Wood H, Voitk M, Voitk A. 2016. The Russula emetica complex in NL – preliminary report. Omphalina 7(7):3–7. Bazzicalupo AL, Buyck B, Saar I, Vauras J, Carmean D, Berbee ML. 2017. Troubles with mycorrhizal mushroom identification where morphological differentiation lags behind barcode sequence divergence. Taxon 66(4):791–810. Bazzicalupo AL, Whitton J, Berbee ML. 2019. Over the hills, but how far away? Estimates of mushroom geographic range extents. Journal of Biogeography 46(7):1547–1557. 218 Beardslee HC. 1918. The Russulas of North Carolina. Journal of the Elisha Mitchell Scientific Society 33:147–197. Beardslee HC. 1934. New and Interesting Fungi. Mycologia 26(3):253–260. Becker HJ, Eberhardt U, Vesterholt J. 2010. Hebeloma hiemale Bres. in Arctic/Alpine Habitats. North American Fungi 5:51–65. Bergemann SE, Miller SL, Garbelotto M. 2005. Microsatellite loci from Russula brevipes, a common ectomycorrhizal associate of several tree species in North America. Molecular Ecology Notes 5(3):472–474. Bergemann SE, Douhan GW, Garbelotto M, Miller SL. 2006. No evidence of population structure across three isolated subpopulations of Russula brevipes in an oak/pine woodland. New Phytologist 170(1):177–184. Berkeley MJ. 1839. XLII. Descriptions of Exotic Fungi in the collection of Sir W. J. Hooker, from Memoirs and Notes of J.F. Klotzsch, with Additions and Corrections. Journal of Natural History 3(19):375–401. Berkeley MJ. 1878. Enumeration of the fungi collected during the Arctic Expedition, 1875-76. Journal of the Linnean Society, Botany 17:13–17. Bigelow HE. 1959. Notes of fungi from northern Canada IV. Tricholomataceae. Canadian Journal of Botany 37(5):769–779. Bigelow HE. 1970. Omphalina in North America. Mycologia 62(1):1–32. Billings WD. 1973. Arctic and alpine vegetation: similarities, differences, and susceptibility to disturbance. Bioscience 23(12):697–704. Billings WD. 1974. Adaptations and origins of alpine plants. Arctic and Alpine Research 6(2):129–142. Billings WD. 1988. Alpine vegetation. In: Barbour MG, Billings WD, eds. North American Terrestrial Vegetation. Cambridge University Press, New York. p. 392– 420. Bills GF. 1984. Southern Appalachian russulas. II. Mycotaxon 21:491–517. Bills GF. 1985. Southern Appalachian russulas. III. The identity of Russula eccentrica and R. morgani (Russulaceae). Brittonia 37:360–365. 219 Bills GF. 1989. Southern Appalachian Russulas. IV. Mycologia 81(1):57–65. Bills GF, Miller Jr OK. 1984. Southern Appalachian Russulas. I. Mycologia 76(60):975– 1002. Binder M, Hibbett DS. 2002. Higher level phylogenetic relationships of homobasidiomycetes (mushroom-forming fungi) inferred from four rDNA regions. Molecular Phylogenetics and Evolution 22(1):76–90. Binder M, Hibbett DS, Larsson K-H, Larsson E, Langer E, Langer G. 2005. The phylogenetic distribution of resupinate forms across the major clades of mushroom-forming fungi (Homobasidiomycetes). Systematics and Biodiversity 3(2):113–157. Bjorbækmo MFM, Carlsen T, Brysting A, Vrålstad T, Høiland K, Ugland KI, Geml J, Schumacher T, Kauserud H, 2010. High diversity of root associated fungi in both alpine and arctic Dryas octopetala. BMC Plant Biology 10(1):e244. Bliss LC. 1962. Adaptations of arctic and alpine plants to environmental conditions. Arctic 15(2):117–144. Bliss LC. 1988. Arctic tundra and polar desert biome. In: Barbour MG, Billings WD eds. North American Terrestrial Vegetation. Cambridge University Press, New York. p. 1–32. Boa ER. 2004. Wild edible fungi: a global overview of their use and importance to people. Non-wood Forest Products 17. Economic Botany 60(1):99–101. Boertmann D, Knudsen H. 2006. Arctic and Alpine Mycology 6. Meddelelser om Grøenland Bioscience 56: 161 p. Bon M. 1972. Contribution a l’etude des Viridantinae Melz. Zv.: Russula subrubens (Lge) n. c. Documents Mycologiques 2(5):33–36. Bon M. 1985. Stage Mycologie Alpine Lanslebourg (Savoie) du 1 au 3 septembre 1984. Bulletin Trimestriel de la Fédération Mycologique Dauphiné-Savoie 96:19– 25. Bon M. 1987. Quelques recoltes mycologiques de la zone alpine au 7ème convegno di micologia. Fiera di Primiero (Italie). Micologia Italiana 17(3):267–270. Bon M. 1988. Clé monographique des russules d’Europe. Documents Mycologiques 18(70–71):1–120. 220 Bon M. 1991. Inventaires des espéces récoltees au stage de mycologie alpine. Bulletin de la Féderation Mycologique du Dauphiné-Savoie 122:25–28. Bon M. 1993. Russules Alpine Nouvelles Ou Intéressantes. Bulletin de la Féderation Mycologique du Dauphiné-Savoie 128:20–24. Bon M. 2000. Essai de clé de détermination des Russules alpines. Bulletin de la Féderation Mycologique du Dauphiné-Savoie 158:9–17. Bon M, Ballarà J. 1996. Aportació a L’estudi de la microflora alpine dels Pirineus (2a part). Revista Catalana de Mycologia 19:139–153. Bon M, Cheype JL. 1987. Mycologie alpine au col du Joly Haute-Savoie; altitude 2000 m. Bulletin Trimestriel de la Fédération Mycologique Dauphiné-Savoie 106:22– 27. Bon M, Noguera JB. 1995. Aportació a l'estudi de la micoflora alpina dels Pirineus (1ª part). Revista Catalana de Micologia 18:39–50. Borgen T. 1993. Svampe 1 Grøenland, Atuakkiorfik, printed in Denmark. Boudier: 112 p. Borgen T. 2006. Distribution of selected basidiomycetes in oceanic dwarf-scrub heaths in the Paamiut area, low arctic South Greenland. In: Boertmann D, Knudsen H, eds. Arctic and Alpine Mycology 6, Grønland Bioscience 56. p. 25–36. Borgen T, Elborne SA, Knudsen H. 2006. A checklist of the Greenland basidiomycetes. In: Boertmann D, Knudsen H, eds. Arctic and Alpine Mycology 6, Grønland Bioscience 56. p. 37–59. Bouckaert R, Vaughan TG, Barido-Sottani J, Duchêne S, Fourment M, Gavryushkina A, et al. 2019. BEAST 2.5: An advanced software platform for Bayesian evolutionary analysis. PLoS computational biology 15(4):e1006650. Bowerman CA, Groves JW. 1962. Notes on fungi from northern Canada V. Gasteromycetes. Canadian Journal of Botany 40(1):239–254. Bresinsky A. 1987. Bemerkenswerte Grosspilzfunde in der Bundersrepublik Deutschland. Zeitschrift für Mykologie 53(2):289–302. Bresinsky A, Kreisel H, Beisenherz M, Eger A. 2000. Mykologisches aus dem Werdenfelser Land: Bovista bovistoides, Lactarius salicis-reticulata. Zeitschrift für Mykologie, 66(2): 123 p. 221 Bresinsky A, Stangl J, Einhellinger A. 1980. Beiträge zur Revision M. Britzelmayr's "Hymenomyceten aus Südbayern" 14. Die Gattung Russula unter besonderer Berücksichtigung ihrer Arten in der Umgebung von Augsburg. Zeitschrift für Mykologie 46:131–156. Brown R. 1823. A list of plants collected in Melville Island in the year 1820; by the officers of the voyage of discovery under the orders of Captain Parry. Chloris Melvilliana. Printed by W. Clowes, Northumberland-Court, Strand. London: 49 p. Brunner I. 1989. Two New Species of Russula (Stirps Atropurpurea) Associated with Alnus crispa in Alaska. Mycologia 81(5):667–676. Budd GE, Jensen S. 2000. A critical reappraisal of the fossil record of the bilaterian phyla. Biological Reviews 75(2):253–295. Bueno de Mesquita CP, Martinez del Río CM, Suding KN, Schmidt SK. 2018a. Rapid temporal changes in root colonization by arbuscular mycorrhizal fungi and fine root endophytes, not dark septate endophytes, track plant activity and environment in an alpine ecosystem. Mycorrhiza 28(8):717–726. Bueno de Mesquita CP, Sartwell SA, Ordemann EV, Porazinska DL, Farrer EC, King AJ, Spasojevic MJ, Smith JG, Suding KN, Schmidt SK. 2018b. Patterns of root colonization by arbuscular mycorrhizal fungi and dark septate endophytes across a mostly-unvegetated, high-elevation landscape. Fungal Ecology 36:63–74. Bunnell FL, Miller Jr OK, Flanagan PW, Benoit RE. 1980. The Microflora: Composition, biomass, and environmental relations. In: Brown J, Miller PhC, Tieszen LL, Bunnell FL, eds. An Arctic ecosystem. The coastal tundra at Barrow, Alaska. US/IBP Synthesis Series 12. p 255–290. Burlingham GS. 1913. The Lactarieae of the Pacific coast. Mycologia 5(6):305–311. Burlingham GS. 1915. Russula Pers. North American Flora 9(4):201–236. Burlingham GS. 1921. Some New Species of Russula. Mycologia 13(3):129–134. Burlingham GS. 1924. Notes on species of Russula. Mycologia 16(1):16–23. Burlingham GS. 1936. New or noteworthy species of Russula and Lactaria. Mycologia 28(3):253–257. Burlingham GS. 1939. Two new species of Russula together with the spore ornamentation of some of our American Russulas. Mycologia 31(4):490–498. 222 Burlingham GS. 1944. Studies in North American Russulae. Mycologia 36(1):104–120. Buyck B. 1989. Valeur taxonomique du bleu de crésyl pour le genre Russula. Bulletin trimestriel de la Société mycologique de France 105:1–6. Buyck B. 1991. The study of microscopic features in Russula 1. spores and basidia. Russulales newsletter 1:8–26. Buyck B. 1994. Russula II (Russulaceae). In: Rammeloo J, Heinemann P, eds. Flore illustrée des Champignons d’Afrique Centrale. Ministère de l’Agriculture, Jardin Botanique National de Belgique, Meise 16. p 411–542. Buyck B. 2007. A new initiative towards the study of Russula in the eastern USA. Pagine Di Micologia 27:81–86. Buyck B. 2019. Russulales News, The Russulales News Team, http://www2.muse.it/russulales-news. – Check with mycologia website format Buyck B, Adamčík S. 2011. Type studies of Russula species described by W.A. Murrill, 1. R. roseiisabellina, R. sericella, and R. obscuriformis. Mycotaxon 115:131–144. Buyck B, Adamčík S. 2013. Type studies in Russula subsection Lactarioideae from North America and a tentative key to North American Species. Cryptogamie, Mycologie 34(3):259–279. Buyck B, Hofstetter V. 2011. The contribution of tef-1 sequences for species delimitation in the Cantharellus cibarius complex in the southeastern USA. Fungal Diversity 49(1):35–46. Buyck B, Hofstetter V, Berbeken A, Walleyn R. 2008. Walking the thin line between Russula and Lactarius: the dilemma of Russula subsect. Ochricompactae. Fungal Diversity 28:15–40. Buyck B, Jančovičová S, Adamčík S. 2015. The Study of Russula in the Western United States. Crytogamie, Mycologie 36(2):193–211. Buyck B, Thoen D, Watling R. 1996. Ectomycorrhizal fungi of the Guinea–Congo region. Proceedings of the Royal Society of Edinburgh, Section B: Biological Sciences 104:313–333. Buyck B, Wang XH, Adamcikova K, Cabon M, Jancovicova S, Hofstetter V, Adamcik S. 2020. One step closer to unravelling the origin of Russula: subgenus Glutinosae subg. nov. Mycosphere 11(1):285–305. 