Effects of Forest Management and Experimental Dead Wood Removal on Macrofungal Communities in Boreal, Mixedwood Forests of Ontario

Home , Amanita sinicoflava, Byssonectria, Ciboria rufofusca, Clitocybe dealbata, Cortinarius cyanites, Cortinarius trivialis

Effects of forest management and experimental dead wood removal on macrofungal communities in boreal, mixedwood forests of Ontario

Melanie Croydon-Sugarman

A thesis submitted in conformity with the requirements for the degree of Master of Science in Forestry

Faculty of Forestry University of Toronto

Ó Copyright by Melanie Croydon-Sugarman 2019

Effects of forest management and experimental dead wood removal on macrofungal communities in boreal, mixedwood forests of Ontario

Melanie Croydon-Sugarman

Master of Science in Forestry

Faculty of Forestry University of Toronto

2019

Croydon-Sugarman, Melanie. 2019. Effects of forest management and experimental dead wood removal on macrofungal communities in boreal, mixedwood forests of Ontario. Master of Science in Forestry, Faculty of Forestry, University of Toronto.

Abstract

Boreal macrofungi are an ecologically diverse group of organisms that are threatened by forest management, including harvest-associated reductions in coarse woody debris (CWD) supplies. In this thesis, I compared richness and composition of macrofungal communities in old-growth and post-logged mixedwood stands in boreal northeastern Ontario, including post-logged stands in which downed CWD availability had been experimentally manipulated. Canonical

Correspondence Analysis showed a strong distinction between macrofungal communities of unlogged and logged plots, correlated with harvest-related reductions in large-diameter CWD supplies. Rarefaction revealed that macrofungal richness, particularly of saprotrophs, was higher in unlogged compared to logged plots, especially those with experimentally-reduced CWD supplies. These results demonstrate the negative effects of post-harvest decreases in CWD on macrofungal richness and suggest that forest management in Canada may, over time, lead to the harvest-related biodiversity losses of boreal Europe. Management practices that better emulate habitat conditions in old-growth stands, including availability of large-diameter CWD, are discussed.

Acknowledgements

I would sincerely like to thank my supervisors, Dr. Jay Malcolm and Dr. Jean-Marc Moncalvo, for the opportunity to pursue this project; for Dr. Malcolm’s continuous support and insight throughout its duration; and for Dr. Moncalvo’s guidance regarding the molecular identification of the fungi collected. I would also like to thank my committee member Dr. Sandy Smith for her valuable insights provided during this thesis.

For invaluable assistance during the DNA sequencing portion of this project, I extend a huge thank-you to Simona Margaritescu at the Royal Ontario Museum. I could not have completed this huge body of work without her.

Thanks also to the Kapuskasing field crew, especially Paul Piascik, for assistance during the collecting periods and tremendous amount of help processing my samples.

Finally, thank you to everyone in the Wildlife Ecology Lab and the Graduate Department of Forestry for fostering a wonderfully supportive environment and close-knit community.

Funding for this project is provided by the Sustainable Forest Management Network, Natural Science and Engineering Research Council of Canada, Ivey Foundation, Tembec, Canadian Forest Service, Ontario Ministry of Natural Resources, and the Faculty of Forestry.

iii

Table of Contents

Abstract ...... ii Acknowledgements ...... iii List of Tables ...... vi List of Figures ...... vii List of Appendices ...... viii Chapter 1. General Introduction ...... 1 Fungi in boreal forests ...... 4 Forest management effects on fungi in Fennoscandia ...... 10 Forest management effects on fungi in Canada ...... 16 Chapter 2. Effects of forest management and experimental dead wood removal on macrofungal communities in boreal, mixedwood forests of northeastern Ontario ...... 21 Introduction ...... 21 Methods ...... 29 Study sites ...... 29 Coarse downed woody debris manipulation experiment...... 30 Downed woody debris sampling...... 31 Additional habitat variables ...... 32 Fruiting body sampling ...... 33 DNA extraction and ITS sequencing ...... 35 BLAST sequence alignment and phylogenetic analysis ...... 37 Statistical analyses ...... 39 Results ...... 41 Molecular identification ...... 41 Fruiting body sampling ...... 42 Macrofungal community variation ...... 44 Rarefaction ...... 45 Richness of ecological guilds ...... 46 Discussion ...... 48 Fruiting body sampling ...... 48 Molecular identification ...... 53

Community composition ...... 55 Ecological guilds ...... 60 Chapter 3. General Conclusions ...... 64 Literature Cited ...... 65 Tables ...... 96 Figures ...... 99 Appendices ...... 103

List of Tables

Table 1. Characteristics of the boreal mixedwood plots sampled for fungi in the vicinity of Kapuskasing, Ontario in 2011, including location, treatment, year of origin, stand replacing disturbance type, basal areas of common tree species, percent deciduous cover, position on shrub-derived ecological gradients, and shrub density. See text for details.

Table 2. Significance levels from permutation tests for canonical correspondence analyses of fungal communities of boreal mixedwood stands sampled in three sampling periods constrained by various habitat variables (n.s. = not significant [p > 0.05]). Coarse woody debris (CWD) variables were measured by wood volume; environmental gradients corresponding to light exposure/conifer composition (DCA1) and soil moisture gradient (DCA2) were from detrended correspondence analyses on shrub community variation (see text for details).

Table 3. Mean (± SE) species richness in each of four ecological guilds and the total number of morphospecies across treatment plots (FR = full-removal, HR = half-removal, CO = control, UL = unlogged) during three sampling periods in mixedwood forests of boreal northeastern Ontario. Letters in common indicate no significant difference between treatment means (Tukey’s HSD, α=0.05). ECM = ectomycorrhizal morphospecies; WS = wood saprotrophs; LS = litter saprotrophs; PARA = parasites; Total = total richness.

List of Figures

Figure 1. Canonical correspondence analyses conducted on macrofungal species assemblages in unlogged and logged plots in mixedwood forests of northeastern Ontario from sampling periods 1 (upper), 2 (mid) and 3 (lower). Asterisks denote environmental variables with significant effects in constraining community composition (* = p<0.05, ** = p<0.01, *** = p<0.005, **** = p<0.001, ***** = p<0.0005). vtot = total CWD volume; vlc_p_u1 = large-diameter, early-decay conifer CWD; vlc_p_u3 = large-diameter, late-decay conifer CWD; vld_p_u1 = large-diameter, early-decay deciduous CWD; vld_p_u3 = large-diameter, late-decay deciduous CWD; vsc_p_u1 = small-diameter, early-decay conifer CWD; vsc_p_u3 = small-diameter, late-decay conifer CWD; vsd_p_u1 = small-diameter, early-decay deciduous CWD; vsd_p_u3 = small-diameter, late-decay deciduous CWD; perc_dec = percent basal area of deciduous trees; shrub_st = shrub stem density; DCA1 = light exposure and conifer composition gradient (low end = shade- intolerant, high end = shade-tolerant); DCA2 = soil moisture gradient (low end = hydric conditions, high end = mesic conditions).

Figure 2. Sample-based species accumulation curves and 95% confidence intervals based on macrofungal fruiting body surveys in four treatment groups (UL plots = black circles, CO plots = red circles, HR plots = green triangles, FR plots = blue triangles) from 37 mixedwood boreal forest sites near Kapuskasing, Ontario across sampling periods 1 (upper), 2 (mid) and 3 (lower). Fruitbodies were identified based on morphological characteristics and ITS sequence similarity with top BLAST hits and construction of maximum likelihood trees.

Figure 3. Box plot showing average species richness of wood saprotrophs in the four treatments (FR = full CWD removal; HR = half CWD removal; CO = control; UL = unlogged) in mixedwood forests of northeastern Ontario sampled in the first sampling period. F-value and p- value (upper left) are from an analysis of variance; letters in common indicate no significant difference between treatment means (Tukey’s HSD, a=0.05).

Figure 4. Redundancy analyses conducted on macrofungal ecological guilds in unlogged and logged plots in mixedwood forests of northeastern Ontario from sampling periods 1 (upper), 2 (mid) and 3 (lower). ECM = ectomycorrhizal morphospecies, litter_sap = litter saprotrophs, para = parasites, total_species = total number of morphospecies, wood_sap = wood saprotrophs. Asterisks denote environmental variables with significant effects in constraining community composition (* = p<0.05, ** = p<0.01, *** = p<0.005, **** = p<0.001, ***** = p<0.0005). vtot = total CWD volume; vlc_p_u1 = large-diameter, early-decay conifer CWD; vlc_p_u3 = large- diameter, late-decay conifer CWD; vld_p_u1 = large-diameter, early-decay deciduous CWD; vld_p_u3 = large-diameter, late-decay deciduous CWD; vsc_p_u1 = small-diameter, early-decay conifer CWD; vsc_p_u3 = small-diameter, late-decay conifer CWD; vsd_p_u1 = small-diameter, early-decay deciduous CWD; vsd_p_u3 = small-diameter, late-decay deciduous CWD; perc_dec = percent basal area of deciduous trees; shrub_st = shrub stem density; DCA1 = light exposure and conifer composition gradient (low end = shade-intolerant, high end = shade-tolerant); DCA2 = soil moisture gradient (low end = hydric conditions, high end = mesic conditions).

vii

List of Appendices

Appendix I. ITS sequences of fruiting bodies sampled from unlogged and logged plots in mixedwood forests of northeastern Ontario. The collection number, phylum and morphospecies name are listed, as well as the top BLAST hit from GenBank or UNITE, its accession number, species name and country of origin.

Appendix II. Morphospecies of fruiting bodies sampled from unlogged and logged plots in mixedwood forests of northeastern Ontario, together with the total number of occurrences recorded and ecological guild assignments. Ones indicate assignment to the respective guild.

viii 1

Chapter 1. General Introduction

Conservation of biodiversity and ecosystem function are increasingly recognized as critical components of forest management (Montreal Process 1995, Larsson and Danell 2001,

Lindenmayer and Franklin 2002, CBD 2009, Aerts and Honnay 2011). As one of the largest terrestrial ecosystems, forests cover almost one third of earth’s land area and contain more than

80% of terrestrial biodiversity (United Nations International Year of Forests, 2011). They mediate global biogeochemical cycles (Hartmann et al. 2012) and comprise some of the largest global carbon (C) sinks and stores, helping to mitigate the effects of global warming (Bonan

2008, Miles and Kapos 2008, Fahey et al. 2010). Globally, forest ecosystems and biodiversity are threatened by forest management, specifically harvesting (Esseen et al. 1997, Siitonen 2001,

Grove 2002, Millennium Ecosystem Assessment 2005), that alters natural stand dynamics and composition (Brumelis and Carleton 1988, Carleton and MacLellan 1994) and reduces the availability of dead wood supplies (Bader et al. 1995, Siitonen et al. 2000, Sippola et al. 2001).

The boreal forest is the planet's largest terrestrial biome (Dahlberg 2002), covering 11% of the land area (Bonan and Shugart 1989) and comprising approximately one quarter of closed- canopy forests globally (Burton et al. 2010). It is located between 45° and 70°N in a circumpolar belt across North America (Canada and Alaska) and Eurasia (Russia and Fennoscandia) (Stocks

1991, Golammer and Furyaev 1996, Weber and Stocks 1998, Stocks et al. 2001, Lindahl et al.

2006) and is dominated by conifers including Picea, Pinus, Abies and Larix (Dahlberg 2002).

Although plant species richness is relatively poor (Kaila et al. 1995, Dahlberg 2002), these forests include speciose groups of saproxylic (wood-using) insects and fungi. The dead wood resources of boreal forests are therefore of particular interest from a biodiversity perspective, especially for fungal communities (see below, Fungi in boreal forests).

In natural (unmanaged) boreal forests, dead wood is produced from periodic disturbances including fire (Johnson 1992, Linder et al. 1997, Amiro et al. 2001, 2002, Karjalainen and

Kuuluvainen 2002), windthrow, and snow breakage, as well as by gap-forming insect outbreaks, disease, drought, and natural tree mortality (Gromtsev 2002, Selikhovkin 2005). Disturbances can produce large volumes of both downed woody debris (DWD) and standing dead trees (snags)

(Spies et al. 1988, Uotila et al. 2001) that decompose at varying rates according to diameter, tree species and local abiotic conditions, as well as through succession of the decay community

(Harmon et al. 1986, Bader et al. 1995, Kruys et al. 1999, Kruys and Jonsson 1999, Tarasov

1999, Siitonen 2001, Heilmann-Clausen and Christensen 2004, Nordén et al. 2004, Küffer et al.

2008, Lonsdale et al. 2008). Coarse woody debris (CWD) includes fallen trunks (logs), large branches, stumps and standing dead trees (snags) generally >7.5 cm diameter (Stevens 1997,

Woodall and Williams 2005); fine woody debris (FWD) comprises smaller dead wood <7.5 cm diameter, including small branches and fragmented wood pieces (Küffer and Senn-Irlet 2005).

CWD is integral to ecosystem functioning in boreal forests in that it plays key roles in nutrient cycling and biodiversity provisioning (Harmon and Hua 1991, Samuelsson et al. 1994,

Siitonen 2001). The persistence of this substrate makes it a valuable sink for long-term storage of atmospheric C (Yatskov et al. 2003) and a slow-release source of organic matter and limiting nutrients such as K, N and P (Siitonen 2001, Bani et al. 2018). These nutrients and the CWD substrate itself are used during the decay process for food and habitat (Harmon et al. 1986,

Franklin et al. 1987, Spies et al. 1988, Samuelsson et al. 1994, Siitonen 2001, Jonsson et al.

2005) by a suite of species including bacteria, fungi, invertebrates, small mammals, mosses and lichens (Franklin et al. 1981, Maser and Trappe 1984, Harmon et al. 1986, Samuelsson et al.

1994, Esseen et al. 1997, Crites and Dale 1998, Siitonen 2001, Küffer and Senn-Irlet 2005,

Woodall and Westfall 2005). As a result of the spatial heterogeneity and complex dynamics of decomposition, numerous microhabitats can exist within a single dead wood piece; for example, unequal decomposition along the length of a log can support both early- and late-decay taxa simultaneously (Heilmann-Clausen and Christensen 2003).

Unfortunately, forest management in the boreal forest has in some cases altered the quantity, quality, and continuity of dead wood substrates, with clearcutting reducing the volume of dead wood below levels produced following natural disturbances (Fleming and Freedman

1998, Niemelä 1999, Krankina et al. 2002). CWD is particularly impacted by clearcut harvesting practices (Linder and Östlund 1998, Sippola et al. 1998, Siitonen 2001), as the majority of large- sized timber is removed during harvesting (Müller-Using and Bartsch 2009). These reductions in

CWD negatively affect communities of saproxylic species (Andersson and Hytteborn 1991,

Bader et al. 1995, Høiland and Bendiksen 1996, Kruys et al. 1999, Siitonen 2001, Stenlid et al.

2008). Although harvesting may initially produce relatively large volumes of DWD (Kruys et al.

1999, Siitonen 2001, Toivanen et al. 2012), over time these operations can deplete dead wood supplies as a result of fibre removals and reductions in forest age (Hansen et al. 1991). Because dead wood supplies accumulate slowly as forests age (Spies et al. 1988), short rotation times can also reduce C storage in CWD (Harmon et al. 1990).

Decreases in dead wood availability due to harvest operations are likely to be more severe under aggressive fibre removals for biofuel, including extraction of logging residues

(branches, treetops, stumps) and non-commercial species (Berglund and Åström 2007, Egnell et al. 2007, Allmér et al. 2009, Riffell et al. 2011, Toivanen et al. 2012). Although the long-term, landscape-level effects of such increases in dead wood removal are poorly known, they could exacerbate the effects of harvesting on saproxylic taxa (Rudolphi and Gustafsson 2005, Eräjää et

4 al. 2010, Rabinowitsch-Jokinen and Vanha-Majamaa 2010). For example, slash removal has been shown in the short-term to affect populations of ground beetles, mosses, and lichens

(Åström et al. 2005, Nittérus et al. 2007, Caruso 2008) and removal of logging residues for biofuel has been shown to reduce fungal diversity in boreal forests (Wästerlund and Ingelög

1981, Mahmood et al. 1999, Toivanen et al. 2012).

Fungi in boreal forests

Boreal macrofungi (i.e., those with a visible fruiting body; Watling 1995) comprise a species-rich assemblage with important ecological roles that are highly integrated with those of dead wood (Stokland et al. 2012, Moose et al. 2019). As a result, post-harvest decreases in dead wood supplies may have negative effects on ecosystem functioning in part because of effects on fungal communities (Stenlid et al. 2008, Toivanen et al. 2012). As macrofungi facilitate boreal biodiversity, forest management has the potential to affect both fungi and a suite of fungal-reliant species (Siitonen 2001). Many insects and other arthropods use fungal fruiting bodies for habitat

(e.g., Kaila et al. 1994), while mycelia and fruitbodies are important components of terrestrial food webs (Moore et al. 2004, Peay et al. 2008), providing a primary C source for soil- and wood-inhabiting organisms (Harmon et al. 1986, Crites and Dale 1998, Siitonen 2001, Wardle

2002, Lonsdale et al. 2008).

The lack of vertical mixing amongst boreal soil horizons restricts different ecological guilds of fungi to particular niches depending on whether they assimilate C as saprotrophs or as root-associated taxa such as ectomycorrhizal species (Lindahl et al. 2007, Clemmensen et al.

2015, Bödeker et al. 2016). In this sense, the roles of macrofungi in nutrient cycling are largely determined by their habitat and mode of nutrition (Lindahl et al. 2006) and, in turn, can be

5 grouped into four main ecological guilds: wood saprotrophic, litter saprotrophic, ectomycorrhizal

(ECM) and parasitic. As a result of their direct or indirect dependence on dead wood or wood- associated organisms for some or all of their life cycle (Speight 1989), both saprotrophs and other functional groups (Dix and Webster 1995, Nordén et al. 1999) can be considered saproxylic, and are potentially susceptible to reductions in dead wood availability.

Saprotrophic fungi are the primary decomposers of wood and leaf litter in boreal forests

(Rayner and Boddy 1988), making them important drivers of C and nutrient cycling (Harmon et al. 1986, Lindahl et al. 2006, Lonsdale et al. 2008). Litter decomposition rates and C storage are influenced not only by substrate quality and abiotic factors (Hobbie et al. 2006, Cornwell et al.

2008), but also by the diversity and spatial stratification of saprotrophic organisms (Setala and

McLean 2004, Hattenschwiler et al. 2005); therefore, management-induced changes to saprotrophic fungal communities could affect nutrient cycling in boreal ecosystems.

Wood saprotrophic fungi are the principal agents of wood decomposition in boreal forests

(Rayner and Boddy 1988, Moore et al. 2004), directly modifying resource availability for other functional groups of organisms (Harley 1971, Jones et al. 1994, Krajick 2001, Moore et al.

2004). These fungi colonize recently-dead or decaying wood in successional assemblages that reflect the age, tree species, diameter, and decay class of the wood substrate and interspecific, competitive interactions with other fungi (Boddy et al. 1985, Renvall 1995, Høiland and

Bendiksen 1996, Boddy 2000, Heilmann-Clausen 2001, Siitonen 2001, Küffer and Senn-Irlet

2005, Bässler et al. 2012, Abrego and Salcedo 2013). Their obligate association with dead wood makes these species particularly susceptible to management-induced reductions in dead wood supplies, especially reductions in CWD availability (Samuelsson et al. 1994, Bader et al. 1995,

Bredesen et al. 1997, Jonsson and Kruys 2001, Nilsson et al. 2001).