223 Buyck B, Zoller S, Hofstetter V. 2018. Walking the thin line… ten years later: the dilemma of above- versus below-ground features to support phylogenies in the Russulaceae (Basidiomycota). Fungal Diversity 89:267–292. Caboň M, Eberhardt U, Looney B, Hampe F, Kolařík M, Jančovičová S, Verbeken A, Adamčík S. 2017. New insights in Russula subsect. Rubrinae: phylogeny and the quest for synapomorphic characters. Mycological progress 16(9):877–892. Caboň M, Li GJ, Saba M, Kolařík M, Jančovičová S, Khalid AN, Moreau PA, Wen HA, Pfister DH, Adamčík S. 2019. Phylogenetic study documents different speciation mechanisms within the Russula globispora lineage in boreal and arctic environments of the Northern Hemisphere. IMA Fungus 1(1): 5. – I don’t know what to do with this 5??? Cash EK. 1953. A check list of Alaskan fungi. The Plant Disease Reporter. Supplement 219:1–70. Callaghan TV, Björn LO, Chapin III FS, Chernov Y, Christensen TR, Huntley B, Ims R, Johansson M, Riedlinger DJ, Jonasson S, Matveyeva N. 2005. Arctic tundra and polar desert ecosystems. Arctic climate impact assessment 1:243–352. Chapin III FS, Körner C. 1995. Patterns, causes, changes, and consequences of biodiversity in arctic and alpine ecosystems. In: Chapin III FS, Körner CH, eds. Arctic and Alpine Biodiversity: Patterns, Causes and Ecosystems Consequences. Ecological Studies 113. Springer-Verlag, New York. p. 313–320. Chapin III FS, Shaver GR, Giblin AE, Nadelhoffer KJ, Laundre JA. 1995. Responses of Arctic tundra to experimental and observed changes in climate. Ecology 76:694– 711. Christiansen MP. 1941. Studies in the Larger Fungi of Iceland. The Botany of Iceland 3(2):187–227. Clericuzio M, Fugui H, Fusheng P, Zijie P, Sterner O. 1998. The sesquiterpenoid contents of fruit bodies of Russula delica. Acta chemica scandinavica 52(11):1333–1337. Clericuzio M, Sterner O. 1997. Conversion of velutinal esters in the fruit bodies of Russula cuprea. Phytochemistry 45(8):1569–1572. Comiso JC, Hall DK. 2014. Climate trends in the Arctic as observed from space. Wiley Interdisciplinary Reviews: Climate Change 5(3):389–409. 224 Cooke WB, Fournelle HT. 1960. Some soil fungi from an Alaskan tundra area. Arctic 13(4):266–270. Corbridge JN, Weber WA. 1998. Rocky Mountain lichen primer. University Press of Colorado, Louisville, CO: 56 p. Corriol G. 2008. Checklist of Pyrenean alpine-stage macrofungi. Sommerfeltia 31:29–99. Courty PE, Franc A, Pierrat JC, Garbaye J. 2008. Temporal changes in the ectomycorrhizal community in two soil horizons of a temperate oak forest. Applied and Environmental Microbiology 74(18):5792–580. Cripps CL. 2003. Interesting distributions of ectomycorrhizal alpine fungi along the Rocky Mountain cordillera. Mycological Society of America, Pacific Grove, CA, July 27–31. Cripps CL. 2010. Orson K. Miller Jr., 1930–2006. Mycologia 102(5):1216–1220. Cripps CL, Ammirati J. 2010. Eighth International Symposium on Arctic-Alpine Mycology (ISAM 8), Beartooth Plateau, Rocky Mountains, USA 2008. North American Fungi 5: 1–8. – why does this say 1–8???? Cripps CL, Barge E. 2013. Notes on the genus Lactarius from the Rocky Mountain alpine zone in regard to Finnish arctic-alpine species. Karstenia 53(1-2):29–37. Cripps CL, Eddington LH. 2005. Distribution of mycorrhizal types among alpine vascular plant families on the Beartooth Plateau, Rocky Mountains, USA, in reference to large-scale patterns in arctic-alpine habitats. Arctic, Antarctic, and Alpine Research 37(2):177–188. Cripps CL, Evenson V, Kuo M. 2016. The essential guide to Rocky Mountain mushrooms by habitat. University of Illinois Press, Urbana, Chicago, and Springfield. p. Cripps CL, Eberhardt U, Schütz N, Beker HJ, Evenson VS, Horak E. 2019a. The genus Hebeloma in the Rocky Mountain Alpine Zone. MycoKeys 46:1–54. Cripps CL, Horak E. 1999. Alpine Mycota (Agaricales) Rocky Mountain Tundra, USA: a preliminary report. International Botanical Congress, St. Louis, MO. Aug. 6, 1999. Cripps CL, Horak E. 2002. Alpine species of Lactarius in the Rocky Mountains. Mycological Society of America, Corvallis, OR. June 24–26, 2002. 225 Cripps CL, Horak E. 2005. Amanita in the Rocky Mountain Alpine Zone: Where mycorrhizal mushrooms tower over miniature forests. Joint meeting of Japanese Mycological Society and Mycological Society of America, Hilo, HI. July 30-Aug 5, 2005. Cripps CL, Horak E. 2006. Arrhenia auriscalpium in arctic-alpine habitats: world distribution, ecology, new reports from the southern Rocky Mountains, USA. In: Boertmann D, Knudsen H, eds. Arctic and Alpine Mycology 6. Meddelelser om Grøenland Bioscience 56. p 17–24. Cripps CL, Horak E. 2007. Alpine agarics with Dryas octopetala (Rosaceae) in arctic- alpine habitats of the Rocky Mountains (USA), Mycological Society of American meeting, Baton Rouge, LA August 6–9, 2007. Cripps C, Horak E. 2008. Checklist and Ecology of the Agaricales, Russulales and Boletales in the alpine zone of the Rocky Mountains (Colorado, Montana, Wyoming) at 3000-4000 m asl. Sommerfeltia 31:101–123. Cripps CL, Horak E. 2010. Amanita in the Rocky Mountain alpine zone, USA: New records for A. nivalis and A. groenlandica. North American Fungi 5:9–21. Cripps CL, Horak E, Mohatt K. 2008. Ectomycorrhizal fungi at alpine treeline in the Rocky Mountains: Baseline data and a review in the context of climate change. MTNCLIME, June 9–12. Consortium for Integrated Climate Change Research in Western Mountains. Cripps CL, Larsson E, Horak E. 2010. Subgenus Mallocybe (Inocybe) in the Rocky Mountain alpine zone with molecular reference to European arctic-alpine material. North American Fungi 5:97–126. Cripps CL, Larsson E, Vauras J. 2019b. Nodulose-spored Inocybe from Rocky Mountain alpine zone molecularly linked to European and type specimens Mycologia. Dahlberg A, Bültmann H. 2013. Fungi. Chapter 10. In: Meltofte H, ed. Arctic Biodiversity Assessment. Status and Trends in Arctic Biodiversity. Conservation of Arctic Flora and Fauna (CAFF). Narayana Press, Denmark. p 354–371. Daniewski W, Gumulka M, Ptaszynska K, Skibicki P, Bloszyk E, Drozdz B, Stromberg S, Norin T, Holub M. 1993. Antifeedant activity of some sesquiterpenoids of the genus Lactarius (Agaricales: Russulaceae). European Journal of Entomology 90(1):65–70. 226 Daniewski WM, Gumułka M, Przesmycka D, Ptaszyńska K, Błoszyk E, Drożdż B. 1995. Sesquiterpenes of Lactarius origin, antifeedant structure-activity relationships. Phytochemistry 38(5):1161–1168. Das K, Ghosh A, Chakraborty D, Li J, Qiu L, Baghela A, Halama M, Hembrom ME, Mehmood T, Parihar A, Pencakowski B, Bielecka M, Reczynska K, Sasiela D, Singh U, Song Y, Swierkosz K, Szczesniak K, Uniyal P, Zhang J, Buyck B. 2017. Fungal biodiversity profiles 31–40. Cryptogam Mycologie 38(3):1–56. Dearness J. 1923. Report of the Canadian Arctic Expedition 1913-1918. Volume IV: Botany. Part C: Fungi. Ottawa:1–24. Dearness J. 1928. Report on fleshy fungi collected in August 1926. In: Soper JD, ed. A faunal investigation of southern Baffin Land. National Museum of Canada Bulletin No. 53 Biological Series. p 120–123. De Crop E, Nuytinck J, Van de Putte K, Wisitrassameewong K, Hackel J, Stubbe D, Hyde KD, Roy M, Halling RE, Moreau PA, Eberhardt U. 2017. A multi-gene phylogeny of Lactifluus (Basidiomycota, Russulales) translated into a new infrageneric classification of the genus. Persoonia: Molecular Phylogeny and Evolution of Fungi 38:58–80. De Groot WJ, Thomas PA, Wein RW. 1997. Betula nana L. and Betula glandulosa Michx. Journal of Ecology 85(2):241–264. Dennis RWG. 1970. Fungus flora of Venezuela and adjacent countries. Kew Bulletin Additional Series 3.1–531. Deslippe JR, Hartmann M, Mohn WW, Simard SW. 2011. Long-term experimental manipulation of climate alters the ectomycorrhizal community of Betula nana in Arctic tundra. Global Change Biology 17:1625–1636. Deslippe JR, Hartmann M, Simard SW, Mohn WW. 2012. Long-term warming alters the composition of Arctic soil microbial communities. FEMS microbiology ecology 82(2):303–315. Deslippe JR, Simard SW. 2011. Below‐ground carbon transfer among Betula nana may increase with warming in Arctic tundra. New Phytologist 192(3):689–698. Desmond R. 1995. Kew: the history of the Royal Botanic Gardens. The Harvill Press, London:150–238. Donk MA. 1964. A conspectus of the families of Aphyllophorales. Persoonia 3(2):199– 324. 227 Donk MA. 1971. Progress in the study of the classification of the higher Basidiomycetes. In: Petersen RH, ed. Progress in the study of the classification of the higher basidiomycetes. The University of Tennessee Press, Knoxville. p 3–25. Durand EJ. 1908. The Geoglossaceae of North America. Annales Mycologici 6:387–477. Earle FS. 1902. Mycological studies. I. Bulletin of the New York Botanical Garden 2(7):331–350. Elborne SA, Knudsen H. 1990. Larger fungi associated with Betula pubescens in Greenland. Meddelelser om Grønland Biosciences 33:77–80. Elliot TF, Trappe JM. 2018. A worldwide nomenclature revision of sequestrate Russula species. Fungal Systematics and Evolution 1(1):229–242. Estey RH. 1994. Essays on the early history of plant pathology and mycology in Canada. McGill-Queen's University Press, Montreal, QC. Eversman S. 1995. Lichens of alpine meadows on the Beartooth Plateau, Montana and Wyoming, USA. Arctic and Alpine Research 27(4):400–406. Fagre DB. 2009. Introduction: understanding the importance of alpine treeline ecotones in mountain ecosystems. In: Butler CR, Malanson GP, Walsh SJ, Fagre DB, eds. The changing alpine treeline. Developments in Earth Surface Processes 12. Amsterdam, Elsevier. p. 35–61. Farr DF, Miller Jr OK, 1972. Notes on arctic and subarctic Basidiomycetes. Virginia Journal of Science 23: 120 p. Fatto RM. 1999. Three new species of Russula. Mycotaxon 70():167–175. Fatto RM. 2000. Several Russulas of the Chiricahua Mountains. Mycotaxon 75:265–272. Fatto RM. 2002. Some Russulas of the subsection Urentinae. Mycotaxon 84:229–244. Favre-Bonvin J, Bernillon J. 1982. Structure du stearyl-velutinal, sequiterpenoide naturel de Lactarius velutinus bert1. Tetrahedron Letters 23(18):1907–1908. Favre J. 1955. Les champignons supérieurs de la zone alpine du Parc National Suisse. Resultats des recherches scientifiques enterprises au Parc National Suisse. Ergebnisse der wissenschaftlichen Untersuchungen des schweizerischen National Parks 5: 212 p. 228 Fellner R, Landa J. 1991. Arctic and alpine fungi in Czechoslovakia. Czech Mycology 45: 35 p. Fellner R, Landa J. 1993a. Some species of Cortinariaceae and Russulaceae in the alpine belt of the Belar Tatras – I. In: Petřini O, Laursen GA, eds. Arctic and alpine mycology 3–4. J Cramer, Stuttgart, Berlin. p 33–37. Fellner R, Landa J. 1993b. Some species of Cortinariaceae and Russulaceae in the alpine belt of the Belaer Tatras II. Czech Mycology 47:45–55. Fischer MW, Stolze-Rybczynski JL, Cui Y, Money NP. 2010. How far and how fast can mushroom spores fly? Physical limits on ballistospore size and discharge distance in the Basidiomycota. Fungal biology 114(8):669–75. Flanagan PW, Scarborough AM. 1973. Laboratory and field studies of decomposition organisms and processes in arctic tundra. U.S. Tundra Biome Data Report 73–28. Tundra Biome Center, University of Alaska, Fairbanks. Flanagan PW, Scarborough AM. 1974. Physiological groups of decomposer fungi on tundra plant remains. Soil Organisms and Decomposition. In: Holding AJ, Heal OW, Maclean Jr SF, Flanagan PW. eds. Tundra, IBP Tundra Biome, TBSC Stock. p 159–181. Fogel R. 1975. Insect mycophagy: a preliminary bibliography. Gen. Tech. Rep. PNW- GTR-036. Portland, OR: US Department of Agriculture, Forest Service, Pacific Northwest Forest and Range Experiment Station. 