The majority of wood saprotrophic fungi belong to the order Polyporales in the

Basidiomycota. Polypores typically form hard, often perennial fruitbodies and include the genera

Cerioporus, Daedaleopsis, Fomes, Fomitopsis, Ganoderma, Lentinus, Lenzites, Neofavolus,

Perenniporia, Polyporus, Postia, Rhodofomes and Trametes. Other corticioid (crust-like) genera traditionally placed within the order Aphyllophorales (e.g. Lõhmus 2011) include the genera

Hymenochaetopsis, Stereum and Trichaptum. Wood saprotrophs also include agaricoid

(mushroom-forming) genera such as Galerina, Mycena, Pluteus and Simocybe and members of the Ascomycota including Bisporella, Chlorociborium, Pseudoplectania, Scutellinia and Xylaria.

Primary colonizers of decaying wood include the wood saprotrophic brown and white rot fungi, which break down recalcitrant lignocellulose (Haider and Trojanowski 1980, Kirk and

Fenn 1982) into smaller molecules. Brown rot fungi (e.g., Fomitopsis pinicola) degrade both cellulose and hemicellulose and are typically associated with coniferous wood. In contrast, white rot fungi (e.g., Fomes fomentarius) produce both cellulolytic and lignolytic enzymes and are generally associated with deciduous wood (Goodell et al. 2008). The products of lignocellulose degradation by brown- and white-rot fungi may then be used by other saproxylic fungi (Renvall

1995, Boddy 2001) or become humified as soil organic matter (Crawford et al. 1990, Lumley et al. 2001, Fukasawa et al. 2009, 2010) to be utilized by other guilds of soil fungi.

As obligate saproxylics (but see Allmér et al. 2009), wood saprotrophs require a dead wood substrate throughout their life cycle and are therefore particularly sensitive to variations in dead wood availability, including reductions due to forest management (Siitonen 2001, Sippola et al. 2004, Junninen and Komonen 2011). Due to their specialized substrate requirements

(Renvall 1995, Penttilä et al. 2004), polypores are particularly susceptible to post-harvest reductions in dead wood supply, especially reductions in CWD (Kotiranta and Niemelä 1996,

Ohlson et al. 1997, Penttilä et al. 2004, Junninen et al. 2006). As a result, these fungi are useful indicators of dead wood continuity over time (Kotiranta and Niemelä 1996).

Litter saprotrophic fungi typically colonize litter, humus and soil and contribute to soil humification through the decomposition of cellulose and hemicelluloses (Baldrian 2008).

Members of this ecological guild include the Basidiomycota genera Clitocybe, Collybia,

Crinipellis, Gymnopus, Marasmius, Tephrocybe and Tubaria. Because the decomposition of dead wood is a continuous gradation from larger pieces to smaller, ‘litter-sized’ fragments, the definition of a ‘litter saprotroph’ often includes taxa that could also be described as wood saprotrophs. This functional overlap between wood and litter saprotrophs is evident in the retention of functional ligninolytic enzyme systems, including laccases and magnesium peroxidases, in litter saprotroph species (Baldrian 2006, Baldrian 2008). These enzymes are used to break down lignin directly or to decompose humified polyphenolic compounds that result from lignin degradation by wood saprotrophs (Steffen et al. 2002).

The ability to utilize lignin and lignin-derived compounds as alternative sources of C and

N suggests that litter-decay fungi could also be negatively impacted by reductions in dead wood availability due to forest management. Since these nutrients are translocated through the fungal mycelium (Lindahl and Olsson 2004, Lindahl and Boberg 2008), reductions in lignin or its by- products could affect the litter-decomposing ability of these fungi by reducing additional nutrient reserves. Unfortunately, this possibility has received little research attention as yet, although

Allmér et al. (2009) showed that the richness and frequency of fungi in wood and needle litter was not significantly different in stands with and without slash removal for biofuel after 25 years, suggesting that a reduction in FWD through logging residue removal did not affect the community of litter saprotrophs in the long-term.

Ectomycorrhizal (ECM) fungi, which form mutualistic associations with the roots of trees and shrubs and facilitate below-ground nutrient exchange through extensive fungal-root networks (Courty et al. 2010, Simard et al. 2012), comprise a third functional guild (Marks and

Kozlowski 1973, Sanders et al. 1975, Finlay 2008). In return for photoassimilates produced by the host plant, ECM fungi enhance the acquisition and translocation of soil nutrients including N

(Martin 1985, Chalot and Bran 1998) and P (Smith and Read 2008) across nutrient-poor boreal soils, into host plant roots (Boddy and Watkinson 1995; Hari and Kulmala 2008). Members of this ecological guild include the genera Amanita, Cortinarius, Inocybe, Laccaria, Lactarius,

Russula and Suillus.

Although ECM fungi generally inhabit soil, many species are facultative saproxylics that use well-decayed CWD as refugia for growth and fruiting (Tedersoo et al. 2003). This substrate also provides microsites in which trees and shrubs germinate and establish (Harmon et al. 1986,

Veblen 1989, St. Hilaire and Leopold 1995, Gray and Spies 1997), including late-decay CWD

‘nurse logs’ that host numerous fine roots with ECM root tips (Kirk 1966, Harvey et al. 1978,

1979, Christy et al. 1982, Vogt et al. 1995, Goodman and Trofymow 1998). Favourable microsites in dead wood are characterized by higher moisture, temperature, softness and erosion resistance than forest soils, increasing root growth and mycorrhizal root biomass (Harvey et al.

1978, DeLong et al. 1997).

The ECM state is evolutionarily derived, with multiple origins from both wood and soil saprotrophs (Kohler et al. 2015). The switch to an ECM mode of nutrient acquisition is convergent with the loss of genes that facilitate a saprotrophic lifestyle, including those encoding plant cell wall-degrading enzymes (Kohler et al. 2015, Fesel and Zuccaro 2016). As a result, reversions to saprotrophy are unlikely (Bruns and Shefferson 2004, Tedersoo and Smith 2013).

Despite these losses, certain ECM fungi retain saprotrophic capacities (Gramss et al.

1998, Lindahl and Taylor 2004, Egger 2006, Courty et al. 2007, Courty et al. 2010), including the production of enzymes that break down polyphenolic compounds (Read and Perez-Moreno

2003, Lindahl et al. 2005) and retention of functional genes for enzymes such as laccases and lignin peroxidases that degrade complex organic compounds including lignin (Chambers et al.

1999, Nehls et al. 2001, Lindahl and Taylor 2004, Luis et al. 2005, Bödeker et al. 2009). As a result, although plant-produced carbohydrates are the primary source of energy for ECM fungi, some may utilize hemicellulose, cellulose and humic polymers from litter and soil organic matter

(Durall et al. 1994), or decompose dead wood directly, although not to the extent of saprotrophs

(Colpaert and Van Laere 1996, Talbot et al. 2008).

ECM fungi have been shown to compete with saprotrophic fungi in the colonization of dead wood, absorbing nutrients released by the action of wood saprotrophs (Tedersoo et al.

2003). There is also evidence that interspecific competition, specifically antagonistic effects of mycorrhizal fungi on litter saprotrophs, shapes the realized niches of these functional groups

(Bödeker et al. 2016). Management-induced reductions in dead wood resources could therefore affect both saprotrophic and ECM species, including their richness and composition, interactions among them, and, by extension, mycorrhizal functions necessary for forest productivity and regeneration (Tedersoo et al. 2008).

A final ecological guild comprises parasitic (biotrophic) fungi that require a living host as their source of nutrition (Kendrick 1992). These include taxa that inhabit dead wood (e.g.

Armillaria spp.) or mosses (e.g., Hygrocybe and Rickenella spp.) and entomopathogenic species that attack insects and other arthropods (e.g., Cordyceps). Reductions in dead wood, particularly late-decay CWD utilized as habitat for both mosses (Söderström 1988a, Andersson and

Hytteborn 1991) and diverse invertebrate assemblages (Esseen et al. 1992, 1997), could therefore affect parasitic fungi.

Forest management effects on fungi in Fennoscandia

The great majority of research on the effects of forest management on dead wood supplies and fungal communities has occurred in Fennoscandia (boreal Europe), where increasingly intensive forest management over the past several centuries (Larsson and Danell

2001) and particularly over the past 100-150 years (Östlund 1993) has resulted in the loss and fragmentation of unmanaged forests, truncated forest stand age distributions, and simplified forest structure at stand and landscape levels, particularly through reductions in the availability of dead wood (von Berg 1859, Heikinheimo 1915, Esseen et al. 1997, Östlund et al. 1997, Linder and Östlund 1998, Siitonen 2001, Uotila et al. 2002, Gibb et al. 2005). As a result, many boreal species in the region have become restricted to increasingly small, isolated, and poor-quality forest areas (Penttilä et al. 2006, Berglund and Jonsson 2008) and many are now rare or threatened, particularly those that rely on dead wood (Gärdenfors 2005, Rassi et al. 2010).

Saproxylic fungi, primarily wood saprotrophs, are well-known to be strongly impacted by management-induced reductions in the quantity, quality and continuity of dead wood that has occurred in Fennoscandia, particularly the scarcity of CWD in managed stands (Bader et al.

1995, Lindblad 1998, Siitonen 2001, Penttilä et al. 2004, Sippola et al. 2004, 2005, Hottola and

Siitonen 2008, Hottola et al. 2009). Reduced CWD volume reduces population sizes and population connectivity, increasing the likelihood of local extinctions and jeopardizing the persistence of saproxylic fungi (Siitonen 2001, Penttilä et al. 2006). Polypores in particular are sensitive to such environmental changes, as they are often specialized in their substrate

11 requirements (Renvall 1995, Penttilä et al. 2004). Many are now red-listed in boreal Europe

(Berglund et al. 2011) and serve as indicators of stands with high conservation value (Karström

1993, Aanderaa et al. 1996, Nitare 2000, Niemelä 2005, Halme et al. 2009a, Halme et al. 2009b).

In general, the species richness of wood saprotrophic fungi has been found to decrease with decreasing amounts of dead wood substrates (e.g. Sippola and Renvall 1999, Allen et al.

2000, Humphrey et al. 2000, Edman et al. 2004b, Penttilä et al. 2004, Berglund and Jonsson

2005, Heilmann-Clausen and Christensen 2005, Schmit 2005, Sippola et al. 2005, Similä et al.

2006, Ódor et al. 2006, Hottola et al. 2009). Depending on the type of forest management practice, reductions in CWD volume can occur at the stand and landscape levels concurrently. In

Fennoscandia, an estimated 90-98% reduction in average CWD volume has occurred at the landscape level as a result of forest management, from 60-90 m3 ha-1 in old-growth forests to between 2-10 m3 ha-1 in managed forests (Siitonen 2001). Under current harvesting practices, these stands are unlikely to support taxa requiring a large volume of stand-level CWD, including red-listed polypore species such as those in mature spruce forest stands. In these stands, a threshold volume of dead wood for the occurrence of rare polypores and, therefore, for the occurrence of the maximum number of polypore species, has been estimated to be 20-40 m3 ha-1

(Penttilä et al. 2004, Hottola et al. 2009).

Reductions in the type (quality) of CWD also impact assemblages of saproxylic fungi. A number of specialist species that favour or are restricted to large-diameter, late-decay CWD

(Bader et al. 1995) are particularly at risk and may be rare or locally extinct in managed stands where these logs are uncommon or absent (Siitonen 2001, Penttilä et al. 2004, Sippola et al.

2004, Junninen et al. 2006). In Fennoscandia, polypore richness per dead wood piece has been shown to increase with increasing diameter of the dead wood substrate (Junninen and Komonen

2011 and references therein) and for spruce logs, a threshold diameter of c. 20-30 cm is needed for large-diameter specialist taxa to appear alongside more generalist taxa (Bader et al. 1995,

Renvall 1995, Sippola et al. 2004, Siitonen et al. 2005, Junninen and Komonen 2011).

The relative importance of the diameter of dead wood in comparison to volume is not always straightforward. Although CWD size has been shown to correlate positively with the number of fruiting wood saprotrophic species, this may partly be an effect of volume (a larger volume is likely to contain more species; e.g., Bader et al. 1995, Renvall 1995, Lindhe et al.

2004) or surface area (when equal numbers of logs are compared, CWD has a greater surface area than FWD; Kruys and Jonsson 1999). However, Heilmann-Clausen and Christensen (2004) found that although saproxylic fungal richness increased with CWD size, richness per unit volume decreased; this was due to a larger surface area per unit volume for small vs large CWD and because, for a given volume of CWD, a larger number of pieces of small-diameter dead wood need to be sampled. Both of these aspects correspond to increased colonization opportunities for fungi (Lonsdale et al. 2008). Similarly, when equal volumes of CWD and FWD were compared, the richness of spruce log cryptogams was greater on FWD as a result of increased surface area and number of logs (Kruys and Jonsson 1999). At low total volumes of dead wood, richness was found to increase with the proportion of FWD; at higher volumes of dead wood, richness increased with the proportion of CWD with the appearance of rare taxa

(Kruys and Jonsson 1999).

While most polypore species occur on dead wood at mid-decay stages (e.g., Bader et al.

1995, Høiland and Bendiksen 1997, Lindblad 1998, Kruys et al. 1999, Groven et al. 2002,

Stokland and Kauserud 2004, Siitonen et al. 2005, Sippola et al. 2005, Junninen et al. 2007,

Jönsson et al. 2008,), late-decay stages are required for the majority of red-listed species

(Renvall 1995, Tikkanen et al. 2006) and other late-successional species that require moss- covered and highly-decayed downed CWD (Siitonen 2001). The importance of both volume and quality of CWD in shaping macrofungal communities is evident from red-listed polypores, whose species richness has been found to be best predicted not only by the total number of logs, but in particular by the number of large logs (Hottola et al. 2009). Furthermore, fungi that depend on predecessor fungi (for example, Pycnoporellus fulgens follows colonization by

Fomitopsis pinicola; Niemelä et al. 1995), including certain late-decay taxa (Siitonen 2001), may be impacted by changes in local species composition that result from clearcutting (Sippola and

Renvall 1999, Kauserud et al. 2005, Junninen et al. 2006, 2008).

An additional important qualitative dead wood feature is the species. A large number of saproxylic macrofungi are specialists on either coniferous or deciduous wood. In eastern

Fennoscandia, Nordén et al. (2013) found that most wood saprotrophic fungi surveyed were specialists (>90% occurrences) on either coniferous (36%) or deciduous (42%) dead wood, while only 22% were classified as generalists. Forest management that specifically targets conifers

(MacDonald 1995) may therefore impact conifer-specialist taxa (Stokland and Larsson 2011).

Reductions in the spatiotemporal continuity of CWD at the stand level also occur as a result of harvesting (Siitonen 2001), potentially impacting saproxylic fungal richness by reducing colonization opportunities (Jönsson et al. 2008). Even if new dead wood substrates become available in the future through succession or disturbance, successful colonization cannot occur without an initial population within dispersal range of the substrate (Siitonen 2001). In

Fennoscandia, polypore richness has been shown to increase steeply for the first 20 ha of stand area, indicating that smaller stands are not as likely to contain a species-rich assemblage

(Junninen and Komonen 2011). Polypores are also more likely to establish close to dead wood

14 that has already been colonized (Edman et al. 2004b, Jönsson et al. 2008), with decreases in dead wood continuity increasing the likelihood of local extinctions (Penttilä et al. 2004, Stokland and

Kauserud 2004).

Habitat isolation has been shown to impact polypores in Fennoscandia, with large old- growth spruce forests providing better habitat than smaller fragments with the same area

(Junninen and Komonen 2011). The loss of connectivity is particularly detrimental to the richness of red-listed taxa with high substrate specificity and low dispersal abilities (Henle et al.

2004, Nordén et al. 2013). Studies have shown that the combined effects of habitat loss and decreased dead wood availability not only reduced the population sizes of saproxylic fungi, but also decreased genetic variation (Högberg and Stenlid 1999, Franzén et al. 2007) and spore viability (Edman et al. 2004a).

For polypores, the effects of local extinctions resulting from forest fragmentation

(Sippola et al. 2004, Berglund and Jonsson 2005) have been found to only become obvious after

>50 years (Gu et al. 2002, Sverdrup-Thygeson and Lindenmayer 2003, Paltto et al. 2006, Penttilä et al. 2006, Ranius et al. 2008). This suggests that an extinction debt is likely to occur in habitat patches that have been recently isolated. In Fennoscandia, the old-growth specialist polypore

Rhodofomes rosea (formerly Fomitopsis rosea) has low dispersal abilities, and spore deposition depends on the area of suitable habitat (forest >140 years old) within 3 km (Larsson 1997,

Edman et al. 2004b). Low dispersal is also evidenced in the genetic differentiation among populations from Sweden, Finland and Russia; the highest genetic variation is found in the more pristine Russian forests (Högberg and Stenlid 1999), suggesting that viability decreases with forest management.

Saproxylic fungi that may benefit from forest management include specialists on logging residues including stumps, slash, treetops and branches (Sullivan et al. 2011), those utilizing smaller-diameter dead wood (Siitonen 2001), and generalist species with high dispersal abilities and fewer substrate restrictions (Berglund et al. 2011, Toivanen et al. 2012, Nordén et al. 2013).

Fomitopsis pinicola is an example of a generalist species with high dispersal ability, and is fully outcrossing over hundreds of kilometers, with low differentiation among populations (Högberg et al. 1995, Nordén 1997, Högberg et al. 1999). This species is therefore less likely to suffer from the effects of forest fragmentation.

However, the removal of post-harvest logging residues and non-commercial tree species for biofuel, which increases disturbance intensity by further reducing the volume of available dead wood (Toivanen et al. 2012), may result in effects even on some of these less sensitive taxa.

Associated wood removal can be dramatic: for example, 65% of logging residues were removed from clearcuts in Sweden (Rudolphi and Gustafsson 2005), and 42% of branches and 81% of cut stumps were removed from clearcuts in Finland (Eräjää et al. 2010). Such forest fuel harvesting may reduce post-harvest DWD below a threshold level for both slash specialist species and generalist taxa able to use residues in addition to larger DWD.

While the effects of management-induced reductions in dead wood supplies are well- known for obligate saproxylic fungi, studies of the impacts on facultative saproxylics (including litter saprotrophs and ECM fungi) in the boreal forest are rare. A study from temperate forests in

Germany examined the effects of forest management type and study site location on soil fungi

(Goldmann et al. 2015). Results showed that forest management influenced the distribution of

ECM fungi, with soil C/N ratio and pH being the main factors in shaping ECM community composition. Certain ECM genera were specific to particular study sites and management types,

16 or to coniferous or beech forest. These results underline the importance of host specificity to

ECM fungi (e.g. Ishida et al. 2007) and the interaction of study site characteristics and the effects of forest management on ECM community composition. Conclusions about species richness as a result of harvesting operations remained unclear, however, in part due to low replication (for each of three different sites, four replicate experimental plots were sampled in each of four management types; Goldmann et al. 2015).