21: 36 p. Fogel R, Trappe JM. 1978. Fungus consumption (mycophagy) by small animals. Northwest Science 52: 31 p. Formica A, Farrer EC, Ashton IW, Suding KN. 2014. Shrub expansion over the past 62 years in Rocky Mountain alpine tundra: possible causes and consequences. Arctic, Antarctic, and Alpine Research 46(3):616–631. Fries EM. 1821. Systema Mycologicum, sistens fungorum ordines, genera et species huc usque cognitas. Vol. I. Berling, Lund. Fries EM. 1838. Epicrisis Systematis Mycologici, seu synopsis Hymenomycetum. Typographia Academica, Uppsala. Fries EM. 1874. Hymenomycetes Europaei. Uppsala: Ed Berlingiana: 755 p. 229 Gardes M, Bruns TD. 1993. ITS Primers with enhanced specificity for Basidiomycetes: application to the identification of mycorrhizae and rusts. Molecular Ecology 2:113–118. Gardes M, Dahlberg A. 1996. Mycorrhizal diversity in Arctic and alpine tundra: an open question. New Phytologist 133(1):147–157. Geml J, Gravendeel B, van der Gaag KJ, Neilen M, Lammers Y, Raes N, Semenova TA, de Knijff P, Noordeloos ME. 2014. The contribution of DNA metabarcoding to fungal conservation: diversity assessment, habitat partitioning and mapping red- listed fungi in protected coastal Salix repens communities in the Netherlands. PLoS One 9(6): e99852. Geml J, Laursen GA, Herriott IC, McFarland JM, Booth MG, Lennon N, Chad Nusbaum H, Lee Taylor D. 2010. Phylogenetic and ecological analyses of soil and sporocarp DNA sequences reveal high diversity and strong habitat partitioning in the boreal ectomycorrhizal genus Russula (Russulales; Basidiomycota). New Phytologist, 187(2):494–507. Geml J, Laursen GA, Timling I, McFarland JM, Booth MG, Lennon N, Nusbaum C, Taylor DL. 2009. Molecular phylogenetic biodiversity assessment of arctic and boreal ectomycorrhizal Lactarius Pers. (Russulales; Basidiomycota) in Alaska, based on soil and sporocarp DNA. Molecular Ecology 18(10):2213–2227. Geml J, Morgado LN, Semenova TA, Welker JM, Walker MD, Smets E. 2015. Long- term warming alters richness and composition of taxonomic and functional groups of arctic fungi. FEMS Microbiology Ecology 91(8): fiv095. Geml J, Semenova TA, Morgado LN, Welker JM. 2016. Changes in composition and abundance of functional groups of arctic fungi in response to long-term summer warming. Biology letters 12(11): e20160503. Geml J, Taylor DL. 2013. Biodiversity and molecular ecology of Russula and Lactarius in Alaska based on soil and sporocarp DNA sequences. Scripta Botanica Belgica (51):132–145. Geml J, Timling I, Robinson CH, Lennon N, Nusbaum HC, Brochmann C, Noordeloos ME, Taylor, DL. 2012. An arctic community of symbiotic fungi assembled by long-distance dispersers: phylogenetic diversity of ectomycorrhizal basidiomycetes in Svalbard based on soil and sporocarp DNA. Journal of Biogeography 39(1):74–88. Gillman LS. Miller Jr OK. 1977. A study of the boreal, alpine, and arctic species of Melanoleuca. Mycologia 69(5):927–951. 230 Gorbunova IA. 2014. Biota of Agaricoid and Gasteroid Basidiomycetes of Dryad Tundras of the Altai-Sayan Mountain Area (Southern Siberia). Contemporary Problems of Ecology 7(1):39–44. Graae BJ, Vandvik V, Armbruster WS, Eiserhardt WL, Svenning JC, Hylander K, Ehrlén J, Speed JD, Klanderud K, Bråthen KA, Milbau A. 2018. Stay or go–how topographic complexity influences alpine plant population and community responses to climate change. Perspectives in Plant Ecology, Evolution and Systematics 30:41–50. Grabherr G, Gottfried M, Gruber A, Pauli H. 1995. Patterns and Current Changes in Alpine Plant Diversity. In: Chapin III FS, Körner CH, eds. Arctic and Alpine Biodiversity: Patterns, Causes and Ecosystems Consequences. Ecological Studies 113. Springer-Verlag, New York. p 167–182. Graf F. 1994. Ecology and sociology of macromycetes in snow-beds with Salix herbacea L. in the alpine valley of Radönt (Grisons, Switzerland). Dissertationes Botanicae 235: 242 p. Graf F, Brunner I. 1995. Alpine dwarf willow and its ectomycorrhizal partners: A system for alpine ski slope renaturation. Proceedings High Altitude Revegetation Workshop No. 11, Colorado State University, Fort Collins, March 16–19, 1994. Fort Collins: Colorado Water Resources Institute, Colorado State University:214– 223. Gray SF. 1821. A natural arrangement of British plants. Vol. I. London. Groves JW, Elliott ME. 1971. Notes on fungi from northern Canada VI. Additional records of Discomycetes. Reports from the Kevo Subarctic Research Station 8:22–30. Groves JW, Hoare SC. 1954. Notes on fungi from northern Canada I. Hypocreales and discomycetes. Canadian Field-Naturalist 68:1–8. Groves JW, Thomson SC. 1955. Notes on fungi from northern Canada II. Boletaceae. Canadian Field-Naturalist 69:44–51. Groves JW, Thomson SC, Pantidou M. 1958. Notes on fungi from northern Canada III. Amanitaceae, Hygrophoraceae, Rhodophyllaceae, and Paxillaceae. Canadian Field-Naturalist 72:133–138. 231 Grund DW. 1965. A survey of the genus Russula occurring in Washington State, Doctoral dissertation. Department of Botany, University of Washington, Seattle: 396 p. Grund DW. 1979. New and interesting Russula Pers. ex S.F. Gray occurring in Washington state. Mycotaxon 9(1):93–113. Gulden G. 1983. Studies in Lepista (Fr.) W.G. Smith. Section Lepista (Basidiomycotina, Agaricales). Sydowia: Annales mycologici 36:59–73. Gulden G. 2005. A preliminary Guide to the Macromycetes in the Finse area, Hardangervidda, Norway. Prepared for ISAM VII: 81 p. Gulden G, Høiland K. 2008. ISAM VII at Finse, Norway, 2005. Sommerfeltia 31:7–16. Gulden G, Jenssen KM, Stordal J. 1985. Arctic and alpine fungi (Vol. 1). Lubrecht & Cramer Ltd. Soppkonsulenten. Oslo, Norway. Gulden G, Lange M. 1971. Studies in the macromycete flora of Jotunheimen, the central mountain massif of South Norway. Norwegian Journal of Botany 18:1–46. Gulden G, Torkelsen AE. 1996. Fungi I. Basidiomycota: Agaricales, Gasteromycetales, Aphyllophorales, Exobasidiales and Tremellales. Skrifter-Norsk Polarinstitutt 198:173–206. Guo W. 1992. Resources of wild edible fungi in Tibet, China. Zhongguo Shiyonjun 11:33–34. Guo-Jie L, Rui-Lin Z, Chu-Long Z, Fu-cheng L. 2019. A preliminary DNA barcode selection for the genus Russula (Russulales, Basidiomycota). Mycology 10(2):61– 74. Gyosheva MM, Dimitrova G. 2011. New records of larger fungi established in habitats of glacial relict plants in Bulgaria. Phytologia Balcanica 17(2):165–167. Hadley KS. 1987. Vascular alpine plant distributions within the central and southern Rocky Mountains, USA. Arctic and Alpine research 19(3):242–251. Hallgrimsson H. 1998. Checklist of Icelandic Fungi V: Agarics. Natturufraedistornun Islands. Halling RE. 2001. Ectomycorrhizae: co-evolution, significance, and biogeography. Annals of the Missouri Botanical Garden 88 (1):5–13. 232 Hanson JR. 2008. The Chemistry of fungi. Cambridge, U.K. Royal Society of Chemistry. Hansen L, Knudsen H. 1992. Nordic Macromycetes. Vol. 2. Nordsvamp, Copenhagen. Haselwandter K, Read DJ. 1980. Fungal associations of roots of dominant and sub- dominant plants in high-alpine vegetation systems with special reference to mycorrhiza. Oecologia 45(1):57–62. Heikkila H, Kallio P. 1966. On the problem of subarctic basidiolichens I. Reports from the Kevo Subarctic Research 3:48–74. Heikkila H, Kallio P. 1969. On the problem of subarctic basidiolichens II. Reports from the Kevo Subarctic Research 4:90–97. Heilmann-Clausen J, Verbeken A, Vesterholt J. 1998. The Genus Lactarius. Danish Mycological Society, Denmark. He MQ, Zhao RL, Hyde KD, Begerow D, Kemler M, Yurkov A, McKenzie EH, Raspé O, Kakishima M, Sánchez-Ramírez S, et al. 2019. Notes, outline and divergence times of Basidiomycota. Fungal Diversity:1–263. Henkel TW, Aime MC, Miller SL. 2000. Systematics of pleurotoid Russulaceae from Guyana and Japan, with notes on their ectomycorrhizal status. Mycologia 92(6):1119–1132. Hibbett DS, Donoghue MJ. 1995. Progress toward a phylogenetic classification of the Polyporaceae through parsimony analysis of mitochondrial ribosomal DNA sequences. Canadian Journal of Botany 73(S1):853–861. Hibbett DS, Pine EM, Langer E, Langer G, Donoghue MJ. 1997. Evolution of gilled mushrooms and puffballs inferred from ribosomal DNA sequences. Proceedings of the National Academy of Sciences, USA 94:12002–12006. Hillebrand H. 2004. On the generality of the latitudinal diversity gradient. The American Naturalist 163:192–211. Hintikka V, Niemi K. 1999. Aseptic culture of slowly-growing mycorrhizal Russula and Cortinarius species. Karstenia 39:39–41. Hobbie JE, Hobbie EA. 2006. 15N in symbiotic fungi and plants estimates nitrogen and carbon flux rates in arctic tundra. Ecology 87(4):816–822. Hoeksema JD. 2010. Ongoing coevolution in mycorrhizal interactions. New Phytologist 187:286–300. 233 Hofstetter V, Buyck B, Eyssartier G, Schnee S, Gindro K. 2019. The unbearable lightness of sequenced-based identification. Fungal Diversity 96(1):243–84. Homola RL, Shaffer RL. 1975. A new Russula of the subsection Nigricantes from Northeastern North America. Mycologia 67(2):428–234. Hongsanan S, Hyde KD, Bahkali AH, Camporesi E, Chomnunti P, Ekanayaka H, et al. 2015. Fungal biodiversity profiles 11–20. Cryptogamie, mycologie 36(3):355– 381. Hooker JD. 1860. An Account of the Plants collected by Dr. Walker in Greenland and Arctic America during the Expedition of Sir Francis M'Clintock, RN, in the Yacht ‘Fox.’. Botanical Journal of the Linnean Society 5(18):79–89. Horak E. 1960. Die Pilzvegetation im Gletschervorfeld (2290–2350 m) des Rotmoosferners in den Ötztaler Alpen. Nova Hedwigia 2(4):487–507. Horak E, Miller Jr OK. 1992. Phaeogalera and Galerina in arctic-subarctic Alaska (USA) and the Yukon Territory (Canada). Canadian journal of Botany 70(2):414– 433. Horak E, Moser MM. 2006. Agrocybe praemagna, a new species. Meddelelser om Grønland Bioscience 56:133–138. Horton TR, Bruns TD. 2001. The molecular revolution in ectomycorrhizal ecology: peeking into the black-box. Molecular Ecology 10(8):1855–1871. Hoshino T, Tuno N, Degawa Y, Kasuya T, Yajima Y, Kawahara E, Nose I. 2018. The 10th International Symposium on Arctic and Alpine Mycology. Mycoscience 30:1–2. Hu L, Zeng L. 1992. Investigation on wild edible mushroom resources in Wanxian Country, Sichuan Province. Zhongguo Shiyongjun 11:35–37. Huhtinen S. 1982. Ascomycetes from central and northern Labrador. Karstenia 22:1–8. Huhtinen S. 1985. Mycoflora of Poste-de-la-Baleine, northern Quebec: Ascomycetes. Le Naturaliste Candien 112(4):473–524. Huijsman HSC. 1955. Observations on Agarics. Fungus 25:18–43. Hultén E. 1968. Flora of Alaska and Neighboring Territories. Stanford: Stanford University Press, California: 1008 p. 234 Hutchison LJ, Summerbell RC, Malloch DW. 1988. Additions to the mycota of North America and Quebec: arctic and boreal species from Schefferville, northern Quebec. Le Naturaliste Candien 115:39–56. Hyde KD, Hongsanan S, Jeewon R. et al. 2016. Fungal diversity notes 367–490: taxonomic and phylogenetic contributions to fungal taxa. Fungal Diversity 80:1– 270. Imshaug HA. 1957. Alpine lichens of western United States and adjacent Canada I. The macrolichens. The Bryologist 60(3):177–272. IPCC, editor. 