Forest management effects on fungi in Canada

Canada’s boreal forest comprises some 21-27% of the land area of boreal forests worldwide (Brandt 2009), 8% of the world’s forests (Kurz et al. 2013) and 30% of Canada’s land area (Rowe 1972), comprising approximately 340 million ha (Amiro et al. 2001). In contrast to the long history of intensive harvesting in Fennoscandia, the boreal forest zone in Canada has a relatively recent management history (Fischer et al. 2012) and has not been subjected to extensive land-use change (Kurz et al. 2013). Large areas of primary forest still remain, presumably with their biota intact (Bergeron et al. 2002), providing a useful benchmark against which management effects can be measured. As Canada’s boreal forest constitutes some of the largest tracts of unmanaged forest worldwide (Canadian Forest Service 2005) and stores huge amounts of carbon (Kurz et al. 2013), maintaining this ecosystem intact has implications for global biodiversity and carbon dynamics (Stenlid et al. 2008).

Clearcutting remains the primary method of harvesting in Canadian boreal forests

(Brassard and Chen 2006). Although less is known regarding dead wood dynamics in boreal

Canada compared to other forested regions in North America (Pedlar et al. 2002), clearcutting has been shown to reduce the availability of dead wood below levels resulting from fire-related

17 disturbance, potentially for decades post-harvest (McRae et al. 2001, Moroni 2006, Brassard and

Chen 2008, Fischer et al. 2012). Pedlar et al. (2002) showed that clearcuts had less than one-third of the CWD volume of burned stands and mainly consisted of stumps and small logs instead of the burn-produced pulse of snags. Overall, post-harvest woody debris is smaller and more quickly-decaying than that produced from fire (Brassard and Chen 2008, Hagemann et al. 2009), making it more ephemeral and less likely to support the diversity of saproxylic taxa that use larger dead wood from natural disturbances (Bader et al. 1995, Sippola and Renvall 1999).

Additionally, short rotation times compared to average fire return intervals mean that harvested stands are younger than unlogged stands (Cyr et al. 2009, Bouchard and Pothier 2011), with less time to regenerate dead wood resources through stand development processes such as self-thinning (Sturtevant et al. 1997), and resulting in the loss of old-growth forests at the landscape level (Bergeron et al. 2002). This may extend the effects of reduced dead wood availability to entire forested landscapes (Fischer et al. 2012). Although extensive boreal forest management is more recent in Canada than in Europe, effects on dead wood availability in

Canadian forests and reductions in forest age may be on a similar trajectory to those in

Fennoscandia, or accelerated given the possible exacerbating effects of biofuel harvesting

(Riffell et al. 2011).

Just as extensive commercial forest management in boreal Canada is a more recent phenomenon than in Europe, research on the effects of forest management on fungal communities in Canada is also in its infancy, with only a few studies to date. Desponts et al.

(2004) sampled “pristine” senescent and old-growth stands (approximately 90 years old) and silviculturally-mature second-growth stands (approximately 50 years old) in the boreal fir forests of eastern Quebec. They found that greater habitat diversity in old-growth stands resulted in a

18 significantly more diverse community of fungi compared to managed stands. The greater diversity and frequency of CWD-specialist fungi in old-growth stands were associated with the presence of large-diameter CWD from early- to mid-decay stages. Non-saproxylic fungi were also significantly more diverse in old-growth stands, potentially highlighting the importance of

CWD for the entire fungal community.

In northwestern boreal Quebec, Kebli et al. (2012) found that fungal assemblages were influenced by stand composition and downed CWD volume: fungal communities varied among log species, with spruce logs supporting the greatest diversity and number of fungal operational taxonomic units (OTUs). This again highlights the potential impacts of harvesting operations that reduce the availability of conifer CWD (e.g., MacDonald 1995).

Fischer et al. (2012) compared the richness and diversity of fungal fruitbodies and molecular OTUs from spruce logs in old-growth and mature, managed mixedwood boreal forests in northeastern Ontario. They found that log decay class influenced fungal diversity and that the number of fruitbodies on early-decay logs was significantly greater in unlogged versus logged sites, correlating positively with the site-level volume of recently-decayed conifer DWD. Despite relatively large overall volumes of DWD in logged sites, harvesting reduced the volume of early- decay conifer DWD, which was correlated with reduced fungal diversity of sampled logs approximately 30-60 years post-harvest. In addition, early-decay conifer logs from old-growth stands harboured a greater fungal richness than those of equal diameters from managed stands.

Unfortunately, many aspects of the relationships between management, dead wood availability and boreal fungal community composition and richness remain unknown in Canada.

The unique forest type used by Desponts et al. (2004) makes it difficult to apply their results more broadly: the humid climate and forest mosaic resulting from the pattern of natural

19 disturbances (mainly blowdowns) in the Gaspé Peninsula differ from the fire- and insect- disturbed stands characteristic of other parts of boreal Canada (Johnson 1992, Payette 1992,

Bouchard et al. 2006). Kebli et al. (2012) and Fischer et al. (2012) restricted their sampling to specific CWD substrates; fungal communities on other substrates and effects on ECM and litter saprotrophs, to my knowledge, have not been studied in Canadian mixedwood boreal forests.

Of particular interest are effects of controlled manipulations of dead wood supplies on macrofungal communities, which have not been undertaken. As downed trees (logs) have been shown to host higher species richness of polypores than standing dead trees (Junninen and

Komonen 2011), investigating the effects of reduced DWD supplies are particularly important.

Such studies have the advantage in that they can examine the effects of variation in DWD supplies in the absence of other potentially confounding habitat variation. They also allow the manipulation of DWD supplies to lower levels than those currently observed, allowing investigation of possible threshold effects, and emulate DWD reductions expected under additional rotations or under more aggressive fibre removal practices such as biofuel harvesting.

Effects of DWD variation have not been examined for non-wood decay taxa, including the degree to which litter saprotrophs and ECM taxa respond to changes in DWD availability. These groups are able to utilize dead wood or wood-derived compounds resulting from the action of wood saprotrophs and hence variation in DWD resources may also affect the diversity of these ecological guilds.

To examine the effects of forest management and dead wood removal on macrofungi in boreal mixedwood forests of northeastern Ontario, I used area-based surveys to compare 1) the richness and species composition of the macrofungal communities in unlogged (UL) and post- logged stands, including post-logged stands which had undergone an experimental manipulation

20 of downed CWD and 2) the richness of each of four ecological guilds of macrofungi (wood saprotrophic, litter saprotrophic, ectomycorrhizal and parasitic) in the same stands. This research is presented in the next chapter of the thesis.

Chapter 2. Effects of forest management and experimental dead wood removal on macrofungal communities in boreal, mixedwood forests of northeastern Ontario

Introduction

Old-growth boreal forests are functionally and compositionally heterogeneous landscapes

(Kneeshaw and Gauthier 2003) governed primarily by mortality and regeneration gap dynamics

(Hunter and Parker 1993). The old-growth state begins when the original postfire cohort of trees begins to die and the understory stems begin to be recruited to the canopy (Kneeshaw and

Gauthier 2003, Brassard and Chen 2006, Bergeron and Fenton 2012). These processes result in high structural complexity, including a multi-aged tree community including large, old trees, a multi-layered canopy and an accumulation of coarse woody debris (CWD), including both large snags and logs in various stages of decay (Sturtevant et al. 1997, Siitonen et al. 2000, Kouki et al. 2001, Siitonen 2001, Uotila et al. 2001, Kneeshaw and Gauthier 2003 ). The structural heterogeneity of old-growth stands that results from disturbance and succession creates an array of habitats which, in turn, supports a rich biota (Bergeron and Fenton 2012).

Globally, old-growth boreal forests are threatened by forest management (e.g., Siitonen

2001) and, increasingly, by residue extractions for biofuel (Berglund et al. 2011). Harvesting can have many effects, including reductions in the area and connectivity of old-growth stands, simplified stand age and structural complexity, suppression of natural spatiotemporal disturbance dynamics, and decreases in the availability of dead wood, particularly the volume and diversity of CWD with respect to size and species composition (Sousa 1984, Esseen et al. 1997, Östlund et al. 1997, Linder and Östlund 1998, Virkkala and Toivonen 1999, Kouki et al. 2001, Nilsson et al.

2001, Siitonen 2001, Kuuluvainen 2002, Angelstam and Kuuluvainen 2004, Gibb et al. 2005,

Dove and Keeton 2015).

In boreal Europe (Fennoscandia), the negative effects of forest management on biological communities have been well documented (Siitonen 2001). A long history of intensive forest utilization (von Berg 1859, Heikinheimo 1915, Esseen et al. 1997, Östlund et al. 1997, Linder and Östlund 1998), particularly over the past 100-150 years with the development of modern forestry (Östlund 1993), has resulted in the severe loss, fragmentation and alteration of old- growth forest stands, leading to population declines of many species (Rassi et al. 2001, Aronsson et al. 1995, Kuusinen et al. 1995, Ahlén and Tjernberg 1996). Significantly, forest management has reduced the volume of CWD by up to 90-98% at the landscape level (Siitonen 2001), not only through clearcutting, but also through stand thinning, fire prevention and salvation logging

(Siitonen et al. 2000). Large-diameter, late-decay CWD has been most highly reduced (Siitonen

2001) due to harvest-associated removal of large-diameter trunks (Siitonen et al. 2000) and the short rotation times typical of modern forest management (Hansen et al. 1991). While most of the CWD volume in old-growth stands consists of larger diameter classes and an abundance of logs in intermediate- and late-decay stages, managed stands are skewed towards smaller diameter classes in earlier decay stages (Fridman and Walheim 2000, Siitonen et al. 2000, Edman and

Jonsson 2001, Fraver et al. 2002, Pedlar et al. 2002).

Reductions in CWD availability have been shown to be a primary factor threatening many saproxylic (wood-using) taxa (Esseen et al. 1997, Siitonen 2001, Penttilä et al. 2006) and have contributed to an average 50% reduction in richness of saproxylic species in mature managed versus old-growth stands (range 18-75%; Gustafsson and Hallingbäck 1988, Andersson and Hytteborn 1991, Økland 1994, Siitonen 1994, Lindblad 1998, Martikainen et al. 1999, 2000,

Sippola et al. 2001). Saproxylic species richness has been shown to correlate positively with

CWD volume and the number of logs per site (Bader et al. 1995, Økland et al. 1995, Økland

1996, Martikainen et al. 1999, 2000), while richness per stem increases with diameter and decay stage, until mid-decay stages (Bader et al. 1995, Renvall 1995, Høiland and Bendiksen 1997,

Lindblad 1998, Söderström 1988a, Kruys et al. 1999). Large-diameter logs have been shown to host more specialist and locally and regionally rare species than smaller logs (Gustafsson and

Hallingbäck 1988, Söderström 1988a, Kruys et al. 1999, Sippola et al. 2001). Forest management can also affect the species assemblages in CWD, as community composition is influenced by tree species, diameter and decay stage (Harmon et al. 1986, Bader et al. 1995,

Renvall 1995, Sverdrup-Thygeson and Midtgaard 1998, Virkkala and Toivonen 1999).

Fungi comprise a species-rich group of organisms in boreal forests that are particularly sensitive to the effects of forest management (Siitonen 2001, Niemelä 2005, Berglund et al.

2011). As major drivers of ecosystem processes, fungi are integral to nutrient cycling, decomposition, and biodiversity, and therefore to post-harvest succession, forest regeneration and ecosystem resiliency (Hättenschwiler et al. 2005, Boddy et al. 2008, Lonsdale et al. 2008,

Gessner et al. 2010). The degree to which fungi may be affected by reductions in dead wood supplies due to forest management is in part governed by their nutritional mode. Four main ecological guilds of macrofungi can be distinguished in boreal forests, with varying affinities to dead wood substrates: wood saprotrophic, litter saprotrophic, ectomycorrhizal (ECM) and parasitic fungi. Although the term ‘saproxylic’ has mainly been confined to wood saprotrophs

(e.g., Siitonen 2001), all four guilds may be described as saproxylic as they rely for at least some portion of their life cycle on dead wood (Speight 1989).

Wood saprotrophs (many in the order Polyporales) require a dead wood substrate throughout their life cycle (Sippola et al. 2004), and are therefore particularly sensitive to post- harvest reductions in dead wood supplies (Nitare 2000, Niemelä 2005). Specialist taxa that

24 require specific dead wood types, decay communities, microclimatic conditions and spatiotemporal continuity of dead wood resources are particularly susceptible to management- induced changes in substrate availability (Bader et al. 1995, Niemelä et al. 1995, Renvall 1995,

Kruys et al. 1999, Nordén and Appelqvist 2001, Siitonen et al. 2001, Gu et al. 2002, Penttilä et al. 2006, Boddy et al. 2008, Jönsson et al. 2008). The general absence of suitable substrates and altered environmental conditions in intensively managed forests have led to population declines of many wood-decaying fungi (Gärdenfors 2000, Rassi et al. 2001).

In Fennoscandia, the diversity of polypores has been shown to be affected by the volume, quality and continuity of dead wood substrates (Bader et al. 1995, Lindblad 1998, Penttilä et al.

2004, Sippola et al. 2004, 2005, Hottola and Siitonen 2008, Hottola et al. 2009). These fungi require long-term availability of dead wood at the landscape level (Groven et al. 2002, Sverdrup-

Thygeson and Lindenmayer 2003) and many species are specialized to particular types of dead wood (Niemelä et al. 1995, Renvall 1995, Boddy et al. 2008). The rarity and patchy distribution of large logs (CWD >20 cm diameter) of mid- and late-decay stages in managed stands is especially detrimental to taxa specialized on these substrates, including many red-listed species

(Bader et al. 1995, Renvall 1995, Lindblad 1998, Kruys et al. 1999, Sippola et al. 2001,

Tikkanen et al. 2006, Junninen and Komonen 2011). Rare species that are sensitive to changes in humidity and have limited dispersal abilities are particularly favoured by continuous availability of large CWD (DBH >30 cm), which maintains moist conditions during drought, provides a larger colonization surface area, and is less ephemeral than smaller dead wood (Söderström

1988b, Andersson and Hytteborn 1991, Lesica et al. 1991, Bader et al. 1995, Høiland and

Bendiksen 1996, Lindblad 1998).

Research also indicates that past practices in Fennoscandia have incurred an extinction debt (Berglund and Jonsson 2008), and in the future rare polypore species are likely to become extinct in existing small, old-growth patches (Junninen and Komonen 2011). Because new CWD with a diameter >30 cm may not begin to accumulate until approximately 100 years post- disturbance (Siitonen 2001), large-CWD specialists may not have access to suitable substrates before local populations are reduced or extinct, even if stands are no longer being managed

(Bader et al. 1995). The increasing rarity and loss of connectivity of old-growth stands means that the richness of saproxylic fungi is likely to decrease as a result of reduced population sizes, while the composition of assemblages will likely lose late-decay CWD specialists, including red- listed species, with restricted spore dispersal abilities. Even given sufficient time and accumulation of suitable dead wood resources, mature stands may therefore never reach the richness and diversity of old-growth stands.

In addition to effects on richness, forest management may also impact the succession and community composition of boreal saproxylic fungi within a dead wood substrate (Coates and

Rayner 1985a,b,c, Boddy 2000). While brown-rot fungi are more common in earlier decay stages, white-rot ligninolytic species are more common in late-decay wood when only recalcitrant nutrients are available (Renvall 1995). Reductions in late-decay CWD due to harvesting could therefore reduce the population sizes and richness of white-rot fungi, preventing the release of lignin-locked nutrients into the soil and affecting the functional groups of fungi that rely on them (Durall et al. 1994, Mäkipää et al. 2017). Indeed, many late-successional fungi are reliant on wood decayed by specific fungal predecessors (Niemelä et al. 1995, Stenlid et al.

2008).

While the presence of increased volumes of post-harvest FWD may ameliorate the impact of CWD removal on species richness by serving as refugia for generalist wood-decay taxa and

FWD specialists (Küffer and Senn-Irlet 2005, Küffer et al. 2008, Tikkanen et al. 2009,

Juutilainen et al. 2011), clearcut slash has also been shown to harbour 50% fewer species than

FWD produced in old-growth forests (Allmér 2005).

Although less well studied, forest management may also affect other ecological guilds of fungi in addition to wood saprotrophs, including soil-inhabiting litter saprotrophs and ectomycorrhizal (ECM) taxa. Litter saprotrophs may utilize highly-decayed wood fragments as they become humified as soil organic matter (Mäkipää et al. 2017) and can therefore be viewed as facilitating the final stage of wood incorporation into the soil. They may also colonize wood

(Rayner and Boddy 1988) and needle litter indiscriminately (Allmér et al. 2009), highlighting the importance of dead wood, particularly DWD, availability for these fungi.

Ectomycorrhizal (ECM) fungi mainly inhabit soil, but are often facultative saproxylics

(Tedersoo et al. 2003, Rajala et al. 2012, 2015). Highly-decayed CWD nurse logs function as moist, stable microclimates for developing seedlings, supporting the production of fine roots with ECM root tips (Harvey et al. 1978, 1979, Christy et al. 1982, Vogt et al. 1995, Goodman and Trofymow 1998). Although the ECM state is evolutionarily derived, with reversions to saprotrophy unlikely (Bruns and Shefferson 2004; Tedersoo and Smith 2013), certain ECM fungi retain functional lignin-degrading genes (Chambers et al. 1999, Chen et al. 2001, Nehls et al.

2001, Lindahl and Taylor 2004, Luis et al. 2005, Bödeker et al. 2009) and may utilize lignocellulosic compounds freed from dead wood by the action of wood saprotrophs (Durall et al. 1994) or decompose dead wood directly (Colpaert and Van Laere 1996, Talbot et al. 2008).

The ability of some ECM fungi to utilize wood-derived lignocellulosic compounds as an

27 alternative energy source to the photoassimilates provided by their host plants (Talbot et al.

2008) may be particularly important in developing stands that have limited photosynthetic ability

(Dighton and Mason 1985). Post-harvest reductions in dead wood supplies could therefore impact forest regeneration by limiting the availability of lignin-containing compounds for ECM fungi.

Despite the large body of research in boreal Europe, few studies as yet have examined the effects of forest management on the macrofungal communities of Canada’s boreal forest. The studies that have been undertaken to date suggest that, similar to Fennoscandia, the increased heterogeneity of dead wood substrates in old-growth stands support a higher richness of saproxylic fungi (Desponts et al. 2004), and that management-induced reductions in CWD diameter and decay class correlate with reductions in both fungal richness and diversity (Fischer et al. 2012, Kebli et al. 2012). However, the effects of management on ecological guilds of fungi other than saproxylic taxa have not yet been examined. Such studies are useful in determining the effects of reduced dead wood supplies on fungal communities that may indirectly rely on it

(such as certain litter saprotrophs and ECM taxa), and therefore better assess community-wide effects of harvesting. This is important given that these other ecological guilds may contribute to dead wood decomposition and aid in forest regeneration. Experimental manipulations in woody debris supplies have also not been undertaken, but are of particular value in helping to distinguish causation and correlation. In addition, they can be used to manipulate CWD supplies to levels lower than those currently observed in the landscape. Such studies are of particular importance in Canada, where boreal harvesting is in its first rotation and CWD levels can be expected to be relatively high prior to equilibrating at lower levels following subsequent rotations (Hansen et al. 1991). These studies are relevant in a broader boreal context, as a

28 growing demand for biofuel extraction from logging residues and non-commercial tree species is likely to exacerbate the effects of management-induced decreases in substrate availability for boreal fungi (Berglund et al. 2011, Toivanen et al. 2012).