2014. Climate Change 2014: Synthesis Report. Contribution of Working Groups I, II and III to the Fifth Assessment Report of the Intergovernmental Panel on Climate Change. Geneva, Switzerland. Irlet B, Rieder K. 1985. Cadmium und Blei in Pilzen aus der alpinen Stufe der schweizer Alpen. Mycologia Helvetica 1:393–399. Jalink LM. 2010. Additional notes on the Lycoperdaceae of the Beartooth Plateau. North American Fungi 5:173–179. James TY, Kauff F, Schoch CL, Matheny PB, Hofstetter V, Cox CJ, Celio G, Gueidan C, Fraker E, Miadlikowska J, Lumbsch HT. 2006. Reconstructing the early evolution of Fungi using a six-gene phylogeny. Nature 443(7113):818–822. Jamoni PG, Bon M. 1993. Note di Mycologia alpina: reperti rari e nuovi della zona alpina del Massiccio del Monte Rosa e Dintorni (3a parte). Rivista Micologia 36:3–6. Jamoni PG. 1995. Russulaceae della zona alpine. Proposta di chiavi di determinazione per le specie crescent nella zona alpine delle Alpi. Rivista Di Micologia 2:75–80. Jamoni PG. 2008. Fungi alpini, delle zone alpine superiori e inferiori. Association Micologica Bresadola, Fondazione centro Studi Micologici, Trento:543. Jennings OE. 1936. The Exploration of Southampton Island, Hudson Bay. 3. Botany. 1. Algae and Fungi. Memoirs of the Carnegie Museum 12(3):1–4. Kainer D, Lanfear R. 2015. The effects of partitioning on phylogenetic inference. Molecular Biology and Evolution 32(6):1611–1627. Kallio P. 1980. Some observations on the fungi of the central Québec-Labrador peninsula. McGill Subarctic Research Report 30:1–16. 235 Kankainen E. 1969. On the structure, ecology, and distribution of the species of Mitrula s. lat. Karstenia 9:23–34. Karsten PA. 1888: Symbolae ad mycologiam fennicam. Medd. Soc. Fauna Flora Fennica 16:37–45. Kasuya T. 2010. Lycoperdaceae (Agaricales) on the Beartooth Plateau, Rocky Mountains, USA. North American Fungi 5:159–171. Kauffman CH. 1921. The mycological flora of the higher Rockies of Colorado. Papers of the Michigan Academy of Science, Arts and Letters 1:101–150. Kauffman CH. 1930. The fungus flora of the Siskiyou mountains in Southern Oregon. Papers of the Michigan Academy of Science, Arts and Letters 11:151–210. Kennedy AH, Taylor DL, Watson LE. 2011. Mycorrhizal specificity in the fully mycoheterotrophic Hexalectris Raf. (Orchidaceae: Epidendroideae). Molecular Ecology 20(6):1303–1316. Kernaghan G, Currah RS. 1998. Ectomycorrhizal Fungi at Tree Line in the Canadian Rockies. Mycotaxon 69:39–80. Kernaghan G, Harper KA. 2001. Community structure of ectomycorrhizal fungi across an alpine/subalpine ecotone. Ecography 24 (2):181–188. Kibby G, Fatto R. 1990. Keys to the species of Russula in Northeastern North America, 3rd edition. Somerville, Kibby-Fatto enterprises: 61 p. Killerman VS. 1936. Pize aus Bayern. Denkschriften der Bayer. Botanischen Gesellschaft in Regensburg 20(14):1–86. Kirk MP, Cannon PF, David JC, Stalpers JA. 2001 Dictionary of the Fungi 9th ed. CAB international, Wallingford, U.K.:458–459. Kirk MP, Cannon PF, David JC, Stalpers JA. 2008. Ainsworth & Bisby’s Dictionary of the Fungi. 10th ed. CAB International, Wallingford. Kirk PM. 2017. “Species Fungorum (version October 2017). In: Species 2000 & ITIS Catalogue of Life”. Species 2000 & IT IS. Retrieved (2019-10-21). Klepzig KD, Moser JC, Lombarder FJ, Hofstetter RW, Ayres MP. 2001. Symbiosis and competition: complex interactions among beetles, fungi, and mites. Symbiosis 30:83–96. 236 Knudsen H, Borgen T. 1982. Russulaceae in Greenland. In: Laursen GA, Ammirati JF, eds. Arctic and Alpine Mycology 1, University of Washington Press, Seattle. p. 216–244. Knudsen H, Borgen T. 1992. New and rare taxa of Russula from Greenland. Persoonia- Molecular Phylogeny and Evolution of Fungi 14(4):509–517. Knudsen H, Mukhin VA. 1998. The arctic-alpine agaric element in the Polar Urals and Yamal, Western Siberia. In: Mukhin VA, Knudsen H, eds. Arctic and Alpine Mycology 5. Russian Academy of Sciences, Ural Division. Yekaterinburg Publishers, Yekaterinburg. p 152–162. Knudsen H, Ruotsalainen J, Vauras J. 2012. Russula Pers. In: Knudsen H, Vesterholt J, eds. Funga Nordica, Agaricoid, boletoid and cyphelloid genera. Funga Nordica. Nordsvamp, Copenhagen. p. 107–148. Knudsen H, Stordal J. 1992. Russula Pers. In: Hansen L, Knudsen H, eds. Nordic Macromycetes. Nordsvamp, Copenhagen. p 374–400. Kobayasi Y. 1982. Ecology and distribution of Arctic lower fungi. In: Laursen GA, Ammirati JF, eds. Arctic and Alpine Mycology. University of Washington Press, Seattle. p. 16–26. Kobayasi Y, Hiratsuka H, Aoshima K, Korf R, Soneda M, Tubaki K, Sugiyama J. 1967. Mycological studies of the Alaskan Arctic. Annual Report of the Institute for Fermentation, Osaka. Kobayasi Y, Hiratsuka N, Otani Y, Tubaki K, Udagawa SI, Sugiyama J, Konno K. 1971. Mycological studies of the Angmagssalik region of Greenland. Bulletin of the National Science Museum 14(1):1–96. Kobayashi Y, Tubaki K, Soneda M. 1968. Enumeration of the higher fungi, moulds and yeasts of Spitsbergen. National Science Museum. Koide RT, Shumway DL, Xu B, Sharda JN. 2007. On temporal partitioning of a community of ectomycorrhizal fungi. New Phytologist 174(2):420–429. Kõljalg U, Larsson KH, Abarenkov K, Nilsson RH, Alexander IJ, Eberhardt U, Erland S, Høiland K, Kjøller R, Larsson E, Pennanen T. 2005. UNITE: a database providing web‐based methods for the molecular identification of ectomycorrhizal fungi. New Phytologist 166(3):1063–1068. Körner C. 1995. Alpine plant diversity: A global survey and functional interpretations. In: Chapin III FS, Körner C, eds. Arctic and Alpine Biodiversity: Patterns, Causes 237 and Ecosystem Consequences. Ecological Studies 113. Springer-Verlag, New York. p 45–62. Körner C. 1999. Alpine Plant Life: Functional Plant Ecology of High Mountain Ecosystems. Springer-Verlag, Berlin: 338 p. Körner C. 2003. Alpine plant life: functional plant ecology of high mountain ecosystems; with 47 tables. Springer Science and Business Media, Springer-Verlag, Berlin, Heidelberg. Kreisel H. 1969. Grundzüge eines natürlichen Systems der Pilze. Verlag VEB Gustav Fischer, Jena. 245 p. Kuchler AW. 1964. The Potential Natural Vegetation of the Conterminous United States. American Geographical Society (36), New York. Kühner R. 1975. Agaricales de la zone alpine. Genre Russula Pers. ex SF Gray. Bulletin trimestriel de la Société Mycologique de France 91(3):314–390. Kühner R, Lamoure D. 1986. Catalogue des Agaricales (Basidiomycètes) de la zone alpine du Parc National de la Vanoise et des régions limitrophes. Traveaux scientifiques du Parc national de la Vanoise 15:103–187. Læssoe T, Peterson JH. 2019. Fungi of Temperate Europe. Vol 1. Princeton Press. 813 p. Lamoure D. 1982. Agaricales de la zone alpine du Parc National des Ecrins. Travaux scientifiques du Parc national des Ecrins 2:119–123. Lamoure D, Lange M, Petersen PM. 1982. Agaricales found in the Godhavn area, W. Greenland. Nordic Journal of Botany 2:85–90. Lanfear R, Calcott B, Ho SY, Guindon S. 2012. PartitionFinder: combined selection of partitioning schemes and substitution models for phylogenetic analyses. Molecular Biology and Evolution 29: 1695–1701. Lange J. 1937. Flora Agaricina Danica. Mycologia 29(4):554–556. Lange M. 1957. Macromycetes. Part III. I. Greenland Agaricales (Pars.), Macromycetes caeteri. II. Ecological and plant geographical studies 148(2):1–125. Lange M, Skifte O. 1967. Notes on the macromycetes of Northern Norway. Acta Borealia 23:1–51. 238 Larsson K-H. 2007. Re-thinking the classification of corticioid fungi. Mycological research 111(9):1040–1063. Larsson, E. and Larsson, K-H. 2003. Phylogenetic relationships of russuloid basidiomycetes with emphasis on aphyllophoralean taxa. Mycologia 95(6): 1037- 1065. Larsson K-H, Larsson E, Kõljalg U. 2004. High phylogenetic diversity among corticioid homobasidiomycetes. Mycological Research 108(9):983–1002. Larsson E, Vauras J, Cripps CL. 2014. Inocybe leiocephala, a species with an intercontinental distribution range–disentangling the I. leiocephala–subbrunnea– catalaunica morphological species complex. Karstenia 54:15–39. Larsson E, Vauras J, Cripps CL. 2018. Inocybe praetervisa group–A clade of four closely related species with partly different geographical distribution ranges in Europe. Mycoscience 59(4):277–287. Laursen GA. 1975. Higher fungi in soils of coastal arctic tundra plant communities. Virginia Polytechnic Institute and State University Doctoral Dissertation, Blacksburg, Virginia. Laursen GA, Ammirati JF. 1982a. Arctic and Alpine Mycology: First International Symposium on Arctic-Alpine Mycology. University of Washington Press, Seattle: 559 p. Laursen GA. Ammirati JF. 1982b, Lactarii in Alaskan Arctic tundra. In: Laursen GA, Ammirati JF eds. Arctic and Alpine Mycology, The First International Symposium on Arcto-Alpine Mycology. University of Washington Press, Seattle. p 216–244. Laursen GA, Ammirati JF, Farr DF. 1987a. Hygrophoraceae from arctic and alpine tundra in Alaska. In: Laursen GA, Ammirati JF, Redhead SA, eds. Arctic and Alpine Mycology 2. Environmental Science Research 34. Plenum Press, New York, NY. p. 273–286. Laursen GA, Ammirati JF, Redhead SA. 1987b. Arctic and Alpine Mycology 2. Environmental science research 34. Plenum Press, New York, NY. 364 p. Laursen GA, Burdsall Jr HH. 1976. Notes concerning a new distribution record for Geopora (Pezizales) from Alaskan tundra. Mycotaxon 4(2):329–330. Laursen GA, Chmielewski MA. 1982. The ecological significance of soil fungi in Arctic tundra. In: Laursen GA, Ammirati JF, eds. Arctic and Alpine Mycology, The First 239 International Symposium on Arcto-Alpine Mycology. University of Washington Press, Seattle. p. 432–488. Laursen GA, Miller Jr OK. 1977. The distribution of fungal hyphae in Arctic soil on the International Biological Programme tundra biome site, Barrow, Alaska. Arctic and Alpine Research 9(2):149–156. Laursen GA, Miller Jr OK, Bigelow HE. 1976. A new Clitocybe from the Alaskan arctic. Canadian Journal of Botany 54(9):976–980. Lebel T, Tonkin JE. 2007. Australasian species of Macowanites are sequestrate species of Russula (Russulaceae, Basidiomycota). Australian Systematic Botany 20(4):355– 381. Lesica P, Antibus RK. 1986a. Mycorrhizae of alpine fell-field communities on soils derived from crystalline and calcareous parent materials. Canadian Journal of Botany 64:1691–1697. Lesica P, Antibus RK. 1986b. Mycorrhizal status of hemiparasitic vascular plants in Montana, USA. Transactions of the British Mycological Society 86:341–343. Li G, Zhao D, Li S, Wen H. 2015. Russula chiui and R. pseudopectinatoides, two new species from southwestern China supported by morphological and molecular evidence. Mycological Progress 14:33. Lind J. 1910. Fungi (Micromycetes) collected in Arctic North America (King William Land, King Point and Herschell Isl.) by the Gjöa expedition under Captain Roald Amundsen 1904-1906. Videnskabs-Selskabets Skrifter 1. Mathematisk- Naturvidenskabelig Klasse 9. Christiania (Oslo): Jacob Dybwad. Lind J. 1927. The geographical Distribution of some Arctic Micromycetes. Det Kgl. Danske Videnskabernas Selskab. Biologiske Meddelelser 6(5):1–45. Lind J. 1934. Studies on the geographical distribution of arctic circumpolar micromycetes. Kgl. Danske Videnskabernas Selskab. Biologiske Meddelelser 11(2):1–152. Linder DH. 1947. Fungi. In: Polunin N, ed. Botany of the Canadian Eastern Arctic. Part II: Thallophyta and bryophyta. National Museum of Canada Bulletin No. 97 Biological Series No. 26:234–296. Linkins AE, Antibus RK. 1978. Ectomycorrhizal Fungi of Salix rotundifolia Trautv. II: Impact of Surface Applied Prudhoe Bay Crude Oil on Mycorrhizal Root Respiration and Cold Acclimation. Arctic 31(3):381–393. 