To examine the effects of forest management on macrofungal communities in boreal northeastern Ontario, in this thesis I compared 1) community composition and morphospecies richness and 2) morphospecies richness in ecological guilds between unlogged, old-growth stands and mature, post-logged stands, including post-logged stands that had undergone a large- scale experimental manipulation of DWD supplies. I focused on the oldest post-logged stands in the landscape, as these are closest to what may constitute the oldest stands in future harvested landscapes (see Cyr et al. 2009) and ask the question, has sufficient time elapsed since harvesting for fungal communities to return to those typical of old-growth forests? I used area-based surveys to examine the entire aboveground macrofungal community to determine effects on taxa that utilize dead wood as well as soil macrofungi that may be indirectly impacted by decreases in

DWD availability. Examination of species richness within ecological guilds (ectomycorrhizal, wood saprotrophic, litter saprotrophic and parasitic) is of particular value because it can provide insight into functional ecosystem changes, particularly given the interconnectedness of the various guilds of boreal fungi (see Tedersoo et al. 2003, Lindahl et al. 2006, Koide et al. 2008,

Allmér et al. 2009, Courty et al. 2010).

Based on the literature, I predicted that old-growth sites would support higher richness and more rare species than managed sites. I also expected assemblages of fungi to differ based on stand type: in old-growth stands I expected a higher frequency of species associated with tree senescence and CWD, whereas in logged stands I expected more generalist saproxylic taxa associated with self-thinning (Desponts et al. 2004). I expected differences between old growth

29 and managed stands to be most extreme in those managed stands from which most CWD had been experimentally removed.

Methods

Study sites

Research was conducted in the Gordon Cosens Forest Management Unit within 80 km of

Kapuskasing, northeastern Ontario, Canada (49°25’0” N, 82°26’0” W). Kapuskasing is located at an elevation of 218 m (Henry 2008) in the Clay Belt of the Canadian Boreal Shield Ecozone.

This region is typified by clay deposits (Vincent and Hardy 1977) and consists of gleysol and luvisol soils (Soil Classification Working Group 1998) as well as organic soils (Lafleur et al.

2016). It has an average annual temperature of 1.3°C, with 100 frost free days, and an average annual precipitation of 830 mm, 30% of which falls during the growing season (Lafleur et al.

2016).

All sites were located in closed-canopy, boreal mixedwood forest stands; that is, those composed of a mix of both coniferous and deciduous tree species. Based on the Forest Resource

Inventory (Ontario Ministry of Natural Resources, unpublished), stand canopy cover composition ranged from 30-70% poplar (trembling aspen, Populus tremuloides and balsam poplar, P. balsamifera; mean 44%) and 10-50% spruce (black spruce, Picea mariana and white spruce, P. glauca; mean 23%). Additional common species included balsam fir (Abies balsamea; mean 20%) and white birch (Betula papyrifera; mean 10%), whereas uncommon species included white cedar (Thuja occidentalis) and jack pine (Pinus banksiana) (mean <2%).

Study sites were established in two stand types: unlogged (UL), old-growth stands disturbed by fire in the mid- and late-1800s (4 sites) or in 1924 (1 site) and stands harvested via

30 clearcutting either by horses between 1943 and 1959 (5 sites) or by skidders between 1967 and

1975 (4 sites) (Table 1). Nearest neighbour distances between sites averaged 7 km (range 1-36 km). Within each site, two or three 150 × 150-m (2.25 ha) plots were established that served as the unit of replication. Plots within a site were spaced at least 150 m from each other and located at least 100 m from roads. In each unlogged site, two plots were established, for a total of 10 plots. In each of the managed sites, three plots were established for a total of 27 plots. In the centre of each plot, a 120 × 120-m plot was established that served as the focus of sampling.

Coarse downed woody debris manipulation experiment

In August-October of 2006 and 2007, an experimental manipulation of CWD (downed wood only) was undertaken at the post-logged sites. Among the three plots established per site, three treatments were assigned at random: 1) control (CO) plots in which no CWD was removed; half-removal (HR) plots in which half the volume of CWD was removed; and 3) full-removal

(FR) plots in which all CWD was removed in 2006 or 2007 and every two years thereafter

(Table 1).

In each plot, three parallel skid trails were created to facilitate CWD removal; these were spaced approximately 50 m apart (allowing for skid distances <25 m) and were established away from low-lying areas and with as little tree cutting as possible, to minimize site damage. Wood removals took place using a cable skidder and were piled >75 m away from plot edges. At the time of the initial manipulation, wood that was too decayed to skid was broken into small pieces with chainsaws; these were eventually removed from the FR plots four years after the initial removal (i.e., in 2010 and 2011). To mimic skidder activity, skid trails in CO plots were lightly bladed.

Analyses by Piascik (2013) indicated that the removal experiment was successful in reducing wood volumes as planned and in disrupting pre-existing patterns of variation.

Specifically, in CO plots, pre- and post-manipulation volumes of total CWD remained relatively constant (49 and 45 m3 ha-1 pre- and post-manipulation, respectively) and before-and-after measurements were significantly correlated (R2 = 84%, p = 0.0004). By contrast, in HR plots the total CWD volume was reduced on average by 45% (from 61 to 33 m3 ha-1 on average) and in

FR plots on average by 81% (67 to 13 m3 ha-1 on average). As a result of this reduction, before- and-after CWD measurements across all sites were not significantly correlated, even when including the highly-correlated measurements in control plots (R2 = 6%, p = 0.22, n = 27 plots;

Piascik 2013).

Downed woody debris sampling

Variation in the quantity and quality of CWD was expected to be a main correlate of fungal community variation, hence detailed measurements of CWD (>7 cm diameter) were undertaken in the unlogged sites in 2005 and 2006 and in the managed sites in 2010 and 2011

(following the manipulation). CWD was sampled using the line-intersect method (Van Wagner

1968). In each 120-by-120 m plot, seven, 120-m-long transects spaced at 15-m intervals were sampled in both the x and y dimensions, for a total of 1680 m of line-intersect per plot. For each piece of CWD intercepted, the diameter, decay stage (“early” or “late”) and species were recorded. Decay class groupings correspond roughly to decomposition classes I-III (early-decay) and IV-V (late-decay) in Maser et al. (1979). Early-decay wood had firm-to-soft outer layers but was largely intact and could not be kicked apart; late-decay wood had soft, considerably decayed or missing outer layers and could be relatively easily kicked apart. When CWD could not be

32 identified as coniferous or deciduous, it was assumed to be distributed between those groups in the same ratio obtained for identified pieces of the same size and decay class (Piascik 2013).

Following Piascik (2013), I calculated nine CWD variables to summarize inter-plot variation in CWD supplies. The nine variables were volumes of: 1) all CWD, 2) small-diameter, early-decay coniferous, 3) small-diameter, late-decay coniferous, 4) large-diameter, early-decay coniferous, 5) large-diameter, late-decay coniferous, 6) small-diameter, early-decay deciduous,

7) small-diameter, late-decay deciduous, 8) large-diameter, early-decay deciduous, and 9) large- diameter, late-decay deciduous. Small-diameter CWD was that below the overall median diameter of CWD (12 cm); large diameter CWD was equal to or above the median diameter.

Additional habitat variables

I additionally measured other key habitat variables that could be expected to influence fungal community variation: percentage basal area of deciduous trees, shrub/sapling stem density, and two primary environmental gradients as revealed by an ordination of understory shrub/sapling communities (see below). These variables were measured in 2006-2007 and reflected fundamental site characteristics that were unlikely to have changed over the short time frame between their sampling and the present study (summer 2013). A central 90 × 90 m (15-m spacing) grid in each plot served as a reference for tree and shrub sampling. Basal areas of live trees ≥7 cm in diameter at breast height were sampled via BAF 2 prism sweeps at every second plot station (16 locations per plot). To sample shrubs and saplings (≥1.0 m in height and <7 cm dbh), a 21 × 21 plot with 5-m spacing was superimposed on the 15-m plot, extending 15-m beyond it on all sides. Stem counts (at 1 m in height) by species were taken in 1 m2 areas at the

441 plot intersections.

To define primary ecological gradients in the study sites, I used plot scores from the first two axes of a Detrended Correspondence Analysis (DCA) undertaken on plot-specific total counts of each shrub/sapling species (see Vanderwel et al. 2010). The first axis (DCA1; 31.3% of total inertia) represented a gradient in light exposure and conifer composition, with shade- intolerant deciduous species such as golden rod, trembling aspen, and wild red raspberry at the low end of the axis and shade tolerant coniferous species such as black spruce, balsam fir, and eastern cedar at the high end. The second axis (DCA2; 14.8% of total inertia) represented a gradient in soil moisture, with indicators of hydric conditions such as speckled alder and alder- leaved buckthorn at the low end of the axis and indicators of mesic, fresh conditions such as beaked hazel and mountain maple at the high end.

Fruiting body sampling

I was interested in large-scale, area-based sampling across substrates, hence I used area- based fruiting body sampling rather than molecular techniques. Fruiting body sampling has been useful in examining community-wide effects of harvesting in other studies (eg. Diamond and

May 1976, Pimm and Askins 1995, Rosenzweig 1995, Hanski 2000), whereas molecular techniques provide a more localized, substrate-specific view of a fungal community, although they have the benefit of detecting non-fruiting species or those that fruit infrequently (Straatsma et al. 2001). As fungal communities sampled with molecular techniques have been shown to be more stable at the level of forest stand, compared to higher variability at finer scales (Izzo et al.

2005, Koide et al. 2007), fruiting body surveys could serve as a proxy for molecular sampling methods at larger scales, at least for dominant fungal taxa.

Fruitbodies of macroscopic fungi >0.5 cm in diameter and situated up to 2 m from the ground (on snags and live trees) were collected in all plots during three sampling periods in the summer of 2013. Sampling periods were on average 8 days in length (range 6-10 days) and were distributed evenly throughout the summer (sampling period 1: June 20-31, sampling period 2:

July 26-31, sampling period 3: August 19-28).

The order in which sites were surveyed was randomized per sampling period. To minimize possible effects of temporal variation, the two or three plots per site were surveyed as synchronously as possible. In sampling period 1, all plots per site were sampled on the same day

(n = 14 sites); in sampling period 2, all plots per site were sampled on the same day (n = 11 sites) or over two days (n = 3 sites); and in sampling period 3 all plots per site were sampled on the same day (n = 8 sites) or over 2-4 days (n = 6 sites).

In sampling period 1, two, 90 × 2-m transects (each with six 15 × 2-m segments) were sampled per plot (that is, 360 m2 in total per plot). Large numbers of fruiting bodies in sampling periods two and three required that smaller areas be sampled; I therefore reduced the sampling area per plot to two 15 × 2-m segments per 90 × 2-m transect, chosen at random (that is, 120 m2 in total per plot per sampling period). The areas sampled were distributed such that any given area in a plot was surveyed in at most one sampling period.

Within a sampling period, the pair of transects surveyed per plot ran parallel to each other. In sampling periods 1 and 2, transects were spaced 30-m apart, and in sampling period 3,

60-m apart. Orientation and spacing of transects were standardized such that the majority of sampling was not on skid trails. Sampling was undertaken by running a line lengthwise along the centre of the transect segment, then slowly searching a 1-m swath on each side of the line for fungi, using a metre stick held perpendicular to the line to confirm distance.

Each fruiting body species unique to a transect was collected and its transect segment, plot, and substrate recorded. Collected fruiting bodies were photographed, dried and grouped into morphospecies based on macro- and microscopic characteristics using a number of online and literature sources. The extensive online database of the Quebec Mycological Society

(www.mycoquebec.org) was a principal resource for assigning species names, due to the proximity and similarity in environment between Quebec and the present study sites in northeastern Ontario.

DNA extraction and ITS sequencing

To confirm fungal species designations, the internal transcribed spacer (ITS) gene region was sequenced for one or several representative specimens of a subset of the putative morphospecies. Due to logistical constraints, I was unable to sequence all of the specimens I collected. Instead, I sequenced morphospecies whose identification seemed most likely to be problematic, although a relatively small number of these proved impossible to successfully sequence. As a result of not being able to sequence every specimen, I use the term

‘morphospecies’ throughout.

The ITS region is a component of the nuclear ribosomal DNA (rDNA) and comprises the highly-variable ITS1 and ITS2 spacer regions as well as the conserved 5.8S gene; it is recognized as the “universal” barcode sequence for species-level identification in fungi (Schoch et al. 2012) due to its ease of amplification (it is both multicopy and contains conserved priming sites beside highly variable regions; Peay et al. 2008), and its degree of variability that corresponds well to interspecific differentiation (Schoch et al. 2012) and morphologically- defined species (Smith et al. 2007) in most groups of fungi.

DNA was extracted from dried fruiting body tissue using the protocol of Dentinger et al.

(2010). From the extracted DNA, the ITS region was amplified via polymerase chain reaction

(PCR) using the forward and reverse primers ITS-1F (Gardes and Bruns 1993) and ITS4 (White et al. 1990), respectively. PCR reactions were performed using a 10 µL reaction volume

(comprising 0.1 units Platinum Taq DNA polymerase, 0.2 units each ITS1F (10µM) and ITS4

(10µM), 1.6 units dNTPs (1.25mM), 1 unit 10X EH buffer, 4.9 units water and 2 units DNA template). PCR amplification was performed using the following cycling program: initial denaturation at 94°C (2 min), followed by 35 cycles of denaturation at 94°C (50 s), annealing at

55°C (50 s), and extension at 72°C (1 min); cycles were terminated with a final extension at

72°C (7 min) and indefinite refrigeration at 4°C.

Positive PCR products were visualized and cleaned using gel electrophoresis: 5 µL loading dye was added to each PCR product, and the solution run in a 1% agarose gel containing

0.005% ethidium bromide for approximately 1 hour at 100 V. Positive bands (i.e., those with successfully-amplified fungal ITS DNA) were detected under UV fluorescence and excised from the gel as described in Dentinger et al. (2011); each band was cut from the gel with a sterile razor, placed in a filter-bottom disposable pipette tip previously trimmed to ~1 cm in length, and the pipette tip then placed in a 1.5 mL microfuge tube. Tubes were spun in a centrifuge for 10 min at 10 000 g and the resulting liquid (containing the amplified DNA) was collected and run through a second round of PCR. Cleaned amplicons were ethanol-precipitated and resuspended in HiDi formamide prior to unidirectional dye-terminator sequencing using the ABI BigDye kit

(Foster City, CA). Produced sequences were then run on an ABI PRISM 3100 DNA Analyzer

(Department of Natural History, Royal Ontario Museum, Toronto, ON).

Chromatograms of the ITS sequences were manually edited for ambiguous reads using

Sequencher 2.1 (GeneCodes, Ann Arbor, MI) and trimmed to begin and end with the conserved motifs flanking the ITS-1F and ITS4 primers: 5′-(...gat)CATTA and GACCT(caaa...)-3′, respectively (Dentinger et al. 2011). Forward and reverse sequence pairs were merged into contigs using the “Assemble Automatically” (algorithm) with “Minimum Match Percentage” set at 90 (or lower for sequences that did not align as well), “Minimum Overlap” set to 100 bp and contig consensus type set to “Consensus Inclusively”.

BLAST sequence alignment and phylogenetic analysis

Consensus sequences were exported and compared to reference database sequences from the National Center for Biotechnology Information (NCBI; Benson et al. 2009) and UNITE

(Kõljalg et al. 2005) using the blastn (nucleotide BLAST) alignment algorithm (Altschul et al.

1997). Alignments were optimized for highly similar sequences (megablast), and sequences from uncultured and environmental samples were excluded, as they were rarely assigned to genus or species level. The high intraspecific variation of ITS sequences has led previous authors to establish a cut-off of >97% similarity (as in bacterial species designations) between a sample and reference sequence to consider it conspecific; however, in this study I used an intraspecific cutoff of >98%, as other sources have stated that this more stringent approach is warranted, although not universally applicable (Nilsson et al. 2008).

BLAST reference sequences producing significant alignments to the query sequence were sorted by Maximum Score and the top 20 with >60% query cover, >90% identity, and an e-value of 0.0 were selected to construct a maximum-likelihood (ML) phylogenetic tree. In cases with more than 20 alignments with >98-99% identity to the query sequence, all subject sequences

38 with >98-100% (>80% query cover, e-value 0.0) were selected, and the two sequences with lowest maximum score were also selected, to create an outgroup.

If the query sequence was placed as the outgroup in the ML tree (in cases with 20 alignments >90% identity or with >20 alignments with >98-99% identity), then the two BLAST sequences with lowest maximum score for that particular query sequence were chosen as an outgroup, and the tree re-rooted. For query sequences with fewer than 20 top hits with identity

>90%, up to 20 top hits with >80% identity (>60% query cover, e-value 0.0) were selected, with two additional below (e-value <0.0) as the outgroup. If identities were all <90% or e-values all

<0.0, the top 20 with >60% query cover and >80% identity were used to build a rough tree.

Sequences were excluded if they were chimeric, clearly misidentified, or described only to phylum, class, or order.

Each ITS query sequence was aligned with the 20 (or more) top BLAST hits obtained using MEGA 6.0 software (Tamura et al. 2013); alignments were manually trimmed and re- aligned. Each query sequence and its aligned subject sequences were used to construct a maximum-likelihood phylogenetic tree using default settings, with 1000 bootstrap replications to confirm each morphospecies taxon. Species epithets were confirmed in the presence of high

(>98%) identity to the query sequence, which generally corresponded to >80% bootstrap support, but were otherwise preceded by ‘cf.’ or ‘aff.’ to indicate taxonomic uncertainty. Trees were rooted and rotated if needed using the FigTree (v1.4.3) software (Rambaut 2016).

Sequences with the same top BLAST hit (either the same reference sequence or a different, but infraspecific, sequence) were treated as a single morphospecies (due to being macro- and microscopically indistinguishable) and were designated by numbers following the morphospecies name (e.g., Gymnopus confluens 1, Gymnopus confluens 2); in these cases, the

39 top BLAST hits were combined to create one maximum-likelihood tree per morphospecies. For samples that failed to sequence or produced sequences with no significant BLAST results

(according to the above criteria), species designations were based only on macro- and microscopic characteristics.

Statistical analyses

The original morphospecies presence-absence data per transect were converted into a matrix of species presence-absence per plot and sampling period; species were then grouped into four ecological guilds (wood saprotrophic, litter saprotrophic, ECM and parasitic) and the total number of morphospecies per guild per plot and sampling period was made into a second matrix.

The primary source for guild assignment was the database of the Quebec Mycological Society

(www.mycoquebec.org). Litter saprotrophs were defined as those species that utilized humus and highly-decayed woody debris within the humus layer, whereas wood saprotrophs were those species that colonized individual pieces of dead wood that were not highly integrated with the soil layer. Some taxa were assigned to more than one ecological guild; for example, a species utilizing both logs and humus as a substrate would be considered both a wood and litter saprotroph.

Analyses comparing richness and community composition between unlogged (UL) and logged (CO, HR and FR) plots were conducted separately for the two matrices (i.e., morphospecies presence/absence and guild species richness). Because of the strong change in community composition from one sampling period to the next, sampling periods were analyzed separately.