240 Linkins AE, Antibus RK. 1982. Mycorrhizae of Salix rotundifolia in coastal Arctic tundra. In: Laursen GA, Ammirati JF, eds. Arctic and Alpine Mycology, The First International Symposium on Arcto-Alpine Mycology. University of Washington Press, Seattle. p. 509–531. Lipson DA, Schadt CW, Schmidt SK, Monson RK. 1999. Ectomycorrhizal transfer of amino acid-nitrogen to the alpine sedge Kobresia myosuroides. The New Phytologist 142(1):163–167. Liu YJ, Whelen S, Hall BD. 1999. Phylogenetic relationships among ascomycetes: Evidence from an RNA polymerase II subunit. Molecular Biology and Evolution 16(12):1799–1808. Looney BP. 2014. Molecular annotation of type specimens of Russula species described by W.A. Murrill from the southeast United States. Mycotaxon 129(2):255–268. Looney BP, Adamčík S, Matheny PB. 2019. Miocene and Pliocene speciation of Russula subsection Roseinae in temperate forests of eastern North America. bioRxiv: 770289. Looney BP, Adamčík S, Matheny PB. 2020. Coalescent-based delimitation and species- tree estimations reveal Appalachian origin and Neogene diversification in Russula subsection Roseinae. Molecular Phylogenetics and Evolution: p.106787. Looney BP, Meidl P, Piatek MJ, Miettinen O, Martin FM, Matheny PB, Labbé JL. 2018. Russulaceae: a new genomic dataset to study ecosystem function and evolutionary diversification of ectomycorrhizal fungi with their tree associates. New Phytologist 218(1):54–65. Looney BP, Ryberg M, Hampe F, Sánchez-García M, Matheny PB. 2016. Into and out of the tropics: Global diversification patterns in a hyperdiverse clade of ectomycorrhizal fungi. Molecular Ecology 25(2):630–647. Löve A, Löve D. 1974. Origin and evolution of the arctic and alpine floras. In: Barry R, Ives JD, eds. Arctic and Alpine Environments. Methuen, London. p. 571–603. Ludley KE, Robinson CH. 2008. Decomposer basidiomycota in Arctic and Antarctic ecosystems. Soil Biology and Biochemistry 40(1):11–29. MacArthur RH, Wilson EO. 1967. The theory of island biogeography. Princeton University Press, Princeton, NJ: 203 p. 241 Macoun JM. 1899. XIII. A list of the plants of the Pribilof islands, Bearing sea, with notes on their distribution. The Fur seals and Fur-seal islands of the North Pacific Ocean 3:559–587. Maddison WP, Maddison DR. 2018. Mesquite: a modular system for evolutionary analysis. Version 3.51 http://www.mesquiteproject.org Madeira F, Park YM, Lee J, Buso N, Gur T, Madhusoodanan N, Basutkar P, Tivey AR, Potter SC, Finn RD, Lopez R. 2019. The EMBL-EBI search and sequence analysis tools APIs in 2019. Nucleic acids research:W636-41. Mains EB. 1955. North American hyaline-spored species of the Geoglosseae. Mycologia 47(6):846–877. Malagòn O, Porta A, Clericuzio M, Gilardoni G, Gozzini D, Vidari G. 2014. Structures and biological significance of lactarane sesquiterpenes from the European mushroom Russula nobilis. Phytochemistry 107:126–134. Massee G. 1913. Miles Joseph Berkeley 1803-1889. In: Oliver FW, ed. Makers of British Botany. Cambridge University Press, New York. p. 225–232. Matheny PB. 2005. Improving phylogenetic inference of mushrooms with RPB1 and RPB2 nucleotide sequences (Inocybe; Agaricales). Molecular Phylogenetics and Evolution 35(1):1–20. Matheny PB, Wang Z, Binder M, Curtis JM, Lim YW, Nilsson RH, Hughes KW, Hofstetter V, Ammirati JF, Schoch CL, Langer E, Langer G, McLaughlin DJ, Wilson AW, Frøslev T, Ge ZW, Kerrigan RW, Slot JC, Yang ZL, Baroni TJ, Fischer M, Hosaka K, Matsuura K, Seidl MT, Vauras J, Hibbett DS. 2007. Contributions of rpb2 and tef1 to the phylogeny of mushrooms and allies (Basidiomycota, Fungi). Molecular Phylogenetics and Evolution 43(2):430–451. M'Clintock FL. 1860. The voyage of the "Fox" in the Arctic seas: A narrative of the discovery of the fate of Sir John Franklin and his companions. John Murray, Albemarle Street, London. Melzer V, Zvára J. 1927. České holubinky (Russulae Bohemiae). Arch Príod Vyzk Čech 17:1–126. Melzer V, Zvara J. 1928. České holubinky (Russulae Bohemiae). Flore monographique des Russules de Bohême. Avec un tableau analytique des espèces. Résumé, Bulletin trimestriel de la Société mycologique de France 44:135–146. 242 Miller Jr OK. 1968. Interesting fungi of the St. Elias mountains, Yukon Territory, and adjacent Alaska. Mycologia 60(6):1190–1203. Miller Jr OK. 1969. Notes on gastromycetes of the Yukon Territory and adjacent Alaska. Canadian Journal of Botany 47(2):247–250. Miller Jr OK. 1982a. Higher fungi in Alaskan subarctic tundra and taiga plant communities. In: Laursen GA, Ammirati JF, eds. The First International Symposium on Arcto-Alpine Mycology. University of Washington Press, Seattle. p. 123–149. Miller Jr OK. 1982b. Mycorrhizae, mycorrhizal fungi and fungal biomass in subalpine tundra at Eagle Summit, Alaska. Holarctic Ecology 5(2):125–134. Miller Jr OK. 1987. Higher fungi in tundra and subalpine tundra from the Yukon territory and Alaska. In: Laursen GA, Ammirati JF, eds. Arctic and Alpine Mycology 2. Plenum Press, New York, NY. p. 287–297. Miller Jr OK. 1993. Observations on the genus Cystoderma in Alaska. In: Petrini O, Laursen GA, eds. Arctic and alpine Mycology 3. Bibliotheca Mycologica 150. J. Cramer in der Gebrüder Borntraeger Verlagsbuchhandlung, Berlin-Stuttgart. p. 161–169. Miller Jr OK. 1998. Hebeloma in the Arctic and alpine tundra in Alaska. In: Mukhin VA, Knudsen H, eds. Arctic and alpine mycology 5. Proceedings of the 5th International Symposium on Arcto-Alpine Mycology. Labytnangi, Russia. Yekaterinburg Publishers, Yekaterinburg, Russia. p 86–97. Miller Jr OK, Burdsall Jr HH, Laursen GA, Sachs IB. 1980. The status of Calvatia cretacea in arctic and alpine tundra. Canadian Journal of Botany 58(24):2533– 2542. Miller Jr OK, Evenson VS. 2001. Observations on the alpine tundra species of Hebeloma in Colorado. Gilbertson Honorary Volume. Harvard Papers in Botany 6:155–162. Miller Jr OK, Laursen GA. 1974. Belowground fungal biomass on US tundra biome sites at Barrow, Alaska. In: Holding AJ, Heal OW, Maclean Jr SF, Flanagan PW, eds. Soil Organisms and Decomposition in Tundra, Proceedings of the Microbiology, Decomposition, and Invertebrate Working Groups Meeting, Stockholm. p 151– 158. Miller Jr OK, Laursen GA. 1978. Ecto- and ectendo-mycorrhizae of arctic plants at Barrow, Alaska. In: Tieszen LL, ed. Vegetation and production ecology of an 243 Alaskan Arctic tundra. Ecological Studies 29. Springer, New York, NY. p. 229– 237. Miller Jr OK, Laursen GA, Calhoun WF. 1974. Higher Fungi in Arctic plant communities. US Tundra Biome, Ecosystem Analysis Studies, US International Biological Program, US Arctic Research Program:74–76. Miller Jr OK, Laursen GA, Farr DF. 1982. Notes on Agaricales from Arctic tundra in Alaska. Mycologia 74(4):576–591. Miller Jr OK, Laursen GA, Murray BM. 1973. Arctic and alpine agarics from Alaska and Canada. Canadian Journal of Botany 51(1):43–49. Miller Jr OK, Linkins AE, Chmielewski MA. 1978. Fungal biomass responses in oil perturbated tundra at Barrow, Alaska. Arctic 31(3):394–407. Miller SL, Aime MC, Henkel TW. 2012. Russulaceae of the Pakaraima Mountains of Guyana 2. New species of Russula and Lactifluus. Mycotaxon 121(1):233–253. Miller SL, Buyck B. 2002. Molecular phylogeny of the genus Russula in Europe with a comparison of modern infrageneric classifications. Mycological Research, 106(3):259–276. Miller SL, Larsson E, Larsson K-H, Verbeken A, Nuytinck J. 2006. Perspectives in the new Russulales. Mycologia 98(6):960–970. Miller SL, McClean TM, Walker J, Buyck B. 2001. A molecular phylogeny of the Russulaceae including agaricoid, gasteroid and pleurotoid taxa. Mycologia 93(2):344–354. Moreau PA. 2002. A la découverte des champignons de zone alpine. Bulletin Mycologique et Botanique Dauphiné-Savoie 166:5–37. Morgado LN, Semenova TA, Welker JM, Walker MD, Smets E, Geml J. 2015. Summer temperature increase has distinct effects on the ectomycorrhizal fungal communities of moist tussock and dry tundra in Arctic Alaska. Global change biology 21(2):959–972. Moser M. 1978. Kleine Kryptogamenflora Bd. IIb/2. Die Röhrlinge und Blätterpilze. Stuttgart, New York: Gustav Fischer Verlag. Moser M. 1993. Studies on North American Cortinarii. III. The Cortinarius flora of dwarf and shrubby Salix associations in the alpine zone of the Windriver Mountains, Wyoming, USA. Sydowia 45:275–306. 244 Moser M, McKnight KH. 1987. Fungi (Agaricales, Russulales) from the alpine zone of Yellowstone National Park and the Beartooth Mountains with special emphasis on Cortinarius. In: Laursen GA, Ammirati JF, eds. Arctic and Alpine Mycology 2. Plenum Press, New York, NY. p. 299–317. Moser MM, McKnight KH, Ammirati JF. 1995. Studies on North American Cortinarii I. New and interesting taxa from the greater Yellowstone area. Mycotaxon 60:301– 346. Moser MM, McKnight KH, Sigl M. 1994. The genus Cortinarius (Agaricales) in the Greater Yellowstone Area: mycorrhizal host associations and taxonomic considerations. In: Despain DG, ed. Plants and their environments, Proceedings of the First Biennial Scientific Conference on the Greater Yellowstone Ecosystem. p. 239–246. Moser M, Jülich W. 1985–2005. Farbatlas der Basidiomyceten. Stuttgart, New York: G. Fischer. Mueller GM, Wu QX. 1997. Mycological contributions of Rolf Singer: field itinerary, index to new taxa, and list of publications. Fieldiana Botany 38: 124 p. Mukhin VA, Knudsen H. 1998. Arctic and Alpine Mycology 5. Russian Academy of Sciences, Ural Division. Yekaterinburg Publishers, Yekaterinburg. Mullis KB, Faloona FA, Scharf SJ, Saiki RK, Horn GT, Erlich H. 1986. Specific enzymatic amplification of DNA in vitro: the polymerase chain reaction. Cold Spring Harbor Symposia on Quantitative Biology, Vol. LL:263–273. Mundra S, Bahram M, Eidesen PB. 2016. Alpine bistort (Bistorta vivipara) in edge habitat associates with fewer but distinct ectomycorrhizal fungal species: a comparative study of three contrasting soil environments in Svalbard. Mycorrhiza 26(8):809–818. Murray DF. 1995. Causes of arctic plant diversity: origin and evolution. In: Chapin III FS, Körner C, eds. Arctic and Alpine Biodiversity: patterns, Causes and Ecosystem Consequences. Springer-Verlag, Berlin. p. 21–32. Murrill W. 1938. New Florida agarics. Mycologia 30(4):359–371. Murrill WA. 1941. More Florida novelties. Mycologia 33(4):434–448. Murrill WA. 1943. More new fungi from Florida. Lloydia 6(3):207–221. 