Multivariate analyses using Canoco (v. 4.5) were conducted to determine the effects of management history and habitat variables on the assemblages of macrofungi. Unconstrained ordination using Detrended Correspondence Analysis (DCA) (Lepš and Šmilauer 2003, Ter

Braak and Šmilauer 2002) was first performed on the species matrix for each sampling period to determine whether a unimodal or linear response model best approximated the composition of the sampled fungal community. As the gradient lengths of the first DCA axes were >3 standard deviations for all sampling periods (3.9, 4.0 and 3.7 for sampling periods 1, 2 and 3, respectively), I used a unimodal model. Constrained ordinations were performed on the species and habitat variable matrices using Canonical Correspondence Analysis (CCA) and on the guild richness and habitat variable matrices using Redundancy analysis (RDA; constrained ordination using a linear response model) (Ter Braak and Šmilauer 2002, Lepš and Šmilauer 2003).

For both analyses, Monte Carlo permutation tests (with 9999 permutations) were performed to determine the significance of the first and combined ordination axes for each sampling period. Monte Carlo permutation tests (9999 permutations) were also performed on each of nine CWD and four habitat variables to determine their individual significance as correlates of community variation in each sampling period.

Rarefaction analyses were conducted to compare species richness among UL, CO, HR and FR plots while controlling for sampling effort; the x-axis represented the number of samples and the y-axis represented the cumulative number of morphospecies.

Finally, analysis of variance (ANOVA) was used to test whether management history

(UL, CO, HR and FR) had a significant effect on the number of morphospecies of macrofungi in each of four ecological guilds (wood saprotrophic, litter saprotrophic, ECM and parasitic).

Tukey’s HSD post-hoc test was used as a pairwise comparison of treatments. If the proportion of

41 plots with no species was >20% for a guild in a given sampling period, a median test was performed as a nonparametric counterpart to ANOVA. These analyses were performed using

SAS v. 9.4.

Results

Molecular identification

In total, 541 morphospecies were assigned, of which 263 were sequenced (Appendix I).

Of these 263, 217 aligned with >98% sequence similarity to a reference sequence from the NCBI or UNITE databases and could therefore reliably be used to construct maximum-likelihood phylogenetic trees to confirm morphospecies identities. Forty-six sequences aligned with 80-

97% sequence similarity to a reference sequence, and therefore species designations are less definite. Of the 263 sequenced morphospecies, 74 reference sequences were from specimens collected in boreal Europe (Estonia, Finland, Greenland, Iceland, The Netherlands, Norway,

Sweden) and 24 reference sequences were from specimens collected in eastern Canada

(Newfoundland and Labrador, Nova Scotia, Ontario, Quebec). Cortinarius was the genus with the largest number of morphospecies corresponding to unique reference sequences from boreal

Europe (20). Of the remaining 278 morphospecies that were not sequenced, 132 were identified with certainty using macroscopic and microscopic characteristics, while 146 either failed to sequence (29) or were non-sequenced specimens that were re-examined and determined to be additional morphospecies (117).

Fruiting body sampling

In total, 541 morphospecies (3781 occurrences) from 205 genera (Appendix II) were collected across sampling periods; 142 (564 occurrences) were collected in sampling period 1,

253 (1207 occurrences) in sampling period 2, and 465 (2010 occurrences) in sampling period 3.

Of the total number of morphospecies encountered across sampling periods, 399, 288, and 76 had zero occurrences in sampling periods 1, 2 and 3, respectively, and none were present in all plots.

In total, 45% (246) of the morphospecies sampled were represented by only a single occurrence. In each sampling period, between 35-40% of the taxa were encountered only once

(50 of 142, 102 of 253 and 168 of 465 morphospecies in sampling periods 1, 2 and 3, respectively). Only a small number of morphospecies occurred >10 times (17, 33, and 57 species in sampling periods 1, 2, and 3, respectively). Only 82 morphospecies were present in all three sampling periods, whereas 27, 33 and 243 taxa were unique to sampling periods 1, 2 and 3, respectively. Fifteen morphospecies were shared between sampling periods 1 and 2; 18 were shared between sampling periods 1 and 3; and 123 were shared between sampling periods 2 and

The majority of taxa collected were members of the Basidiomycota (485 morphospecies);

Ascomycota (56 spp.) comprised the remainder. Of the 205 genera, 128 were represented by a single morphospecies. Wood saprotrophs were the most morphospecies-rich ecological guild

(231 morphospecies), followed by ectomycorrhizal taxa (ECM; 180 morphospecies), litter saprotrophs (162 morphospecies), and parasites (27 morphospecies). Of the wood saprotrophs,

40 were also considered to be litter saprotrophs, and 12 were also considered parasitic; a small number of other taxa overlapped in ecological group (4 ECM-wood saprotrophs, 3 ECM-litter

43 saprotrophs, 1 ECM-parasitic, 1 litter-saprotrophic-parasitic and 1 litter-wood saprotrophic- parasitic). Morphospecies with >10 occurrences across sampling periods comprised 37 wood saprotrophs, 32 litter saprotrophs, 27 ECM taxa, 12 litter-wood saprotrophs, 4 wood saprotroph- parasites, 1 ECM-wood saprotroph and 1 parasite.

The most morphospecies-rich genera included the ECM genera Cortinarius (58 spp.),

Russula (24 spp.), Inocybe (20 spp.) and Lactarius (15 spp.) and the litter and wood saprotrophic genera Mycena (34 spp.), Entoloma (22 spp.), Gymnopus (11 spp.) and Crepidotus (10 spp.). The remaining 197 genera each had 9 or fewer morphospecies. The most frequently-encountered morphospecies overall was the wood saprotroph Hymenochaetopsis tabacina, with 61 occurrences across plots and sampling periods. On a per sampling period basis, the morphospecies with the highest number of occurrences (in parentheses) across plots was the wood saprotroph Jackrogersella multiformis (21) in sampling period 1, the litter saprotroph

Hemimycena aff. gracilis (29) and the wood and litter saprotroph Mycena leptocephala (29) in sampling period 2, and the litter saprotroph Gymnopus confluens (30) in sampling period 3.

Across the unlogged plots, the most common morphospecies (number of occurrences in parentheses) was the wood saprotroph Hymenochaetopsis tabacina (26); per sampling period, the most frequently-encountered morphospecies in unlogged plots were Hymenochaetopsis tabacina in sampling period 1 (8) and sampling period 2 (10) and Cortinarius cf. decipiens in sampling period 3 (10). Across logged plots, the most common morphospecies (number of occurrences in parentheses) was the wood saprotroph Cerioporus varius (41); per sampling period, the most frequently-encountered morphospecies in logged plots was Trichaptum fuscoviolaceum in sampling period 1 (39); Mycena leptocephala in sampling period 2 (22); and Gymnopus confluens in sampling period 3 (22).

Macrofungal community variation

Canonical Correspondence Analysis showed a relatively strong distinction between logged and unlogged plots in all three sampling periods (Fig. 1). These differences were associated with significantly higher wood volumes for at least one CWD variable in unlogged compared to logged plots. Generally, unlogged plots were characterized by a high total volume of CWD, high volumes of large-diameter coniferous CWD (in both early- and late-decay stages), and a high volume of large-diameter, late-decay deciduous CWD. In contrast, logged plots contained higher volumes of small CWD (both coniferous and deciduous, in early and late stages of decay). A distinct fungal community was associated with an unusually high volume of large- diameter, early-decay deciduous CWD and high percent basal area of deciduous trees in two plots (ML10CO and ML10HR). Environmental variation as represented by the nine CWD and four habitat variables explained between 24-30% of the total variation in the fungal community

(29.5%, 23.9% and 25.9% for sampling periods 1, 2 and 3, respectively). The influence of these environmental variables was reflected by the highly significant constraint observed for all combined canonical axes in sampling periods 1 and 2 (p<0.0005; Table 2).

The primary CCA axis was significant in sampling period 1 (p<0.001) and explained approximately 4-5% of the total variation in fungal community composition (5.4%, 4.1% and

4.1% for sampling periods 1, 2 and 3, respectively). This axis represented a gradient of the volume of large-diameter, early-decay deciduous CWD (sampling period 1: p<0.0005, sampling period 2: p<0.05), percent basal area of deciduous trees (sampling period 1: p<0.001, sampling period 2: p<0.05, sampling period 3: p<0.05) and the volume of small-diameter, late-decay deciduous CWD (sampling period 1: p<0.005). These variables were positively correlated with each other and distinguished a fungal community associated with high values of these variables

45 in two logged plots (ML10CO and ML10HR). The second CCA axis explained approximately 4-

5% of the total variation in community composition (4.5%, 3.9% and 3.6% for sampling periods

1, 2 and 3, respectively) and separated fungal communities of small CWD in logged plots from those of large CWD in unlogged plots. Wood-removal treatments (CO, HR and FR) within logged sites had no clear effect on their macrofungal communities and did not strongly correlate with any environmental variable.

Permutation tests revealed that the macrofungal communities of unlogged plots were significantly associated with a high total volume of CWD (sampling period 1: p<0.0005, sampling period 3: p<0.005) and a high volume of large-diameter, late-decay deciduous CWD

(sampling period 1: p<0.005, sampling period 2: p<0.001), the values of which were highly positively correlated. Macrofungal assemblages in unlogged plots were also significantly shaped by a high volume of large-diameter coniferous CWD in early- (sampling period 1: p<0.05) and late-decay stages (sampling period 1: p<0.05) and with shade-tolerant and mesic conditions represented, respectively, by the high ends of axes DCA1 (sampling period 1: p<0.005, sampling period 2: p<0.05, sampling period 3: p<0.0005) and DCA2 (sampling period 1: p<0.01).

Rarefaction

Species-accumulation curves indicated that in all three sampling periods, unlogged (UL) plots were more species-rich than control (CO), half-removal (HR) and full-removal (FR) logged plots (Fig. 2). In sampling period 1, UL plots were significantly more species-rich than FR plots: at 9 samples, the number of species in UL plots was 91, whereas FR plots had a total of 57 species (CO and HR plots had a total of 82 and 73 species, respectively). In sampling periods 2 and 3, UL plots were still more species-rich than logged plots, but not significantly so: at 9

46 samples, the total number of species in UL, CO, HR and FR plots in sampling period 2 were 145,

125, 139 and 120, respectively and in sampling period 3 were 272, 226, 230 and 243, respectively. The total number of species tripled from sampling period 1 to sampling period 3 for

UL, HR and FR plots and more than doubled for CO plots.

Richness of ecological guilds

Over all three sampling periods, the number of wood saprotrophs, litter saprotrophs, and the total number of species averaged higher in UL plots than in any of the logged plots, with UL plots consistently more species-rich than CO, HR and FR plots (Table 3, Fig. 3). This was not true of the richness of ECM or parasitic macrofungi in any of the sampling periods.

Redundancy analysis on the ecological guild and habitat matrices revealed that in all sampling periods, there was no significant constraint on community composition observed for either the first or all combined canonical axes. However, environmental variation as represented by the nine CWD and four habitat variables explained between 41-48% of the total variation in ecological guilds (44.0%, 47.7% and 40.7% for sampling periods 1, 2 and 3, respectively).

In all three sampling periods, the number of wood saprotrophs was strongly associated with unlogged plots (Fig. 4). In sampling periods 1 and 3, unlogged plots were also associated with the total number of morphospecies, the number of litter saprotrophs, the total volume of

CWD (p<0.005, p<0.05), the volumes of large-diameter early- (p<0.05, p<0.0005) and late- decay coniferous CWD (p<0.05, p<0.005), and the volume of large-diameter, late-decay deciduous CWD (p<0.005, n.s.). In sampling period 2, unlogged plots were again highly- associated with the volumes of large-diameter coniferous CWD and DCA1, although ecological guilds were influenced significantly by the percent basal area of deciduous trees (p<0.005) and

47 the volume of large-diameter, early-decay deciduous CWD (p<0.05), which negatively correlated with the number of ECM and litter saprotroph morphospecies.

Results from ANOVA showed that in all three sampling periods, the number of wood saprotrophs differed significantly amongst treatment plots (p = 0.046, 0.041 and 0.005 for sampling periods 1, 2 and 3, respectively) and explained over 20% of the variation amongst plots

(R2 = 0.21, 0.22 and 0.32, respectively; Table 3). Pairwise differences between plots showed that in sampling period 2, UL plots were significantly more species-rich than FR plots (Tukey’s

HSD, a=0.05) and contained, on average, 7 more species; in sampling period 3, UL plots were significantly more species-rich than both HR and FR plots (Tukey’s HSD, a=0.05) and contained, on average, 10 and 11 more species, respectively. Pairwise differences between UL and logged plots were not significant at a=0.05 in sampling period 1, despite an overall significant ANOVA result.

The number of litter saprotrophs differed significantly among plots in sampling periods 1

(p = 0.007) and 3 (p = 0.002) and explained, respectively, 31% and 36% of the variation amongst plots. In sampling period 1, pairwise differences indicated that UL plots were significantly more species-rich than CO plots and FR plots (Tukey’s HSD, a=0.05), with an average of 4 and 5 more species, respectively. In sampling period 3, UL plots were significantly more species-rich than all logged plots (Tukey’s HSD, a=0.05), with an average of 19, 13, and 15 more species than CO, HR and FR plots, respectively.

Plots also differed significantly in the total number of morphospecies in sampling periods

1 (p = 0.0179) and 3 (p = 0.0352) and explained 26% and 23% of the variation amongst plots, respectively. In sampling period 1, pairwise differences indicated that UL plots were significantly more species-rich than FR plots (Tukey’s HSD, a=0.05) and contained an average

48 of 8 more species. In sampling period 3, UL plots were significantly more species-rich than CO plots, with an average of 26 more species.

For ECM species, a median test revealed that the number of species above and below the median were not significantly different among plots in sampling period 1; ANOVA results showed that species richness did not differ significantly amongst plots in sampling periods 2 and

3. Similarly, a median test was not significant for parasitic taxa in sampling periods 1 and 2, and an ANOVA was not significant in sampling period 3.

Discussion

Fruiting body sampling

From the total number of morphospecies and fruiting bodies sampled, species richness increased from sampling period 1 to sampling period 3. This was evident from the large number of morphospecies with zero occurrences in the first two sampling periods compared with sampling period 3 and the large number of species unique to sampling period 3 (although there was substantial overlap in species between sampling periods 2 and 3). These results are consistent with previous studies demonstrating that fungal fruiting peaks between late summer and autumn (Eveling et al. 1990) or in early autumn (Straatsma et al. 2001).

The increasing richness across sampling periods was largely due to the appearance of

ECM taxa, particularly in sampling period 3, corresponding to seasonal changes in tree physiology (Hansen et al. 1997, Waring and Running 1998, Kagawa et al. 2006) that facilitate development of ECM fruiting bodies (Högberg et al. 2010). Higher allocation of photosynthate to below ground has been shown to occur in August relative to June and directly facilitates ECM fruiting body production (Högberg et al. 2010). These results are consistent with other studies on

49 the phenology of ECM fruiting body production that demonstrate a seasonal peak in fruiting from mid-summer to early autumn (Sato et al. 2012).

The large amount of rain in July 2013 (99 mm over 16 days) likely contributed to increased fruiting body production in August of that summer; the association between mean annual precipitation and fungal richness has been previously reported (Tedersoo et al. 2014), as has a correlation between precipitation and fruiting body production (Lodge et al. 2004, Durall et al. 2006). This increased richness was apparent despite the shortened transect lengths in sampling periods 2 and 3. Had the area surveyed in these sampling periods been equivalent to that in sampling period 1, it is likely that the increased richness in the latter two sampling periods would have been even more striking.

Nearly half of the morphospecies surveyed overall and per sampling period were singletons and nearly half of the genera sampled consisted of a single morphospecies. Of the fungi sampled per sampling period, only a small number of morphospecies occurred >10 times.

This taxonomic rarity is consistent with other studies of fungal ecology that report an uneven distribution of morphospecies in which the majority of taxa are rare (Tofts and Orton 1998,

Horton and Bruns 2001, Ferrer and Gilbert 2003, Arnold et al. 2007). In a 21-year study on fruiting body production in Switzerland, rare species of macrofungi were shown to occur in particularly speciose years (Straatsma et al. 2001). The large number of rare species in the present study may similarly reflect the highly-productive season, but emphasizes the need for additional and consecutive years of sampling, as highly-productive seasons may not occur annually (Straatsma et al. 2001). Over a seven-year survey of fungal fruiting bodies in Austria,

2448 records of 886 species were encountered in 13, 1-ha forest and grassland plots (Straatsma and Krisai-Greilhuber 2003). The number of species encountered per year varied from 264 to

525, and the number per hectare therefore varied from approximately 20.3 to 40.4, or 0.002 to

0.004 species per m2, respectively. In the present study, the number of morphospecies per m2 varied from 0.39 to 3.9 across sampling periods, indicating that summer 2013 was particularly productive for fruiting bodies.

The predominance of Basidiomycota among the specimens collected in part reflects the typically larger size of fungi within this phylum compared to Ascomycota, of which fewer species form macroscopic fruitbodies (Watling 1995). The size cutoff for morphospecies in the present study (0.5 cm diameter) no doubt resulted in the exclusion of certain saproxylic and soil- inhabiting fungi. For example, certain microscopic species of Ascomycota are important contributors to the decomposition of dead wood, including soft-rot Phialophora species (Kebli et al. 2012). Certain corticioid (crust-forming) species of Basidiomycota may not have been collected, due to their morphological similarity to microscopic Ascomycota (i.e., molds); these include saproxylic species of Tomentella and Athelia, which include both wood saprotrophic and

ECM taxa (Tedersoo et al. 2003, Kebli et al. 2012).

The logistical constraint of being able to sequence only a subset of the macrofungi collected may also have underestimated the number of species collected, particularly for speciose genera with a large number of morphologically-similar species, such as Cortinarius. Although this would modify species richness counts, it would likely not alter the ecological signals in the present study due to the (general) consistency of ecological guilds within genera, and the ability to macroscopically identify fungi at the generic level. The use of molecular sampling techniques rather than area-based fruiting body surveys also would have altered the composition of macrofungi sampled by capturing taxa that fruit infrequently or not at all (Straatsma et al. 2001).

Additionally, although molecular sampling techniques are inappropriate for area-based surveys

51 due to their localized nature, they would likely have sampled a higher richness of fungi within specific substrates (O’Brien et al. 2005).

Wood saprotrophs comprised the most species-rich ecological guild, represented by a breadth of species-poor genera that has been observed elsewhere in boreal forests (Lõhmus

2011). In contrast, ECM fungi were distributed among a smaller number of more speciose genera, similarly to other boreal forest surveys (Durall et al. 2006). The ECM genus Cortinarius was the most species-rich genus sampled, reflecting the high diversity of ECM fungi in boreal forests (Tedersoo et al. 2014). Of the 58 Cortinarius species encountered, 24 were represented by only a single occurrence and 51 had <10 occurrences, corresponding to the composition of

ECM assemblages with a few common and many rare species (Horton and Bruns 2001, Taylor

2002).