245 Murrill WA. 1945a. More fungi from Florida. Lloydia 7(4):175–189. Murrill WA. 1945b. New Florida fungi. Quarterly Journal of the Florida Academy of Sciences 8(2):171–189. Murrill WA. 1946. More Florida fungi. Lloydia 8(4):263–290. Nara K, Nakaya H, Hogetsu T. 2003. Ectomycorrhizal sporocarp succession and production during early primary succession on Mount Fuji. New Phytologist 158(1):193–206. Nares G. 2011. Narrative of a Voyage to the Polar Sea During 1875-6 in HM Ships Alert and Discovery: With Notes on the Natural History (Vol. 1). Cambridge University Press. Nauta M. 2010. Notes on Mollisioid Ascomycetes from the Beartooth Plateau, Rocky Mountains USA. North American Fungi (5)5:181–186. Neatby LH, Mercer K. 2008. Sir John Franklin. The Canadian Encyclopedia. Historica Canada. Retrieved April 9, 2019 from https://www.thecanadianencyclopedia.ca/en/article/sir-john-franklin Newman EI, Reddell P. 1987. The distribution of mycorrhizas among families of vascular plants. New Phytologist 106(4):745–751. Nguyen NH, Landeros F, Garibay-Orijel R, Hansen K, Vellinga EC. 2013. The Helvella lacunosa species complex in western North America: cryptic species, misapplied names and parasites. Mycologia 105(5):1275-86. Niezdoiminogo EL. 2003. Biogeographical review of agaricoid fungi from Russian Arctic. Mikologiya i Fitopatologiya 37:28–35. Noordeloos M, Gulden G. 1992. Studies in the genus Galerina from the Shefferville area on the Québec-Labrador Peninsula, Canada. Persoonia-Molecular Phylogeny and Evolution of Fungi 14(4):625–639. Oberwinkler F. 1977. Beiträge zur Biologie der niederen Pflanzen. Neue Vorstellungen über die Verwandschaftengruppen und die Stammesgeschichte der Laubmoose: 59–104. Ohenoja E. 1972. Preliminary note on the botanical research at Rankin Inlet, 1971. Musk- Ox 10:67. 246 Ohenoja E. 1975. Leotia, Cudonia, Spathularia and Neolecta (Ascomycetes) in Finland. Annals Botanici Fennici 12:123–130. Ohenoja E. 2000. Ecological aspects of the larger fungi in Northern Finnish Lapland and Adjacent parts of Norway. Micologia duemila:397–405. Ohenoja E, Ohenoja M. 1993. Lactarii of the Franklin and Keewatin districts of the Northwest Territories, arctic Canada. In: Petrini O, Laursen GA, eds. Arctic and alpine mycology 3–4. J Cramer, Stuttgart, Berlin. p. 179–192. Ohenoja E, Ohenoja M. 2010. Larger fungi of the Canadian arctic. North American Fungi 5:85–96. Ohenoja E, Ruotsalainen AL, Vauras J. 2018. Mycological records from ISAM 9, Kevo, Finland. Mycoscience 59(4):263–267. Ohenoja E, Vauras J, Ohenoja M. 1998. The Inocybe species found in the Canadian Arctic and west Siberian sub-arctic, with ecological notes. In: Mukhin VA, Knudsen H, eds. 1998. Arctic and Alpine Mycology 5. Russian Academy of Sciences, Ural Division. Yekaterinburg Publishers, Yekaterinburg. p. 106–121. Ortega A, Esteve-Raventós F. 2001. On the presence of Russula laccata in Sierra Nevada (Andalucia, Southern Spain) and ITS taxonomic relationships with R. norvegica. Mycotaxon 77:39–45. Osmundson TW, Cripps CL, Mueller GM. 2005. Morphological and molecular systematics of Rocky Mountain alpine Laccaria. Mycologia 97(5):949–972. Overholtz L. 1919. Some Colorado fungi. Mycologia 11(5):245–258. Parmelee JA. 1968. Fungi in the Canadian Arctic. North 15(2):1–5. Parmalee JA. 1969. Fungi of central Baffin Island. The Canadian Field Naturalist 83:48– 53. Parmelee JA. 1989. The rusts (Uredinales) of arctic Canada. Canadian Journal of Botany 67(11):3315–3365. Parmelee JA, Ginns J. 1986. Parasitic microfungi on vascular plants in the Yukon and environs. International Journal of Mycology and Lichenology 2(2):293–347. Peck CH. 1873. Descriptions of new species of fungi. Bulletin of the Buffalo Society of Natural Sciences 1:41–72. 247 Peck CH. 1879. Report of the Botanist (1878). Annual Report on the New York State Museum of Natural History 32:17–72. Peck CH. 1898. New species of Alabama fungi. Bulletin of the Torrey Botanical Club 25:368–372. Peck CH. 1903. Report of the State botanist 1902. Bulletin of the New York State Museum for Natural History 67:3–194. Peck CH. 1906. Edible fungi. Bull. Bulletin of the New York State Museum 105:36–44. Peck CH. 1907. New York Species of Russula. Bulletin of the New York State Museum 116:67–98. Peck CH. 1911. New species and varieties of extralimital fungi. Bulletin of the New York State Museum 150:50–65. Pegler D, Fiard JP. 1979. Taxonomy and ecology of Lactarius (Agaricales) in the lesser Antilles. Kew Bulletin 33:601–628. Peintner U. 1998. Lead and cadmium contents of Basidiomycetes along a heavily trafficated high mountain pass road in the Austrian Alps. In: Mukhin VA, Knudsen H, eds. 1998. Arctic and Alpine Mycology 5. Russian Academy of Sciences, Ural Division. Yekaterinburg Publishers, Yekaterinburg. p. 122–127. Peintner U. 2008. Cortinarius alpinus as an example for morphological and phylogenetic species concepts in ectomycorrhizal fungi. Sommerfeltia 31:161–177. Persoon CH. 1796. Seu descriptiones tam novorum, quam notabilium fungorum exhibitae. Observationes Mycologicae. 116 p. Persoon CH. 1797. Tentamen dispositionis methodicae Fungorum. Wolf, Leipzig: 76 p. Peters HA. 1962. Studies in the Genus Russula Fr. In northern California, Master thesis. San Francisco State College, San Francisco. Peterson PM. 1977. Investigations on the ecology and phenology of the macromycetes in the Arctic. Meddelelser om Grøenland Bioscience 199:2–72. Petrini O, Laursen GA. 1993. Arctic and Alpine Mycology 3–4. Bibliotheca Mycologica 150: 269 p. 248 Phookamsak R, Hyde KD, Jeewon R, Bhat DJ, Jones EG, Maharachchikumbura SS, Raspé O, Karunarathna SC, Wanasinghe DN, Hongsanan S, Doilom M. 2019. Fungal diversity notes 929–01035: taxonomic and phylogenetic contributions on genera and species of fungi. Fungal Diversity 95(1):1–273. Polunin N. 1934. The Flora of Akpatok Island, Hudson Strait. Journal of Botany 72:197– 204. Polunin N. 1940. Botany of the Canadian eastern Arctic. Part 1. Pteridophyta and Spermatophyta. National Museum of Canada Bulletin No. 92 Biological Series No. 24: 395 p. Polunin, N. 1947. Botany of the Canadian Eastern Arctic. Part II: Thallophyta and bryophyta. National Museum of Canada Bulletin No. 97 Biological Series No. 26. Rambaut A, Suchard MA, Xie D, Drummond AJ. 2014. Tracer 1.6 Available from: http://beast.bio.ed.ac.uk/Tracer. Rammeloo J, Walleyn R. 1993. The edible fungi of Africa south of Sahara: a literature review. Jardin Botanique National de Belgique. Rau TG. 1977. Fungi in decomposing litter from tundra plants near Barrow, Alaska. Virginia Polytechnic Institute and State University Doctoral Dissertation, Blacksburg, Virginia. Read DJ, Haselwandter K. 1981. Observations on the mycorrhizal status of some alpine plant communities. New Phytologist, 88(2):341–352. Redhead SA. 1980. Gerronema pseudogrisella. Fungi Canadenses No. 170. Agriculture Canada, Ottawa. Redhead SA. 1984. Arrhenia and Rimbachia, expanded generic concepts, and a reevaluation of Leptoglossum with emphasis on muscicolous North American taxa. Canadian Journal of Botany 62(5):865–892. Redhead SA. 1989. A biogeographical overview of the Canadian mushroom flora. Canadian Journal of Botany 67(10):3003–3062. Redhead SA, Baillargeon G. 1999. Fungal database development and early historical records of mushrooms from Canada. McIlvainea 14(1):73–82. Redhead SA, Miller Jr OK, Watling R, Ohenoja E. 1982. Marasmius epidryas. Fungi Canadensis No. 213. Agriculture Canada, Ottawa. 249 Redhead SA, Norvell LL. 1993. Notes on Bondarzewia, Heterobasidion and Pleurogala. Mycotaxon 48:371–380. Reid DA, 1972. Coloured illustrations of rare and interesting fungi. V. Fungorum Rariorum Icones Coloratae. Reumaux P, Bidaud A, Moënne-Loccoz P. 1996. Russules rares ou méconues. Editions Federation Mycologique Dauphiné, Savoy, Frangy. 296 p. Retzer JL. 1956. Alpine soils of the Rocky Mountains. Journal of Soil Science 7:22–32. Roberts C. 2007. Russulas of Southern Vancouver Island coastal forests, Doctoral dissertation. Univiersity of Victoria, Victoria: 667 p. Robinson CH. 2001. Cold adaptation in Arctic and Antarctic fungi. New Phytologist 151(2):341–353. Romagnesi H. 1967. Les russules d’Europe et d’Afrique du Nord. Bordas, Paris: 998 p. Ronikier A. 2008. Contribution to the biogeography of arctic-alpine fungi: first records in the Southern Carpathians (Romania). Sommerfeltia 31:191–211. Ronikier A, Adamčík S. 2009. Critical review of Russula species (Agaricomycetes) known from Tatra National Park (Poland and Slovakia). Polish Botanical Journal, 54(1):41–53. Ronikier M, Mleczko P. 2006. Observations on the mycorrhizal status of Polygonum viviparum in the Polish Tatra Mts. (Western Carpathians). Acta Mycologica 41(2):209–222. Ronikier A, Ronikier M. 2010. Biogeographical patterns of arctic-alpine fungi: distribution analysis of Marasmius epidryas, a typical circumpolar species of cold environments. North American Fungi 5:23–50. Ronquist F, Teslenko M, van der Mark P, Ayres DL, Darling A, Höhna S, Larget B, Liu L, Suchard MA, Huelsenbeck JP. 2012. MRBAYES 3.2: Efficient Bayesian phylogenetic inference and model selection across a large model space. Syst. Biol. 61:539-542. Rostrup E, Simmons HG. 1906. Fungi collected by HG Simmons on the 2nd Norwegian Polar expedition, 1898–1902. In: Report of the second Norwegian Arctic Expedition in the “Fram”, 1898–1902 2(9). Videnskabs-Selskabet I Kristiania. p. 1–10. 250 Ruotsalainen J, Huhtinen S. 2015. Type studies in Russula 1: on two species described by Kühner. Karstenia 55:61–68. Ruotsalainen J, Vauras J. 1994. Novelties in Russula: R. olivobrunnea, R. intermedia and R. groenlandica. Karstenia 34(1):21–34. Ryberg M, Andreasen M, Björk RG. 2011. Weak habitat specificity in ectomycorrhizal communities associated with Salix herbacea and Salix polaris in alpine tundra. Mycorrhiza 21(4):289–296. Ryberg M, Larsson E, Molau U. 2009. Ectomycorrhizal Diversity on Dryas octopetala and Salix reticulata in an Alpine Cliff Ecosystem, Arctic, Antarctic, and Alpine Research 41(4):506–514. Saccardo PA, Peck CH, Trelease W. 1904. The Fungi of Alaska. Harriman Alaska Expedition. Cryptogamic Botany 5:44–49. Saiki RK, Gelfand DH, Stoffel S, Scharf SJ, Higuchi R, Horn GT, Mullis KB, Erlich HA. 1988. Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science 239:487–491. Sande D, de Oliveira GP, Moura MAF, de Almeida Martins B, Lima MTNS, Takahashi JA. 2019. Edible mushrooms as a ubiquitous source of essential fatty acids. Food Research International: 108524. Sarnari M. 1998–2005. Monografia illustrata del genere Russula in Europa 1 & 2. Bresadola, Trento. Savile DBO. 1961. The botany of the northwestern Queen Elizabeth Islands. Canadian Journal of Botany 39(4):909–942. Savile DBO. 1963. Mycology in the Canadian arctic. Arctic 16(1):17–25. Savile DBO. 1972. Arctic adaptations in plants. Canada Department of Agriculture Research Branch, Monograph (6):61–75. Savile DBO, Parmelee JA. 1964. Parasitic fungi of the Queen Elizabeth Islands. Canadian Journal of Botany 42(6):699–722. Schadt CW. 2002. Studies on the fungal associations of the alpine sedge Kobresia myosuroides in Colorado. Ph.D. thesis, University of Colorado, Boulder. Schadt CW, Martin AP, Lipson DA, Schmidt SK. 2003. Seasonal dynamics of previously unknown fungal lineages in tundra soils. Science 301(5638):1359–1361. 251 Schmid-Heckel H. 1985. Zur Kenntnis der Pilze in den Nördlichen Kalkalpen. Mykologische Untersuchungen im Nationalpark Berchtesgaden. 201 p. Schmid-Heckel H. 1988. Pilze in den Berchtesgadener Alpen. Nationalparkverwaltung Berchtesgaden. Schmidt SK, Naff CS, Lynch RC. 2012. Fungal communities at the edge: ecological lessons from high alpine fungi. Fungal Ecology 5(4):443–452. Schmidt SK, Sobieniak-Wiseman LC, Kageyama SA, Halloy SRP, Schadt CW. 2008. Mycorrhizal and dark-septate fungi in plant roots above 4270 meters elevation in the Andes and Rocky Mountains. Arctic, Antarctic, and Alpine Research, 40(3):576–583. Schoch CL, Seifert KA, Huhndorf S, Robert V, Spouge JL, Levesque CA, Chen W, Fungal Barcoding Consortium. 2012. Nuclear ribosomal internal transcribed spacer (ITS) region as a universal DNA barcode marker for Fungi. Proceedings of the National Academy of Sciences 109(16):6241-6246. Seaver R, Shope PF. 1930. A mycological foray through the mountains of Colorado, Wyoming, and South Dakota. Mycologia 22(1):1–8. Semenova TA, Morgado LN, Welker JM, Walker MD, Smets E, Geml J. 2016. Compositional and functional shifts in arctic fungal communities in response to experimentally increased snow depth. Soil Biology and Biochemistry 100:201– 209. Senn-Irlet BI. 1987. Pilze aus der alpinen stufe des Val d'Anniviers (Wallis). Bulletin de la Murithienne 105:87–106. Serreze MC, Barry RG. 2011. Processes and impacts of Arctic amplification: A research synthesis. Global and planetary change 77(1-2):85–96. Shaffer RL. 1962. The subsection Compactae of Russula. Brittonia 14:254–284. Shaffer RL. 1964. The subsection Lactarioideae of Russula. Mycologia 56(2):202–231. Shaffer RL. 1970. Notes on the subsection Crassotunicatae and other species of Russula. Lloydia 33:49–96. Shaffer RL. 1972. North American Russulas of the subsection Foetentinae. Mycologia 64(5):1008–1053. 252 Shaffer RL. 1975. Some common North American species of Russula subsection Emeticinae. Nova Hedwidia Beiheft 51:207–237. Shiryaev AG, Zmitrovich IV, Ezhov ON. 2018. Taxonomic and Ecological Structure of Basidial Macromycetes Biota in Polar Deserts of the Northern Hemisphere. Contemporary Problems of Ecology 11(5):458–471. Silvestro D, Michalak I. 2012. raxmlGUI: a graphical front-end for RAxML.Organisms Diversity & Evolution 12(4):335–337. Singer R. 1938. Contribution à l’étude des Russules. 3. Quelques Russules américaines et asiatiques. Bulletin trimestriel de la Société Mycologique de France 54:132–177. Singer R. 1939a. Contribution a l’étude des Russules. 4. Quelques Russules américaines et asiatique (suite). Bulletin trimestriel de la Société Mycologique de France 55:226–232. Singer R. 1939b. Contribution a l’étude des Russules. 4. Quelques Russules américaines et asiatique (suite). Bullatin trimestriel de la Société Mycologique de France 55:233–283. Singer R. 1942. Type Studies on Basidiomycetes I. Mycologia 34(1):64–93. Singer R. 1943. Type Studies on Basidiomycetes II. Mycologia 35(2):142–163. Singer R. 1948 [1946]. New and interesting species of Basidiomycetes. II. Papers of the Michigan Academy of Science, Arts and Letters 32:103–151. Singer R. 1951. The Agaricales (mushrooms) in modern taxonomy. Lilloa 22:5–832. Singer R. 1952. Russulaceae of Trinidad and Venezuela. Kew Bulletin 7(3):295–301. Singer R. 1962. The Agaricales in modern taxonomy. J. Cramer, Weinheim: 915 p. Singer R. 1984. Tropical Russulaceae II. Lactarius section Panuoidei. Nova Hedwigia 40:435–447. Singer R. 1986. The Agaricales in modern taxonomy. Koeltz Scientific Books Koenigstein, Federal Republic of Germany: 981 p. Sitta N, Davoli P. 2012. Edible ectomycorrhizal mushrooms: International markets and regulations. In: Zambonelli A, Bonito G, eds. Edible ectomycorrhizal mushrooms. Springer, Berlin, Heidelberg. p 355–380. 253 Skifte O. 1989. Russula of the island Bjornoya (Bear Island), Svalbard. Opera Botanica 100:233–239. Smith JE, Lebel T. 2001. A comparison of taxonomic keys to species within the genus Russula. Mcllvainea, 15(1):9–22. Smith ME, Douhan GW, Rizzo DM. 2007. Intra-specific and intra-sporocarp ITS variation of ectomycorrhizal fungi as assessed by rDNA sequencing of sporocarps and pooled ectomycorrhizal roots from a Quercus woodland. Mycorrhiza 18(1):15-22. Smith SE, Read DJ. 2008. Mycorrhizal symbiosis. Academic Press, San Diego, California, USA. Solheim W. 1949. Studies on Rocky Mountain Fungi: I. Mycologia 41(6):623–631. Spence HS. 1932. Sub-arctic mushrooms. Canadian Field Naturalist 46:53–54. Sprague R, Lawrence DB. 1959. The fungi on deglaciated Alaskan terrain of known age. 1. Research Studies Washington State University 27(3):110–128. Sprague R, Lawrence DB. 1959–1960. The fungi on deglaciated Alaskan terrain of known age. 2. Research Studies Washington State University 27(4):214–229. Sprague R, Lawrence DB. 1960. The fungi of deglaciated Alaskan terrain of known age. 3. Research Studies Washington State University 28(1):1–20. Stephenson SL, Laursen GA. 1993. A preliminary report on the distribution and ecology of myxomycetes in Alaskan tundra. Bibliotheca Mycologica 150:251–257. Stöver BC, Müller KF. 2010. TreeGraph 2: Combining and visualizing evidence from different phylogenetic analyses. BMC Bioinformatics 11(7). Stucky BJ, 2012. SeqTrace: a graphical tool for rapidly processing DNA sequencing chromatograms. Journal of Biomolecular Techniques 23(3):90. Sturm M, Racine C, Tape K. 2001. Climate change: increasing shrub abundance in the Arctic. Nature 411(6837):546–547. Sturm M, Schimel J, Michaelson G, Welker JM, Oberbauer SF, Liston GE, Fahnestock J, Romanovsky VE. 2005. Winter biological processes could help convert Arctic tundra to shrubland. BioScience 55(1):17–26. 254 Tanabe Y, Watanabe MM, Sugiyama J. 2002. Are Microsporidia really related to Fungi?: A reappraisal based on additional gene sequences from basal fungi. Mycological Research 106(12):1380-1391. Tarnocai C, Canadell JG, Schuur EA, Kuhry P, Mazhitova G, Zimov S. 2009. Soil organic carbon pools in the northern circumpolar permafrost region. Global biogeochemical cycles 23(2):GB2023. Tedersoo L, Sadam A, Zambrano M, Valencia R, Bahram M. 2010. Low diversity and high host preference of ectomycorrhizal fungi in Western Amazonia, a neotropical biodiversity hotspot. The ISME journal, 4(4). p. 465. Tedersoo L, Bahram M, Toots M, Diedhiou AG, Henkel TW, Kjøller R, Morris MH, Nara K, Nouhra E, Peay KG, Polme S. 2012. Towards global patterns in the diversity and community structure of ectomycorrhizal fungi. Molecular Ecology 21(17):4160–4170. Tedersoo L, Nara K. 2010. General latitudinal gradient of biodiversity is reversed in ectomycorrhizal fungi. New Phytologist 185(2):351–354. Thiers HD. 1994. The subgenus Compactae of Russula in California. Mycologia Helvetica 2:107–120. Thiers HD. 1997a. New Species of Russula from California. Mycotaxon 63:349–358. Thiers HD. 1997b. The Agaricales (Gilled fungi) of California. 9. Russulaceae I. Russula. Mad River Press, Eureka. 158 p. Thoen E, Aas AB, Vik U, Brysting AK, Skrede I, Carlsen T, Kauserud H. 2019. A single ectomycorrhizal plant root system includes a diverse and spatially structured fungal community. Mycorrhiza 29(3):1–14. Timling I, Dahlberg A, Walker DA, Gardes M, Charcosset JY, Welker JM, Taylor DL 2012. Distribution and drivers of ectomycorrhizal fungal communities across the North American arctic. Ecosphere 3:1–25. Timling I, Walker DA, Nusbaum C, Lennon NJ, Taylor DL. 2014. Rich and cold: diversity, distribution and drivers of fungal communities in patterned-ground ecosystems of the North American Arctic. Molecular Ecology 23:3258–3272. Tondl F. 1988. Russula nana v západních Tatrách. Mykol. Listy 32:4–8. Trappe JM. 1962. Fungus associates of ectotrophic mycorrhizae. The Botanical Review 28(4):538–606. 255 Trappe JM. 1987. Phylogenetic and ecologic aspects of mycotrophy in the angiosperms from an evolutionary standpoint. Ecophysiology of VA mycorrhizal plants:5–25. Trelease W. 1904. Vol. V. 13. In: Harriman Alaska Expedition, 1899, Vol. V. containing Cryptogamic Botany of Alaska, by William Trelease, New York. Tsuji M, Hoshino T. 2019. Fungi in Polar Regions. CRC Press Taylor and Francis Group, Florida. Twieg BD, Durall DM, Simard SW. 2007. Ectomycorrhizal fungal succession in mixed temperate forests. New Phytologist 176(2):437–447. Väre H. 2017. Finnish botanists and mycologists in the Arctic. Arctic Science 3(3):525– 552. Verbeken A. 1996. Biodiversity of the genus Lactarius Pers. in tropical Africa. PhD. Thesis. Part 1. Ghent University: p. 342. Verbeken A. 1998. Studies in tropical African Lactarius species. 6. A synopsis of the subgenus Lactariopsis (Henn.) R. Heim emend. Mycotaxon 66:387–418. Vesterholt J. 1998. A check-list of fungi recorded from the Faroe Islands. Frodskaparrit 46:33–65. Vidari G, Vita-Finzi P. 1995. Sesquiterpenes and Other Secondary Metabolites of Genus Lactarius (Basidiomycetes): Chemistry and Biological Activity. Studies in natural products chemistry 17:153–206. Vila J, Llistosella J, Llimona X. 1997. Contribució al coneixement dels fongs de l’estatge alpí dels Pireneus de Catalunya. I. Revista Catalana Micol. 20:221–232. Voitk A. 2015. Russula griseascens (Bon and Gaugué) Marti. Omphalina 6(1):14. Walker MD. 1995. Patterns and causes of arctic plant community diversity. In: Chapin FS, Körner C, eds. Arctic and Alpine Biodiversity: Patterns, Causes and Ecosystem Consequences. Ecological Studies 113. Springer-Verlag, New York. p. 3–20. Wang X-H, Halling RE, Hofstetter V, Lebel T, Buyck B. 2018. Phylogeny, biogeography and taxonomic re-assessment of Multifurca (Russulaceae, Russulales) using three- locus data. PLoS ONE 13(11):e0205840. 256 Wang Q, Fan X, Wang M. 2016. Evidence of high-elevation amplification versus Arctic amplification. Scientific reports 6:e19219. Wang X, Shen J, Du J, Liu J. 2006. Marasmane sesquiterpenes isolated from Russula foetens. J. Antibiot. 59(10):669–672. Watling R. 1977. Larger fungi from Greenland. Astarte 10:61–71. Watling R. 1983. Larger cold-climate fungi. Sydowia 36:308–325. Watling R. 1987. Larger arctic-alpine fungi in Scotland. In: Laursen G, Ammirati JF, Redhead SA, eds. Arctic and alpine mycology 2. Plenum Press, New York, NY. p. 17–45. Watling R. 2005. Fungal associates of Salix repens in northern oceanic Britain and their conservation significance. Mycological Research 109(12):1418–1424. Watling R, Miller Jr OK. 1971. Notes on eight species of Coprinus of the Yukon Territory and adjacent Alaska. Canadian Journal of Botany 49(9):1687–1690. Weber WA, Siiushan S. 1955. The lichen flora of Colorado: Cetraria, Cornicularia, Dactylina, Thamnolia. University of Colorado Series in Biology 11:115–119. Weber WA. 2003. The Middle Asian element in the southern Rocky Mountain flora of the western United States: a critical biogeographical review. Journal of Biogeography 30(5):649–685. Wehmeyer LE. 1961. A world monograph of the genus Pleospora and its segregates. Ann Arbor, University Michigan Press. Wells WL, Kempton PE. 1914. Know Alaska’s Mushrooms. Cooperative Extension work in Agriculture and Home Economics. A.S. Buswell, Director University of Alaska and U.S. Department of Agriculture Cooperating, Printed and Distributed under acts of Congress:1–32. White TJ, Bruns TD, Lee S, Taylor JW, 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis MA, Gelfand DH, Sninsky JJ, White TJ, eds. PCR-protocols: A Guide to Methods and Applications. Academic Press, San Diego, p. 315–322. Wickham H. 2016. ggplot2: elegant graphics for data analysis. Springer. Woo B. 1989. Trial field key to the species of RUSSULA in the Pacific Northwest http://www.svims.ca/council/Russul.htm. 257 APPENDICES 258 APPENDIX A C-TAB EXTRACTION PROTOCOL, WITH LAB NOTES 259 CTAB DNA Extraction Protocol from Nguyen et al. (2013) Grind tissue in liquid N with a pinch of fine beads (glass beads 50–100 microns) using a mortal and pestle. Harvest the powder into 2 ml screw cap tubes. 