The most frequently-encountered morphospecies across plots and across sampling periods was the wood saprotroph Hymenochaetopsis tabacina; this species was also the most common in UL plots and one of the top five most common species in logged plots. As H. tabacina is confined to dead deciduous wood including snags, logs and fallen branches (Labbé

2019b), it is unlikely to be affected by previous forest management practices that targeted large- diameter coniferous CWD (MacDonald 1995). As it is not confined to CWD of a certain diameter or decay class, it can be supported by both the high volume of large, late-decay deciduous CWD in UL plots and the volumes of deciduous CWD (both large, early-decay and small, early- and late-decay) in logged plots. Its ubiquity across plots may also reflect the highly- productive conditions for fruiting body production during the sampling year; higher similarity in assemblages amongst different locations in productive compared to poor seasons has been previously observed (Mehus 1986). Such seasonal weather conditions conducive to fruiting body

52 production may obscure observed differences between UL and logged plots and hence additional years of sampling are required to more accurately determine differences in macrofungal richness and composition among sites with different management histories (Straatsma et al. 2001).

Although H. tabacina was the most common morphospecies in UL plots in sampling periods 1 and 2, the ECM species Cortinarius cf. decipiens was the most common morphospecies in sampling period 3. This species is mycorrhizal with poplars on moist soil

(Labbé 2019a) corresponding to the mesic conditions of UL plots. The association of this morphospecies with living deciduous trees highlights the mixedwood condition of UL plots, despite their transition to conifer dominance. The increased frequency of C. cf. decipiens in sampling period 3 also highlights the appearance of ECM taxa later in the season (Höberg et al.

2010).

The most common morphospecies in logged plots across sampling periods was the wood saprotroph Cerioporus varius; this species has been described as preferring larger-diameter wood including fallen logs and large stumps of coniferous and deciduous trees (Labbé 2017) or with a preference for small deciduous logs and sticks (Kuo 2015). The abundance of C. varius in logged plots may be due to the abundance of small-diameter CWD and FWD produced from self- thinning (Sturtevant et al. 1997).

In sampling period 1, the most common morphospecies in logged plots was the wood saprotroph Trichaptum fuscoviolaceum. Although this species is restricted to coniferous DWD

(Labbé 2014b), it may be able to subsist on the high volumes of small-diameter coniferous CWD in logged plots. Similarly, the most common species in sampling period 2, Mycena cf. leptocephala, utilizes both dead wood and litter of coniferous and deciduous trees (Labbé 2018).

Gymnopus confluens was the most frequently-encountered species in sampling period 3; its

53 ability to utilize both coniferous and deciduous humus and litter (Labbé 2014a) also means that it is not substrate restricted.

The smaller number of occurrences of common species in UL compared to logged plots may reflect increased fruiting body production in association with forest management, particularly for saprotrophic species (Ohenoja 1988). Alternatively, it may indicate the presence of a more diverse community of macrofungi in UL plots, with each species represented by fewer occurrences. The increased availability of microhabitats in old-growth stands (Siitonen 2001) could provide a more diverse array of niches, but with potentially fewer resources for fruiting body production for any given taxon.

Species-accumulation curves indicate that in all three sampling periods, UL plots were the most species-rich. This pattern has been seen in Fennoscandia, in which mature natural forests harbor 72-100% more species of polypores than mature managed forests (Junninen and

Komonen 2011 and references therein).

Molecular identification

The large number of maximum-similarity reference sequences from boreal Europe, many of which correspond to ECM taxa, reflects the circumboreal distribution of many ECM fungi and their associations with boreal tree species (e.g., Geml et al. 2012). Several of the sequences obtained matched with reference sequences for macrofungi that have currently only been recorded from Fennoscandia or elsewhere in northern Europe and likely represent taxa new to

Ontario. For example, the top match for the specimen of Cortinarius argenteolilacinus collected in the present study was with a variety of this species (C. argenteolilacinus var. dovrensis) from

Norway (99% similarity). This species and variety have not been previously reported from

Ontario, although it has been found in Alberta (Brandrud et al. 2018). Cortinarius albocyaneus is another species previously described from northern Europe only, and the specimen sequenced in the current study matched closely (99% similarity) with a conspecific specimen from Sweden.

These affinities with Fennoscandian taxa highlight the possibility that the effects of forest management on fungal communities in Canada could parallel those in boreal Europe.

Morphospecies collected in the present study may also represent cryptic taxa that have been previously collected in Canada, but identified using European taxonomy (e.g. Miller and

Buyck 2002 and references therein). Cryptic taxa occur in fungi as a result of slower rates of phenotypic vs reproductive-barrier evolution (Taylor et al. 2006), leading to morphologically- similar but genetically distinct taxa. For example, Mycena pura represents a species complex comprising 11 cryptic species that cannot be distinguished by ITS sequencing (Harder et al.

2013). The two sequences of M. pura in the present study were located in different clades within a maximum-likelihood tree comprised almost entirely of M. pura sequences, suggesting that they represent cryptic taxa within this species complex. Due to their morphological similarity, however, they are both designated as the morphospecies ‘Mycena pura’. Multi-gene phylogenies would aid in delimiting species within these cryptic species complexes (O’Donnell et al. 2003).

The importance of constructing phylogenetic trees to confirm taxonomic identification is evident from the three sequenced specimens of Mycena urania (initially thought to represent distinct morphospecies). Although the sequence with maximum identity from BLAST searches of the NCBI and UNITE databases was a specimen of Mycena metata from Estonia, a maximum- likelihood tree grouped the query sequences with a specimen of M. urania from Sweden. This finding highlights the genetic variability that can occur among conspecific taxa from disparate

55 geographical locations and emphasizes the importance of phylogeography in shaping the current distributions of macrofungi.

Community composition

In this study, variation in the quantity and quality of downed CWD was strongly associated with differences in macrofungal community composition between unlogged (UL) and logged (CO, HR and FR) plots, although not among CO, HR and FR (logged) plots. Macrofungi in old-growth (UL) plots were associated with an abundance of CWD, a diversity of large- diameter CWD, particularly in late-decay stages, and closed-canopy, shade-tolerant, mesic conditions. In contrast, macrofungal assemblages in logged stands were associated with a low total volume of CWD, an abundance of small-diameter CWD and more shade-intolerant and hydric conditions.

As was true in the present study, old-growth stands contain a high volume and diversity of CWD (Spies et al. 1988, Linder et al. 1997, Siitonen et al. 2000), which differentiates them from managed second-growth forests (Franklin et al. 1981, Hansen et al. 1991, Tyrrell and Crow

1994). The high total volume of CWD in UL plots and low total volume in logged plots represents a reduction in dead wood availability due to previous forest management and younger forest age (Bader et al. 1995, Siitonen et al. 2000, Siitonen 2001, Sippola et al. 2001, Krankina et al. 2002). The higher volumes of large coniferous CWD in unlogged plots are also expected given that conifers are targeted by harvesting operations (MacDonald 1995).

Older stands contain larger trees (Delong and Kessler 2000); these will become large- diameter CWD in the absence of forest management through senescence and disturbances

(Sturtevant et al. 1997). This is reflected by the abundance of large, late-decay deciduous CWD

56 and large coniferous CWD in unlogged plots. In contrast, the high volumes of small-diameter

CWD in logged plots reflect the effects of previous harvesting operations and the fact that regenerating trees had not reached large sizes as yet (Fridman and Walheim 2000, Fraver et al.

2002, Pedlar et al. 2002). This is the result of management effectively reverting logged stands to an earlier successional stage (Delong and Kessler 2000) in which self-thinning is the main contributor to CWD accumulation. Self-thinning occurs in earlier stages of stand development

(Brassard and Chen 2006) and results from stem mortality of small, suppressed trees (Lang

1985), producing small, more quickly-decaying CWD (Sturtevant et al. 1997). The high volumes of small, late-decay CWD in logged plots may also be remaining logging slash that is small and quickly-decaying (Fraver et al. 2002), although in the present study the logged plots were old enough such that much of this small-diameter material had already disappeared.

A high volume of large coniferous CWD in UL plots reflects the closed-canopy conditions that characterize old-growth stands and favour shade-tolerant coniferous tree species

(Bergeron and Dubuc 1989, Brassard and Chen 2006). As the canopy closes, conifers increase in abundance with time since fire while shade-intolerant deciduous species decline (Bergeron and

Dubuc 1989, Bergeron and Dansereau 1993, Bergeron and Harvey 1997, Bergeron 2000, De

Grandpré et al. 2000, Gaulthier et al. 2000, Harper et al. 2002). This transition is also reflected in the results of the DCA on shrub/sapling communities, in which shade-tolerant white and black spruce (Picea glauca and Picea mariana, respectively), balsam fir (Abies balsamea) and eastern white cedar (Thuja occidentalis) were associated with UL plots, whereas shade-intolerant trembling aspen (Populus tremuloides) and paper birch (Betula papyrifera) were associated with logged plots.

The persistence of conifer dominance in UL plots is also evidenced by the input of large, early-decay coniferous CWD. Although currently mixedwood stands, they may be succeeding to more confer dominance due to the absence of fire (Bergeron and Dubuc 1989, De Grandpré et al.

2000, Harper et al. 2002, Lesieur et al. 2002), leading to replacement of shade-intolerant deciduous species with shade-tolerant coniferous species. This is in accordance with decreased fire cycles in Ontario boreal forests, which have been on the decline since the 1800s when the stands of the UL plots first established post-fire (Bergeron et al. 2001). Non-stand-replacing disturbance dynamics (e.g., windthrow) are therefore likely maintaining a continuous supply of large-diameter coniferous CWD, whereas frequent disturbance by fire would result in cyclic succession in which species are not replaced (see Brassard and Chen 2006), and shade-intolerant species remain dominant until the next disturbance by fire (Dix and Swan 1971, Carleton and

Maycock 1978, Johnson 1992, Cumming 2001, Chen and Popadiouk 2002).

The association of logged sites with an earlier-successional, more deciduous-dominated phase (Brassard and Chen 2006) reflects the change in species composition and increased deciduous component that occurs as a result of logging to the detriment of conifers. In the open, clearcut areas created by harvesting, recruitment of seedlings is favoured by shade-intolerant deciduous species (Coates 2002, Brassard and Chen 2006), but is poor for conifers; advanced regeneration may be destroyed completely due to logging, further impeding conifer regeneration

(Ruel et al. 2004). Logging may also produce a temporary input of nutrients, which are again beneficial to deciduous trees that are able to outcompete conifers on nutrient-rich sites (Brumelis and Carleton 1988, Carleton and MacLellan 1994). Logged sites may also have an increased abundance of ericaceous shrubs that would impede conifer growth (Ruel et al. 2004). Thus, despite our control of site conditions by controlling the proportion of deciduous and coniferous

58 trees (i.e., all plots are in mixedwood stands), it may be that the fundamentally more deciduous nature of logged sites remained. The hydric condition of some of the logged plots may be due to low-lying microsites in some areas, including in some cases alder swaths along old skid trails that had evidently been subjected to rutting.

The potential impacts of harvesting operations on conifer regeneration coupled with the removal of the original conifers themselves may hinder the transition of logged plots to a later successional stage even given sufficient time; this could affect not only community composition but also reduce the richness of macrofungi associated with conifers and conifer-tolerant conditions (e.g., shade, acidic soils; Bergeron and Dubuc 1989, Ste-Marie and Paré 1999,

Brassard and Chen 2006). This could exacerbate post-harvest reductions in large-diameter, late- decay coniferous CWD in logged plots, with particular impacts on the richness of rare or red- listed fungi that prefer these substrates (Kruys et al. 1999).

For example, the polypore Rhodofomes rosea (formerly Fomitopsis rosea) is a circumboreal, perennial species that produces fruitbodies on conifer wood, especially spruce

(Ryvarden and Gilbertson 1993). Although common in Ontario (e.g., Fischer et al. 2012), this species is threatened in Fennoscandia (Bendiksen et al. 1997, Larsson 1997) due to its confinement to old-growth, coniferous forests (Högberg and Stenlid 1999, Kauserud and

Schumacher 2003) and the rarity of these stands in boreal Europe (e.g., Linder and Östlund

1998). In the present study, R. rosea was sampled in three CO plots, one HR plot and twice in one UL plot. This suggests that the proximity of old-growth to logged stands is great enough so as not to limit spore dispersal (Edman et al. 2004b), and that successful colonization and fruiting body development can occur in logged plots given sufficient volume and quality of coniferous

CWD. Alternatively, R. rosea may be maintained in logged plots if local populations persist on

59 residual post-harvest substrates. If dispersal from surrounding unlogged plots is a limiting factor, however, local extinctions could occur in logged plots due to reduced genetic differentiation and inbreeding effects from smaller population sizes (Högberg et al. 1998), even given sufficient time post-harvest for coniferous dead wood to accumulate. The absence of R. rosea from FR plots suggests that the volume of large-diameter, early-decay coniferous CWD is sufficient to support R. rosea in control and half-removal plots but not in full-removal plots. In turn, this highlights the possibility that intensive harvesting that removes the majority of large-diameter trees (as seen in boreal Europe) could negatively impact the persistence of this species and lead to its increase in red-list status in Canada.

Fischer et al. (2012) found that R. rosea was significantly more common in unlogged than logged sites, and that this was associated with the large mean diameter of conifer logs in the former. Although the occurrence of this relatively rare species was not significantly different among treatments in the present study, this may be due to differences in methods: we surveyed presence/absence of R. rosea per plot, whereas Fischer et al. (2012) surveyed the number of fruiting bodies per spruce log.

The percentage basal area of deciduous trees had a significant effect on macrofungal community composition across sampling periods, and correlated positively with the volume of large, early-decay deciduous CWD, although its effect was primarily due to two unusual logged plots (ML10CO and ML10HR) that had high values for these variables. It appears that these two plots were in an area where a large residual amount of deciduous trees were left after the harvest, which subsequently mostly were windthrown. The result was a relatively high deciduous canopy composition, but an even more striking amount of large-diameter deciduous CWD.

The highly-significant effects of the CWD and habitat variables on the community composition of fungi in sampling period 1 may be partly the result of increased transect length in this sampling period; a larger sampling area surveys more individual pieces of dead wood, representing increased colonization opportunities and, in turn, increased richness of saproxylic fungi (Kruys and Jonsson 1999). Differences in the significance of the environmental variables among the three sampling periods may also highlight seasonal variation in community composition. Sampling period 1 took place in June, hence the fungal community sampled was skewed towards taxa inhabiting dead wood rather than a more even distribution across ecological guilds; this may be due to water retention in CWD compared to soil, which would facilitate growth of wood saprotrophs in the absence of heavy precipitation that only occurred later in the summer. The predominance of wood saprotrophs would therefore strengthen the associations between the sampled fungal community and the measured dead wood variables.

Ecological guilds

The impact of CWD availability on the richness of saproxylic macrofungi is seen from the consistently higher number of wood saprotrophic morphospecies in unlogged compared to logged plots. Across sampling periods, the number of wood saprotrophs decreased with increasing intensity of CWD removal in the manipulation experiment. The number of morphospecies was greatest in UL plots and smallest in FR plots (in both sampling periods 1 and

3, UL>CO>HR>FR, whereas in sampling period 2 UL>HR>CO>FR). These results reflect the strong correlation between saproxylic fungal richness and local CWD volume (Bader et al. 1995,

Ohlson et al. 1997, Sippola and Renvall 1999, Penttilä et al. 2004, Similä et al. 2006) and the

61 importance of a high volume and diversity of CWD for maintaining the high species richness of saproxylic taxa in boreal forests (Siitonen 2001).

In addition to increased dead wood availability, wood saprotrophs in UL plots may be benefitting from the stable, moist microclimate and long-term continuity of old-growth stands, which have been shown to support saproxylic richness (Siitonen 2001). Redundancy Analysis

(RDA) performed on the ecological guilds and environmental variables reflected a strong positive correlation between wood saprotrophs, shade-tolerant, coniferous conditions

(represented by axis DCA1) and UL plots, suggesting that these species benefit from a late- successional stage in stand development.

The impact of forest management reflected in the higher number of litter saprotrophs in

UL compared to FR plots in sampling periods 1 and 3 may result from the definition of this ecological guild in the present study: in addition to macrofungi that strictly utilize leaf and needle litter, I defined ‘litter saprotrophs’ as those species which additionally utilize humus and very fine woody debris confined to the soil. Similar to wood saprotrophs, the effects of CWD removal and changes in site conditions resulting from forest management can be seen from the decreased richness of litter saprotroph species in logged compared to unlogged plots.

As FWD and humus accumulate from the DWD available, sufficient supplies of CWD may also benefit litter saprotrophs that utilize these substrates and the compounds produced during their decomposition at later stages of decay (Steffen et al. 2002, Baldrian 2006, Baldrian

2008). This is reflected in the strong positive association between litter saprotrophs, unlogged plots and volumes of large-diameter, early- and late-decay coniferous CWD. These results suggest that high volumes of coniferous CWD in UL plots are important for maintaining a high species richness of litter saprotrophs that may subsequently utilize highly-fragmented pieces of

62 this dead wood. Alternatively, litter saprotrophs may benefit from the needle litter produced by coniferous trees; carpets of litter may be more abundant and stay moist longer in old-growth, heavily-forested stands where there are also larger volumes of coniferous CWD.

Treatment comparisons that did not follow the pattern UL>CO>HR>FR may have resulted in part from the methodology of the CWD removal experiment. Half removals were based on the volume of CWD originally present; as a result, some HR plots had higher volumes of CWD than CO plots even after manipulation.

In this study, the richness of ECM fungi did not differ between unlogged and logged plots and therefore did not appear to be affected by variation in CWD availability or any differences in site conditions that distinguished old-growth from post-logged stands. Other studies have shown logging to negatively impact ECM richness (Dahlberg et al. 2001). The absence of management effects on the number of ECM species in the present study in part may be due to the survey method undertaken. Fruiting body sampling can miss a large proportion of ECM species that would otherwise be identified by molecular or mycorrhizae sampling techniques (Danielson

1984, Gardes and Bruns 1996, Tedersoo et al. 2003). Additionally, if ECM fungi can colonize

CWD and soil indiscriminately (Tedersoo et al. 2003), their richness may not be affected by decreases in CWD availability due to logging, although community composition is likely to change (Jones et al. 2003). Similarly, many ECM genera are associated with a broad range of hosts (Cullings et al. 2000, Molina et al. 1992), which may inhibit the effects of post-harvest changes in tree composition on ECM richness.

Current harvesting practices in boreal Canada are designed to mimic the effects of natural disturbances (Ontario Ministry of Natural Resources 2001), but have been seen to reduce the volume of dead wood supplies below levels following fire (McRae et al. 2001, Moroni 2006,

Brassard and Chen 2008, Fischer et al. 2012). In addition, Canadian boreal forests are strongly impacted by insects and other non-stand replacing disturbances (Kneeshaw 2001, Natural

Resources Canada 2010), leading to the production of old-growth stands shaped by gap dynamics (Chen and Popadiouk 2002) with uneven age-class distributions and high structural complexity (Brassard and Chen 2006), as well as large volumes of CWD (Linder et al. 1997,

Sippola et al. 1998, Karjalainen and Kuuluvainen 2002).

Selective logging is one method of harvesting that attempts to emulate old-growth conditions. Dove and Keeton (2015) used a form of selective logging termed Structural

Complexity Enhancement (SCE) in a mature, multi-aged Vermont, US forest; SCE is designed to mimic natural disturbance events and accelerate old-growth features such as late-successional stand development and structural heterogeneity. Results indicated that plots harvested by SCE had significantly greater fungal richness eight years post-treatment, compared to conventional selection harvests and controls. Under SCE, fungal diversity was specifically affected by CWD recruitment and continuity of high levels of aboveground biomass. This emphasizes the importance of structural complexity and CWD availability for fungal communities.