1. Add 500 µl 2% CTAB extraction buffer, a dash of 2% PVP-40, and 2.5 µl b- mercaptoethanol to freshly ground tissue. Mix well. 2x CTAB buffer and 2% PVP-40 can be mixed in advanced and stored at room temperature. Upon using, mix with appropriate amount of b-mercaptoethanol. Add b-mercaptoethanol under a fume hood. 2. Incubate at 65 ºC for 30 minutes, shake sample every 10 minutes. For difficult or poorly preserved samples, incubate for at least 1 hour. 3. Add 500 µl isoamyl-chloroform, shake well and shake on the Nutator for 20 minutes, invert tube occasionally. Phenol-isoamyl-chloroform can also be used at this step. 4. Spin down at max speed for 5 minutes. 5. Transfer the top phase to a fresh tube and repeat steps 3 and 4 if the top phase is disturbed. 6. Add 15 µl 5M sodium acetate and 500 µl ice-cold isopropanol. Gently mix the sample. Any acetate salt can be used for this step. Cold incubation from -20ºC to -80ºC for 10 minutes is optional. You can store in -80ºC overnight to increase yield. This step causes the negatively charged DNA to clump at the bottom of the tube. 7. Spin down 3 minutes at max speed. Discard isopropanol by pipetting it out of the tube, being careful to not lose the pellet. Adding acetate salt usually causes the pellet to be goopy. 8. Wash the pellet with 500 µl ice-cold 70% ethanol by filling the same tube, make sure the pellet is loose and has been washed throughout. Incubate at room temperature for at least 5 minutes. If pellet is thick, dislodge the pellet from the side of the tube and flatten it out to allow ethanol to permeate. 9. Spin down for 1 minute and discard solution by pipetting it out of the tube. 10. Wash the pellet with 500 µl 95% ice-cold ethanol. 11. Spin down for 1 minute and discard solution by pipetting it out of the tube. 12. Completely dry the pellet at 65 ºC, either with a dry heater or vacuum centrifuge. You can also dry the pellet by pipetting out all of the ethanol and placing the tube on a clean paper towel in a positive pressure flow hood to air dry. 260 13. Reconstitute with 30–50 µl TE buffer or ddH2O. 14. The sample is now ready for PCR preparation or can be stored at -80ºC. Additional Information 1M Tris (1L pH=8.0); milipore before bottling Tris-HCl 88.75g Tris base 53g dH20 to 1L EDTA 1L .25M pH=8 .5M pH=8 Ethylenedinitrilo-tetracetic acid, dissodium salt 93.05g 186g NaOH pellets 10g 10g dH2O 800ml to 1L **Stir solution vigorously on magnetic stirrer (heated) until EDTA crystals disslove. Adjust pH to 8 with NaOH pellets. Fill to 1 or L, millipore filter and bottle 2% CTAB Ingredients for 200 ml: 2 % CTAB (4.0 g) 100 mM Tris pH 8 (20 ml of 1.0 M sol) 20 mM EDTA (16 ml of 0.25 M sol) 1.4 M NaCl (16.4 g) 1-2 % PVP polyvinylpyrrolidone 40 (4.0 g) – relieves the effects of PCR inhibitors 0.2 % Beta mercaptoethanol Add just before use (20 µl per 10 ml solution) Notes: CTAB is Hexadecyltrimethylammonium bromide. Dissolve it before adding NaCl, with stirring and a little warmth, if necessary. When the NaCl is dissolved, lots of tiny bubbles come out of solution; they rise to the surface very slowly, simulating undissolved material. PVP of 40,000 average molecular weight makes the solution slightly translucent, but no large particles should be present after dissolving. Beta-mercaptoethanol should be kept in the refrigerator in a dry box. 261 APPENDIX B PREPERATION OF PRIMERS ORDERED FROM INTEGRATED DNA TECHNOLOGIES (IDT) 262 Preparing Primers from IDT (Integrated DNA technologies) Created by Ed Barge, Ordering from IDT: go to "order custom DNA oligos", enter # of items (number of primers you are ordering), enter sequence for each primer, enter name for each primer. That's it. You don't need to mess with anything else. Once the primers arrive, briefly spin down in centrifuge to make sure all of the primer powder is at bottom of tube (you won't see anything though, as there is such a small amount of material), and then dilute to 100 µM with molecular grade water. To do this, add a volume of molecular grade water equal to 10 times the number of nanomoles (nm) of primer (nm is noted on the side of the primer tube from IDT). This is your stock solution, which should be frozen. To make the 10 µM working solution, dilute from the above stock 1:10 in molecular grade water. 263 APPENDIX C SEQTRACE PROTOCOL 264 SeqTrace Protocol Created by Ed Barge, updated by Chance Noffsinger Program downloadable at https://code.google.com/archive/p/seqtrace/downloads 1. Click “File” a. Select “New project” i. Choose Location of Trace Files ii. Set Forward to “F_” iii. Set Reverse to “R_” iv. Add Forward and Reverse Primers sequences v. Select “OK” 2. Click on the Large Blue Plus Sign a. Select all files b. Click “Add” 3. Select pair of forward and reverse sequences a. Click “Traces” i. Select “Group selected Forward/Reverse files” 1. Name Sample and hit “OK” 2. Repeat for all Forward and Reverse files 4. Select Group a. Click “Traces” i. Select “View selected trace files” 1. Edit 2. Save the working sequence to project by clicking the Blue floppy disk or file -> “Save working sequence to project” 3. Repeat for all Groups 5. After all are edited a. Click “Sequences” i. Select “Export Sequences” ii. Click either “From all trace files..” or “From selected trace files” 6. Name the file a. Select Location b. Save as FASTA c. Will open in notepad with clean sequences 265 APPENDIX D CODE REQUIRED TO RUN THE DESKTOP VERSION OF MUSCLE ON WINDOWS 266 Download Muscle from https://www.drive5.com/muscle/downloads.htm. Copy and paste the command prompt into the folder containing the muscle.exe file. Open command prompt from this folder and enter the following commands. Replace “file_name” with the actual file name. muscle -in file_name.txt -fastaout file_name _aln.fas muscle -in file_name _aln.fas -out file_name _refine.fas -refine muscle -in file_name _refine.fas -out file_name _refine_2.fas -refine muscle -in file_name _refine_2.fas -out file_name _refine_3.fas -refine muscle -in file_name _refine_3.fas -out file_name _refine_4.fas -refine 267 APPENDIX E MAXIMUM LIKELIHOOD PHYLOGENIES PRODUCED 268 Maximum likelihood phylogeny of the genus Russula produced using select sequences from the multi-locus dataset produced by Looney et al. (2016) in addition to ITS and RPB2 sequences generated in this study. Thickened branches lead to clades receiving ≥ 75% bootstrap support (BS). AA = Arctic-alpine habitats. 269 Maximum likelihood phylogeny of the Russula crown clade combining ITS and RPB2 data. Thickened branches lead to clades receiving ≥ 75% bootstrap support (BS). AA = Arctic-alpine habitats. 270 Maximum likelihood phylogeny of the Russula core clade combining ITS and RPB2 data. Thickened branches lead to clades receiving ≥ 75% bootstrap support (BS). AA = Arctic- alpine habitats. 271 Maximum likelihood phylogeny of the Brevipes clade combining ITS and RPB2 data. Thickened branches lead to clades receiving ≥ 75% bootstrap support (BS). AA = Arctic- alpine habitats. 272 APPENDIX F BEAST PROTOCOL 273 Some alignment files were additionally analyzed in BEAST (Bouckaert et al. 2019) following these instructions. In BEAUti all data partitions were selected, and the Tree models and Clock models were linked for all partitions. Under the site models tab, the Gamma category count was set to 4, the shape parameter was set to 1.0, and the Proportion Invariant sites was set to 0.2. The boxes for Proportion Invariant, Substitution Rate, and for Shape parameter were all checked, and the Subst Model was set to RB. The settings for the first site model was then cloned for all other partitions. A Relaxed Clock Log Normal and the Birth Death models were selected. The chain length was set to 50 million with Pre-Burnin set to zero. Trees were logged every 50,000 chains. The xml file was than run in BEAST. 274 APPENDIX G BEST-FITTING SUBSTITUTION MODELS DETERMINED BY PARTITION FINDER FOR BAYESIAN ANALYSES 275 Best-fitting substitution models determined by Partition Finder for Bayesian analysis of the Broad Russula phylogeny. Partition Substitution Model ITS GTR+I+G RPB1 1st Codon HKY+G RPB1 2nd Codon K80+I RPB1 3rd Codon K80+I RPB2 1st Codon GTR+I+G RPB2 2nd Codon HKY+I+G RPB2 3rd Codon GTR+G LSU GTR+I+G Best-fitting substitution models determined by Partition Finder for Bayesian analysis of the Russula core clade. Partition Substitution Model ITS SYM+I+G RPB2 1st Codon GTR+I RPB2 2nd Codon HKY+G RPB2 3rd Codon SYM+G Best-fitting substitution models determined by Partition Finder for Bayesian analysis of the Russula crown clade. Partition Substitution Model ITS GTR+I+G RPB2 1st Codon GTR+I+G RPB2 2nd Codon GTR+I+G RPB2 3rd Codon HKY+G Best-fitting substitution models determined by Partition Finder for Bayesian analysis of the Brevipes clade. Partition Substitution Model ITS GTR+G RPB2 1st Codon GTR+G RPB2 2nd Codon HKY+G RPB2 3rd Codon SYM+G 276 APPENDIX H R CODE FOR MAPS OF RUSSULA SPECIES DISTRIBUTIONS 277 ## install necessary packages library(ggplot2) library(ggmap) library(maps) library(mapdata) ## Latitude and longitude for Russula subrubens collections, W is negative, South is negative lon <- c(-109.40645, -105.5816675, -107.5378, -106.5640, 9.5018, -105.6200, -105.8792, -106.3992) lat <- c(45.00593, 40.0605419, 37.9339, 39.1086, 56.2639, 40.0892, 39.6636, 38.7606) species_loc <- c("Beartooth Plateau", "Niwot Ridge", "Cinnamon Pass", "Independence Pass", "Denmark", "Blue lake", "Loveland Pass", "Mineral Basin") species_cor <- data.frame(species_loc, lon, lat) species_cor ## f is the boarder surrounding the points I have included in my matrix, larger the ## number, bigger the map species_bbox <- make_bbox(lon = nana_cor$lon, lat = nana_cor$lat, f = .4) species_bbox ## zoom - low numbers (3 = world level) easy for R to load, higher numbers hard to load ## (20 = house level), play with to get good resolution species_map <- get_map(location = species_bbox, maptype = "terrain", source = "google", zoom = 4) ## map, can add labels to points with geom_text(data = pascua_cor, aes(label = paste(" ", ## as.character(pascua_loc), sep="")), angle = 60, hjust = 0, color = "black") ggmap(species_map) + geom_point(data = species_cor, color = "red", size = 2.5) + ggtitle("Russula subrubens")