In contrast, Bader et al. (1995) showed that selective logging significantly decreased the availability of large-diameter logs in late stages of decay 100 years post-harvest, and that the number of species of wood-inhabiting fungi decreased with increasing degree of cutting. Sippola et al. (2001) similarly found that the number of polypore species was significantly lower in selectively-logged compared to primeaval stands. This suggests that selective logging may fail to support the diversity of macrofungi in old-growth stands, although there is evidence that certain old-growth indicator polypores may be preserved through selective logging, including green tree retention (Kotiranta and Niemelä 1996).

Chapter 3. General Conclusions

The present study provides evidence of the negative effects of clearcutting on saprotrophic macrofungi in old-growth boreal mixedwood forests of northeastern Ontario. In the studied sites, old-growth plots hosted larger volumes of large-diameter coniferous CWD, and as a result supported a higher number of macrofungi. The overlap in species composition and harvesting effects between the current study and Fennoscandian studies, particularly the negative effects of clearcutting on macrofungi associated with CWD (Penttilä et al. 2006), suggests that harvesting in boreal Ontario could lead to similar increases in red-list status for susceptible taxa.

Coarse woody debris is not only integral to biodiversity and nutrient cycling in boreal forests (Harmon and Hua 1991, Samuelsson et al. 1994, Siitonen 2001), but also facilitates post- disturbance recovery and regeneration (Sernander 1936, Hytteborn et al. 1987, Hofgaard 1993,

Takahashi 1994, Lee et al. 1997, Kuuluvainen and Juntunen 1998, Parent et al. 2003, Lampainen et al. 2004) with late-decay CWD ‘nurse logs’ (Burns and Honkala 1990) functioning as seedbeds for developing seedlings (LeBarron 1950). This substrate is also an important structural component of forests (Kruys and Jonsson 1999, Hottola et al. 2009) that reduces erosion

(Harmon and Hua 1991), contributes to water storage (Fraver et al. 2002, Karjalainen and

Kuuluvainen 2002) and aids in soil development following humification (Siitonen 2001).

Retaining adequate volumes of CWD is therefore important for a number of ecosystem functions in Canadian boreal forests and to prevent the extensive loss of functionality and diversity seen in

European boreal forests due to over-harvesting (Siitonen 2001).

Literature Cited

Aanderaa, R., J. Rolstad, and S.M. Søngen. 1996. Biological diversity in forests. Norges Skogeierforbund og A/S Landbruksforlaget, Oslo, Norway.

Abrego, N. and I. Salcedo. 2013. Variety of woody debris as the factor influencing wood- inhabiting fungal richness and assemblages: is it a question of quantity or quality? Forest Ecology and Management 291:377-385.

Aerts, R. and O. Honnay. 2011. Forest restoration, biodiversity and ecosystem functioning. BMC Ecology 11, 29.

Ahlén, I. and M. Tjernberg, editors. 1996. Rödlistade Ryggradsdjur i Sverige. Artfakta, Artdatabanken, SLU, Uppsala.

Allen, R.B., P.K. Buchanan, P.W. Clinton, and A.J. Cone. 2000. Composition and diversity of fungi on decaying logs in a New Zealand temperate beech (Nothofagus) forest. Canadian Journal of Forest Research 30:1025-1033.

Allmér, J., 2005. Fungal communities in branch litter of Norway spruce. Doctoral dissertation. Dept. of Forest Mycology and Pathology, SLU. Acta Universitatis agriculturae Sueciae, vol. 2005, 125 pp.

Allmér, J., J. Stenlid, and A. Dahlberg. 2009. Logging-residue extraction does not reduce the diversity of litter-layer saprotrophic fungi in three Swedish coniferous stands after 25 years. Canadian Journal of Forest Research 39(9):1737-1748.

Altschul, S.F., T.L. Madden, A.A. Schaffer., J. Zhang, Z. Zhang, W. Miller, and D.J. Lipman. 1997. Gapped BLAST and PSIBLAST: a new generation of protein database search programs. Nucleic Acids Research 25:3389-3402.

Amiro, B.D., M.D. Flannigan, B.J. Stocks, and B.M. Wotton. 2002. Perspectives on carbon emissions from Canadian forest fires. Forestry Chronicle 78:388-390.

Amiro, B.D., B.J. Stocks, M.E. Alexander, M.D. Flannigan and B.M. Wotton. 2001. Fire, climate change, carbon and fuel management in the Canadian boreal forest. International Journal of Wildland Fire 10:405-413.

Andersson, L.I. and H. Hytteborn. 1991. Bryophytes and decaying wood - a comparison between managed and natural forest. Holoarctic Ecology. 14:121-130.

Angelstam, P. and T. Kuuluvainen. 2004. Boreal forest disturbance regimes, successional dynamics and landscape structures - a European perspective. Ecological Bulletin 41:117- 136.

Arnold, A.E., D.A. Henk, R.L. Eells, F. Lutzoni, and R. Vilgalys. 2007. Diversity and phylogenetic affinities of foliar fungal endophytes in loblolly pine inferred by culturing and environmental PCR. Mycologia 99:185-206.

Aronsson, M., T. Hallingäck, and J-E. Matsson. 1995. Swedish Red Data Book on Plants 1995. Swedish Threatened Species Unit, SLU, Uppsala. (In Swedish with an English summary.)

Åström, M., M. Dynesius, K. Hylander, and C. Nilsson. 2005. Effects of slash harvest on bryophytes and vascular plants in southern boreal forest clear-cuts. Journal of Applied Ecology 42:1194-1202.

Bader, P., S. Jansson, and B.G. Jonsson. 1995. Wood-inhabiting fungi and substratum decline in selectively logged boreal spruce forests. Biological Conservation 72:355-362.

Baldrian, P. 2006. Fungal laccases - occurrence and properties. FEMS Microbiology Reviews 30:215-242.

Baldrian, P. 2008. Wood-inhabiting ligninolytic basidiomycetes in soils: ecology and constraints for applicability in bioremediation. Fungal Ecology 1(1):4-12.

Bani, A., S. Pioli, M. Ventura, P. Panzacchi, L. Borruso, R. Tognetti, G. Tonon, and L. Brusetti. 2018. The role of microbial community in the decomposition of leaf litter and deadwood. Applied Soil Ecology 126:75-84.

Bässler, C., J. Müller, M. Svoboda, A. Lepšová, C. Hahn, H. Holzer, and V. Pouska. 2012. Diversity of wood-decaying fungi under different disturbance regimes - a case study from spruce mountain forests. Biodiversity and Conservation 21:33-49.

Bendiksen, E., K. Høiland, T.E. Brandrud, and J.B. Jordal. 1997. Truede og Sårbare Sopparter i Norge: en kommentert rødliste. Fungiflora, Oslo.

Berg, E. 1859. Die Wälder Finland. Jahrbuch der Könischen Sächsischen Academie für Forst- und Landwirthe 13:1-118.

Berglund, M. and M. Åström. 2007. Harvest of logging residues and stumps for bioenergy production - effects on soil productivity, carbon budget and species diversity. Mid Sweden University and Swedish Forest Agency, Sweden, p 19.

Berglund, H., J. Hottola, R. Penttilä, and J. Siitonen. 2011. Linking substrate and habitat requirements of wood-inhabiting fungi to their regional extinction vulnerability. Ecography 34:864-875.

Berglund, H. and B.G. Jonsson. 2005. Verifying an extinction debt among lichens and fungi in northern Swedish boreal forests. Conservation Biology 19:338-348.

Berglund, H. and B.G. Jonsson. 2008. Assessing the extinction vulnerability of wood-inhabiting fungal species in fragmented northern Swedish boreal forests. Biological Conservation 141:3029-3039.

Bergeron, Y. 2000. Species and stand dynamics in the mixed woods of Quebec’s southern boreal forest. Ecology 81:1500-1516.

Bergeron, Y. and P.-R. Dansereau. 1993. Predicting the composition of Canadian southern boreal forest in different ﬁre cycles. Journal of Vegetation Science 4:827-832.

Bergeron, Y. and M. Dubuc. 1989. Succession in the southern part of the Canadian boreal forest. Vegetation 79:51-63.

Bergeron, Y. and N.J. Fenton. 2012. Boreal forests of eastern Canada revisited: old growth, nonfire disturbances, forest succession, and biodiversity. Botany 90:509-523.

Bergeron, Y., S. Gauthier, V. Kafka, P. Lefort, and D. Lesieur. 2001. Natural ﬁre frequency for the eastern Canadian boreal forest: consequences for sustainable forestry. Canadian Journal of Forest Research. 31:384-391.

Bergeron, Y. and B.D. Harvey. 1997. Basing silviculture on natural ecosystem dynamics: an approach applied to the southern boreal mixedwood forest of Quebec. Forest Ecology and Management 92:235-242.

Bergeron, Y., A. Leduc, B.D. Harvey, and S. Gauthier. 2002. Natural fire regime: a guide for sustainable management of the Canadian boreal forest. Silva Fennica 36(1):81-95.

Boddy, L. 2000. Interspecific combative interactions between wood decaying basidiomycetes. FEMS Microbiology Ecology 31:185-194.

Boddy, L. 2001. Fungal community ecology and wood decomposition processes in angiosperms: from standing tree to complete decay of coarse woody debris. Ecological Bulletins 49:43- 56.

Boddy, L., J. Frankland, P. van West, editors. 2008. Ecology of Saprotrophic Basidiomycetes. Academic Press, London.

Boddy, L., O.M. Gibbon, and M.A. Grundy. 1985. Ecology of Daldinia concentrica: effect of abiotic variables on mycelial extension and interspecific interactions. Transactions of the British Mycological Society 85:201-211.

Boddy, L. and S.C. Watkinson. 1995. Wood decomposition, higher fungi, and their role in nutrient redistribution. Canadian Journal of Botany 73:1377-1383.

Bödeker, I.T.M., B.D. Lindahl, Å. Olson, and K.E. Clemmensen. 2016. Mycorrhizal and

saprotrophic fungal guilds compete for the same organic substrates but affect decomposition differently. Functional Ecology 30:1967-1978.

Bödeker, I.T.M., C.M.R. Nygren, A.F.S. Taylor, Å. Olson, and B.D. Lindahl. 2009. Class II peroxidase-encoding genes are present in a phylogenetically wide range of ectomycorrhizal fungi. The ISME Journal 3:1387-1395

Bonan, G.B. 2008. Forests and climate change: forcings, feedbacks, and the climate benefits of forests. Science 320:1444-1449.

Bonan, G.B. and H.H. Shugart. 1989. Environmental factors and ecological processes in boreal forests. Annual Review of Ecology and Systematics 20:1-28.

Bouchard, M., D. Kneeshaw, and Y. Bergeron. 2006. Forest dynamics after successive spruce budworm outbreaks in mixedwood forests. Ecology 87:2319-2329.

Bouchard, M. and D. Pothier. 2011. Long-term influence of fire and harvesting on boreal forest age structure and forest composition in eastern Québec. Forest Ecology and Management 261:811-820.

Brandrud, T.E., G. Schmidt-Stohn, K. Liimatainen, T. Niskanen, T.G. Frøslev, K. Soop, D. Bojantchev, I. Kytövuori, T.S. Jeppesen, F. Bellù, et al. 2018. Cortinarius sect. Riederi: taxonomy and phylogeny of the new section with European and North American distribution. Mycological Progress 17:1323-1354.

Brandt, J.P. 2009. The extent of the North American boreal zone. Environmental Reviews 17:101-161.

Brassard, B.W. and H.Y.H. Chen. 2006. Stand structural dynamics of North American boreal forests. Critical Reviews in Plant Sciences 25:115-137.

Brassard, B.W. and H.Y.H. Chen. 2008. Effects of forest type and disturbance on diversity of coarse woody debris in boreal forest. Ecosystems 11(7):1078-1090.

Bredesen, B., R. Haugan, R. Aanderaa, I. Lindblad, B. Økland, and O. Rosok. 1997. Wood- inhabiting fungi as indicators on ecological continuity within spruce forests of southeastern Norway. Blyttia 55:131-140.

Brumelis, G. and T.J. Carleton. 1988. The vegetation of postlogged black spruce lowlands in central Canada. I. Trees and tall shrubs. Canadian Journal of Forest Research 18:1470- 1478.

Bruns, T. and R.P. Shefferson. 2004. Evolutionary studies of ectomycorrhizal fungi: milestones and future directions. Canadian Journal of Botany 82:1122-1132.

Burns, R.M. and B.H. Honkala. 1990. Silvics of North America. Agriculture Handbook 654,

Vol. 1 and 2. Forest Service, USDA, Washington, DC.

Burton, P.J., Y. Bergeron, B.E.C. Bogdanski, G.P. Juday, T. Kuuluvainen, B.J. McAfee, A. Ogden, V.K. Teplyakov, R.I. Alfaro, D.A. Francis, et al. 2010. Sustainability of boreal forests and forestry in a changing environment. In: Mery, G., P. Katila, G. Galloway, R.I. Alfaro, M. Kanninen, M. Lobovikov, and J. Varjo (Eds.), Forests and society - responding to global drivers of change. IUFRO World Series Volume 25, Vienna, pp. 247-282.

Carleton, T.J. and P.F. Maycock. 1978. Dynamics of the boreal forest south of James Bay. Canadian Journal of Botany 56:1157-1173.

Carleton, T.J. and P. MacLellan. 1994. Woody vegetation responses to ﬁre versus clear-cutting logging: a comparative survey in the central Canadian boreal forest. Ecoscience 1:141- 152.

Caruso, A. 2008. Lichen diversity on stems, slash and stumps in managed boreal forests. Ph.D. Dissertation, Acta Universitatis Agriculturae Sueciae. 2008:3. Swedish University of Agricultural Sciences, Uppsala, pp. 1652-6880.

Chalot, M. and A. Bran. 1998. Physiology of organic nitrogen acquisition by ectomycorrhizal fungi and ectomycorrhizas. FMES Microbiology Review 22:21-44.

Chambers, S.M., R.M. Burke, P.R. Brooks, and J.W.G. Cairney. 1999. Molecular and biochemical evidence for manganese-dependent peroxidase activity in Tylospora fibrillosa. Mycological Research 103:1098-1102.

Chen, D.M., A.F.S. Taylor, R.M. Burke, and J.W.G. Cairney. 2001. Identification of genes for lignin peroxidases and manganese peroxidases in ectomycorrhizal fungi. New Phytologist 152:151-158.

Chen, H.Y.H. and R.V. Popadiouk. 2002. Dynamics of North American boreal mixedwoods. Environmental Reviews 10:137-166.

Christy, E.J., P. Sollins, and J. Trappe. 1982. First-year survival of Tsuga heterophylla without mycorrhizae and subsequent ectomycorrhizal development on decaying logs and mineral soil. Canadian Journal of Botany 60:1601-1605.

Clemmensen, K.E, R.D. Finlay, A. Dahlberg, and J. Stenlid. 2015. Carbon sequestration is related to mycorrhizal fungal community shifts during long-term succession in boreal forests. New Phytologist 205:1525-1536.

Coates, K.D. 2002. Tree recruitment in gaps of various size, clearcuts and undisturbed mixed forest of interior British Columbia, Canada. Forest Ecology and Management 155:387- 398.

Coates, D. and A.D.M. Rayner. 1985a. Fungal population and community development in cut beech logs. I. Establishment via the aerial cut surface. New Phytologist 101:153-171.

Coates, D. and A.D.M. Rayner. 1985b. Fungal population and community development in cut beech logs. II. Establishment via the buried cut surface. New Phytologist 101:173-181.

Coates, D. and A.D.M. Rayner. 1985c. Fungal population and community development in cut beech logs. III. Spatial dynamics, interactions and strategies. New Phytologist 101:183- 198.

Colpaert, J.V. and A. Van Laere.1996. A comparison of the extracellular enzyme activities of two ectomycorrhizal and a leaf-saprotrophic basidiomycete colonizing beech leaf litter. New Phytologist 134:133-141.

Conventional on Biological Diversity (CBD). 2009. Connecting biodiversity and climate change mitigation and adaptation. Second Ad Hoc Technical Expert Group on Biodiversity and Climate Change, CBD Technical Series No. 41. CBD Secretariat, Montreal, QC.

Cornwell, W.K., J.H.C. Cornelissen, K. Amatangelo, E. Dorrepaal, V.T. Eviner, O. Godoy, S.E. Hobbie, B. Hoorens, H. Kurokawa, N. Pérez-Harguindeguy, et al. 2008. Plant species traits are the predominant control on litter decomposition rates within biomes worldwide. Ecology Letters 11:1065-1071.

Courty, P.-E, N. Bréda, J. Garbaye. 2007. Relation between oak tree phenology and the secretion of organic matter degrading enzymes by Lactarius quietus ectomycorrhizas before and during bud break. Soil Biology and Biochemistry 39:1655-1663.

Courty, P.-E., M. Buee, A.G. Diedhiou, P. Frey-Klett, F. Le Tacon, F. Rineau, M.P. Turpault, S. Uroz, and J. Garbaye. 2010. The role of ectomycorrhizal communities in forest ecosystem processes: new perspectives and emerging concepts. Soil Biology & Biochemistry 42:679-698.

Crawford, R.H., S.E. Carpenter, and M.E. Harmon. 1990. Communities of filamentous fungi and yeasts in decomposing logs of Pseudotsuga menziesii. Mycologia 82:759-765.

Crites, S. and M.R.T. Dale. 1998. Diversity and abundance of bryophytes, lichens, and fungi in relation to woody substrate and successional stage in aspen mixedwood boreal forests. Canadian Journal of Botany 76:641-651.

Cumming, S.G. 2001. Forest type and wildﬁre in the Alberta boreal mixedwood: what do ﬁres burn? Ecological Applications 11:97-110.

Cyr, D., S. Gauthier, Y. Bergeron, and C. Carcaillet. 2009. Forest management is driving the eastern North American boreal forest outside its natural range of variability. Frontiers in Ecology and the Environment 10:519-524.

Dahlberg, A. 2002. Effects of fire on ectomycorrhizal fungi in Fennoscandian boreal forests. Silva Fennica 36(1):69-80.

Dahlberg, A., J. Schimmel, A.F.S. Taylor, and H. Johannesson. 2001. Post-fire legacy of ectomycorrhizal fungal communities in the Swedish boreal forest in relation to fire severity and logging intensity. Biological Conservation 100:151-161.

Danielson, R.M. 1984. Ectomycorrhizal associations in jack pine stands in northeastern Alberta. Canadian Journal of Botany 62:932-939.

De Grandpré, L., J. Morissette, and S. Gauthier. 2000. Long-term post-ﬁre changes in the northeastern boreal forest of Quebec. Journal of Vegetation Science 11:791-800.

Delong, S.C. and W.B. Kessler. 2000. Ecological characteristics of mature forest remnants left by wildfire. Forest Ecology and Management 131:93-106.

DeLong, H.B., V.J. Lieffers, and B.V. Blenis. 1997. Microsite effects on ﬁrst-year establishment and overwinter survival of white spruce in aspen-dominated boreal mixedwoods. Canadian Journal of Forest Research 27:1452-1457.

D.A. Benson, I. Karsch-Mizrachi, D.J. Lipman, J. Ostell, E.W. Sayers. 2010. GenBank. Nucleic Acids Research 38:D46-D51.

Dentinger, B.T.M., Y.D. Maryna, and J.-M. Moncalvo. 2011. Comparing COI and ITS as DNA barcode markers for mushrooms and allies (Agaricomycotina). PLoS ONE 6(9):e25081.

Desponts, M., G. Brunet, L. Bélanger, and. M. Bouchard. 2004. The eastern boreal old-growth balsam fir forest: a distinct ecosystem. Canadian Journal of Botany 82:830-849.

Diamond, J.M. and R.M. May. 1976. Island biogeography and the design of nature reserves. In: May, R.M. (Ed.), Theoretical ecology: principles and applications. W.B. Saunders, Philadelphia, pp. 163-186.

Dighton, J. and P.A. Mason. 1985. Mycorrhizal dynamics during forest tree development. In: Moore, D., L.A. Caselton, D.A. Wood, and J.C. Frankland (Eds.), Development biology of higher fungi. Cambridge University Press, Cambridge, pp. 117-139.

Dix, R. L. and J.M.A. Swan. 1971. The roles of disturbance and succession in upland forest at Candle Lake, Saskatchewan. Canadian Journal of Botany 49:657-676.

Dix, N. J. and J. Webster. 1995. Fungal Ecology. Chapman & Hall, London.

Dove, N.C. and W.S. Keeton. 2015. Structural complexity enhancement increases fungal species richness in northern hardwood forests. Fungal Ecology 13:181-192.

Durall, D.M., S. Gamiet, S.W. Simard, L. Kudrna, and S.M. Sakakibara. 2006. Effects of

clearcut logging and tree species composition on the diversity and community composition of epigeous fruit bodies formed by ectomycorrhizal fungi. Canadian Journal of Botany 84:966-980.

Durall, D.M., A.W. Todd, and J.M. Trappe. 1994. Decomposition of C-14-labeled substrates by ectomycorrhizal fungi in association with Douglas fir. New Phytologist 127:725-727.

Edman, M., M. Gustafsson, J. Stenlid, and L. Ericson. 2004a. Abundance and viability of fungal spores along a forestry gradient - responses to habitat loss and isolation. Oikos, 104:35- 42.

Edman, M. and B.G. Jonsson. 2001. Spatial pattern of downed logs and wood-decaying fungi in an old-growth Picea abies forest. Journal of Vegetation Science 12:609-620.

Edman, M., N. Kruys, and B.G. Jonsson. 2004b. Local dispersal sources strongly affect colonization patterns of wood-decaying fungi on spruce logs. Ecological Applications 14:893-901.

Egger, K.N. 2006. The surprising diversity of ascomycetous mycorrhizas. New Phytologist 170:421-423.

Egnell, G., R. Hyvönen, L. Högbom, T. Johansson, T. Lundmark, B. Olsson, E. Ring, and F. von Sydow. 2007. Environmental aspects of stump-harvest - compilation of knowledge and knowledge gaps. Energy Agency Report 40, 154 pp. (In Swedish.)

Eräjää, S., P. Halme, J.S. Kotiaho, A. Markkanen, and T. Toivanen. 2010. The volume and composition of dead wood on traditional and forest fuel harvested clear-cuts. Silva Fennica 44:203-211.

Esseen, P.-A., B. Ehnström, L. Ericson, and K. Sjöberg. 1992. Boreal forests the focal habitats of Fennoscandia. In: Hansson, L. (Ed.), Ecological principles of nature conservation. Elsevier, pp. 252-325.

Esseen, P.A., B. Ehnström, L. Ericson, and K. Sjöberg. 1997. Boreal forests. Ecological Bulletins 46:16-47.

Eveling, D.W., R.N. Wilson, E.S. Gillespie, and D. Bataille. 1990. Environmental effects on sporocarp counts over fourteen years in a forest area. Mycological Research 94:998- 1002.

Fahey, T.J., P.B. Woodbury, J.J. Battles, C.L. Goodale, S.P. Hamburg, S.V. Ollinger, and C.W. Woodall. 2010. Forest carbon storage: ecology, management, and policy. Frontiers in Ecology and the Environment 8:245-252.

Fesel, P.H. and A. Zuccaro. 2015. Beta-glucan: Crucial component of the fungal cell wall and elusive MAMP in plants. Fungal Genetics and Biology 90:53-60.

Ferrer, A. and G.S. Gilbert. 2003. Effect of tree host species on fungal community composition in a tropical rain forest in Panama. Diversity and Distributions 9:455-468.

Finlay, R. 2008. Ecological aspects of mycorrhizal symbiosis: with special emphasis on the functional diversity of interactions involving the extraradical mycelium. Journal of Experimental Botany 59:1115-1126.

Fleming, T. and B. Freedman. 1998. Conversion of natural, mixed-species forests to conifer plantations: implications for dead organic matter and carbon storage. Ecoscience 5:213- 221.

Fischer, A.L., J.M. Moncalvo, J.N. Klironomos, and J.R. Malcolm. 2012. Fruiting body and molecular rDNA sampling of fungi in woody debris from logged and unlogged boreal forests in Northeastern Ontario. Ecoscience 19:374-390.

Franklin, J.F., K. Cromack Jr., W. Denison, A. McKee, C. Maser, J. Sedell, F. Swanson, and G. Juday. 1981. Ecological characteristics of old-growth Douglas-fir forests. USDA For. Serv. Gen. Tech. Rep. PNW-118. Pacific Northwest For. and Range Exp. Stn. Portland, Ore., 48 pp.

Franklin, J.F., H.H. Shugart, and. M.E. Harmon. 1987. Tree death as an ecological process: the causes, consequences, and variability of tree mortality. BioScience 37:550-556.

Franzén, I., R. Vasaitis, R. Penttilä, and J. Stenlid. 2007. Population genetics of the wood- decaying fungus Phlebia centrifuga P. Karst. in fragmented and continuous habitats. Molecular Ecology 16:3326-3333.

Fraver, S., R.G. Wagner, and M. Day. 2002. Dynamics of coarse woody debris following gap harvesting in the Acadian forest of central Maine, U.S.A. Canadian Journal of Forest Research 32:2094-2105.

Fridman, J. and M. Walheim. 2000. Amount, structure, and dynamics of dead wood on managed forestland in Sweden. Forest Ecology and Management 131:23-36.

Fukasawa, Y., T. Osono, and H. Takeda. 2009. Microfungus communities of Japanese beech logs at different stages of decay in a cool temperate deciduous forest. Canadian Journal of Forest Research 39:1606-1614.

Fukasawa, Y., T. Osono, and H. Takeda. 2010. Beech log decomposition by wood-inhabiting fungi in a cool temperate forest floor: a quantitative analysis focused on the decay activity of a dominant basidiomycete Omphalotus guepiniformis. Ecological Research 25(5):959-966.

Gärdenfors, U. (Ed.) 2000. The 2000 Red List of Swedish Species. ArtDatabanken, SLU, Uppsala, 397 pp.

Gärdenfors, U. (Ed.) 2005. The 2005 Red List of Swedish Species. ArtDatabanken, SLU, Uppsala, 496 pp.

Gardes, M. and T. Bruns. 1993. ITS primers with enhanced specificity for basidiomycetes: application to identification of mycorrhizae and rusts. Molecular Ecology 2:113-118.

Gardes, M. and T. Bruns. 1996. Community structure of ectomycorrhizal fungi in a Pinus muricata forest: above- and below-ground views. Canadian Journal of Botany 74:1572- 1583.

Gauthier, S., L. De Grandpré, and Y. Bergeron. 2000. Differences in forest composition in two boreal forest ecoregions of Quebec. Journal of Vegetation Science 11:781-790.

Geml, J., I. Timling, C.H. Robinson, N. Lennon, H.C. Nusbaum, C. Brochmann, M.E. Noordeloos, and D.L. Taylor. 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 39:74-88.

Gessner, M.O., C.M. Swan, C.K. Dang, B.G. McKie, R.D. Bardgett, D.H. Wall, and S. Hättenschwiler. 2010. Diversity meets decomposition. Trends in Ecology and Evolution 25:372-380.

Gibb, H., J.P. Ball, T. Johansson, O. Atlegrim, J. Hjälten, and. K. Danell. 2005. Effects of management on coarse woody debris volume and composition in boreal forests in northern Sweden. Scandinavian Journal of Forest Research 20:213-222.

Goldammer, J.G., and V.V. Furyaev. 1996. Fire in ecosystems of boreal Eurasia: ecological impacts and links to the global system. In: Goldammer, J.G. and V.V. Furyaev (Eds.), Fire in ecosystems of boreal Eurasia. Kluwer Academic Publishers, Norwell, Mass., pp. 1-20.

Goldmann, K., I. Schöning, F. Buscot, and T. Wubet. 2015. Forest management type influences diversity and community composition of soil fungi across temperate forest ecosystems. Frontiers in Microbiology 6:1300.

Goodell, B., Y. Quian, and J. Jellison. 2008. Fungal decay of wood: soft rot-brown rot-white rot. In: Schultz, T., D. Nicholas, H. Militz, M.H. Freeman, and B. Goodell (Eds.), Development of commercial wood preservatives: efficacy, environmental, and health issues. ACS Symposium Series 982, American Chemical Society, Washington, D.C., pp. 9-31.

Goodman, D.M. and J.A. Trofymow. 1998. Distribution of ectomycorrhizas in microhabitats in mature and old-growth stands of Douglas fir on southeastern Vancouver Island. Soil Biology and Biochemistry 30:2127-2138.

Gramss, G., T.H. Günther, and W. Fritsche. 1998. Spot tests for oxidative enzymes in ectomycorrhizal, wood-and litter decaying fungi. Mycological Research 102:67-72.

Gray, A.N. and T.A. Spies. 1997. Microsite controls on tree seedling establishment in conifer forest canopy gaps. Ecology 78:2458-2473.

Gromtsev, A. 2002. Natural disturbance dynamics in the boreal forests of European Russia: a review. Silva Fennica 36(1):41-55.

Grove, S. 2002. Saproxylic insect ecology and the sustainable management of forests. Annual Review of Ecology and Systematics 33:1-23.

Groven, R., J. Rolstad, K.O. Storaunet, and E. Rolstad. 2002. Using forest stand reconstructions to assess the role of structural continuity for late-successional species. Forest Ecology and Management 164:39-55.

Gu, W., R. Heikkilä, and I. Hanski. 2002. Estimating the consequences of habitat fragmentation on extinction risk in dynamic landscapes. Landscape Ecology 17:699-710.

Gustafsson, L. and T. Hallingbäck. 1988. Bryophyte flora and vegetation of managed and virgin coniferous forests in south west Sweden. Biological Conservation 44:283-300.

Hagemann, U., M.T. Moroni, and F. Makeschin. 2009. Deadwood abundance in Labrador high- boreal black spruce forests. Canadian Journal of Forest Research 39:131-142.

Haider, K. and J. Trojanowski. 1980. A comparison of the degradation of IJC-labelled DHP and cornstalk lignins by micro- and macro-fungi and bacteria. In: Kirk, T.K., T. Higuchi, and H.-M. Chang (Eds.), Lignin biodegradation: microbiology, chemistry and potential applications, Vol. 1. CRC Press, Boca Raton, Fl., pp. 111-134.

Halme, P., J. Kotiaho, A.-L. Ylisirniö, J. Hottola, K. Junninen, J. Kouki, M. Lindgren, M. Mönkkönen, R. Penttilä, P. Renvall, et al. 2009a. Perennial polypores as indicators of annual and red-listed polypores. Ecological Indicators 9:256-266.

Halme, P., M. Mönkkönen, J. Kotiaho, A.-L. Ylisirniö, and A. Markkanen. 2009b. Quantifying the indicator power of an indicator species. Conservation Biology 23:1008-1016.

Hansen, A.J., T.A. Spies, F.J. Swanson, and J.L. Ohmann. 1991. Conserving biodiversity in managed forests - lessons from natural forests. BioScience 41:382-92.

Hansen, J., R. Türk, G. Vogg, R. Heim, and E. Beck. 1997. Conifer carbohydrate physiology: updating classical views. In: Rennenberg, H., W. Eschrich, and H. Ziegler (Eds.), Trees - contributions to modern tree physiology. Backhuys, Leiden, pp. 97-108.

Hanski, I. 2000. Extinction debt and species credit in boreal forests: modelling the consequences

of different approaches to biodiversity conservation. Annales Zoolgici Fennici 37:271- 280.

Harder, C.B., T. Laessoe, T.G. Froslev, F. Ekelund, S. Rosendahl, and R. Kjøller. 2013. A three- gene phylogeny of the Mycena pura complex reveals 11 phylogenetic species and shows ITS to be unreliable for species identification. Fungal Biology 117:764-775.

Hari, P. and L. Kulmala. 2008. Advances in global change research 34: boreal forest and climate change. Springer, New York, 582 pp.

Harley, J.L. 1971. Fungi in ecosystems. Journal of Ecology 59:653-668.

Harmon, M.E., W.K. Ferrel, and J.F. Franklin. 1990. Effects on carbon storage of conversion of old-growth forests to young forests. Science 247:699-702.

Harmon, M.E, J.F. Franklin, F.J. Swanson, P. Sollins, S.V. Gregory, J.D. Lattin, N.H. Anderson, S.P. Cline, N.G. Aumen, J.R. Sedell, et al. 1986. Ecology of coarse woody debris in temperate ecosystems. Advances in Ecological Research 15:133-302.

Harmon, M.E. and C. Hua. 1991. Coarse woody debris dynamics in two old-growth ecosystems: comparing a deciduous forest in China and a conifer forest in Oregon. BioScience 41:604-610.

Harper, K.A., Y. Bergeron, S. Gauthier, and P. Drapeau. 2002. Post-ﬁre development of canopy structure and composition in black spruce forests of Abitibi, Québec: a landscape scale study. Silva Fennica 36:249-263.

Hartmann, M., C.G. Howes, D. Vaninsberghe, H. Yu, D. Bachar, R. Christen, R.H. Nilsson, S.J. Hallam, and W.W. Mohn. 2012. Significant and persistent impact of timber harvesting on soil microbial communities in northern coniferous forests. The ISME Journal 6:2199- 2218.

Harvey, A.E., M.F. Jurgensen, and M.J. Larsen. 1978. Seasonal distribution of ectomycorrhizae in a mature Douglas fir/larch forest soil in western Montana. Forest Science 24:203-208.

Harvey, A.E., M.J. Larsen, and M.F. Jurgensen. 1979. Comparative distribution of ectomycorrhizae in soils of three western Montana forest habitat types. Forest Science 25:350-358.

Hättenschwiler, S., A.V. Tiunov, and S. Scheu. 2005. Biodiversity and litter decomposition in terrestrial ecosystems. Annual Review of Ecology, Evolution, and Systematics 36:191- 218.

Heikinheimo, O. 1915. Der Einfluss der Brandwirtschaft auf die Wälder Finlands. Acta Forestalia Fennica 4:1-264 + 149 pp. of appendices. (In Finnish with German summary.)

Heilmann-Clausen, J. 2001. A gradient analysis of communities of macrofungi and slime moulds on decaying beech logs. Mycological Research 105:575-596.

Heilmann-Clausen, J. and M. Christensen. 2003. Fungal diversity on decaying beech logs - implications for sustainable forestry. Biodiversity and Conservation 12:953-973.

Heilmann-Clausen, J. and M. Christensen. 2004. Does size matter? On the importance of various dead wood fractions for fungal diversity in Danish beech forests. Forest Ecology and Management 201:105-117.

Heilmann-Clausen, J. and M. Christensen. 2005. Wood-inhabiting macrofungi in Danish beech- forests - conflicting diversity patterns and their implications in a conservation perspective. Biological Conservation 122:633-642.

Henry, H.A.L. 2008. Climate change and soil freezing dynamics: historical trends and projected changes. Climatic Change 87:421-434.

Hobbie, S.E., P.B. Reich, J. Oleksyn, M. Ogdahl, R. Zytkowiak, C. Hale, and P. Karolewski. 2006. Tree species effects on decomposition and forest floor dynamics in a common garden. Ecology 87:2287-2297.

Hofgaard, A. 1993. Structure and regeneration patterns in a virgin Picea abies forest in northern Sweden. Journal of Vegetation Science 4:601-608.

Högberg, M.N., M.J.I. Briones, S.G. Keel, D.B. Metcalfe, C. Campbell, A.J. Midwood, B. Thornton, V. Hurry, S. Linder, T. Näsholm, et al. 2010. Quantification of effects of season and nitrogen supply on tree below-ground carbon transfer to ectomycorrhizal fungi and other soil organisms in a boreal pine forest. New Phytologist 187:485-493.

Högberg, N., O. Holdenrieder, and J. Stenlid. 1999. Population structure of the wood decay fungus Fomitopsis pinicola. Heredity 83:354-360.

Högberg, N. and J. Stenlid. 1999. Population genetics of Fomitopsis rosea - a wood-decay fungus of the old-growth European taiga. Molecular Ecology 8:703-710.

Högberg, N., J. Stenlid, and P. Angelstam. 1998. Observations of low fertility for Swedish populations of Fomitopsis rosea. In: Högberg, N. Population biology of common and rare saprotrophic wood-decay fungi. Ph.D. Thesis. Swedish University of Agricultural Sciences, Uppsala.

Högberg, N., J. Stenlid, and J.-O. Karlsson. 1995. Genetic differentiation in Fomitopsis pinicola (Schwarts: Fr.) Karst studied by means of arbitrary primed-PCR. Molecular Ecology 4:675-680.

Hörnberg, G.O., U. Zackrisson, U. Segerström, B.W. Svensson, M. Ohlson and R.H.W.

Bradshaw. 1998. Boreal swamp forests biodiversity ‘hotspots’ in an impoverished forest landscape. BioScience 48:795-802.

Høiland, K. and E. Bendiksen. 1996. Biodiversity of wood-inhabiting fungi in a boreal coniferous forest in Sor-Trondelag County, Central Norway. Nordic Journal of Botany 16(6):643-659.

Horton, T.R. and T.D. Bruns. 2001. The molecular revolution in ectomycorrhizal ecology: peeking into the black box. Molecular Ecology 10:1855-1871.

Hottola, J., O. Ovaskainen, and I. Hanski. 2009. A unified measure of the number, volume and diversity of dead trees and the response of fungal communities. Journal of Ecology 97:1320-1328.

Hottola, J. and J. Siitonen. 2008. Significance of woodland key habitats for polypore diversity and red-listed species in boreal forests. Biodiversity and Conservation 17:2559-2577.

Humphrey, J.W., A.C. Newton, A.J. Peace, and E. Holden. 2000. The importance of conifer plantations in northern Britain as a habitat for native fungi. Biological Conservation 96:241-252.

Hunter, J.C. and V.T. Parker. 1993. The disturbance regime of an old-growth forest in coastal California. Journal of Vegetation Science 4:19-24.

Hytteborn, H., J.R. Packham, and T. Verwijst. 1987. Tree population dynamics, stand structure and species composition in the montane virgin forest of Vallibacken, northern Sweden. Vegetatio 72:3-19.

Ishida, T.A., K. Nara, and T. Hogetsu. 2007. Host effects on ectomycorrhizal fungal communities: insight from eight host species in mixed conifer-broadleaf forests. New Phytologist 174:430-440.

Izzo, A., J. Agbowo, and T.D. Bruns. 2005. Detection of plot-level changes in ectomycorrhizal communities across years in an old-growth mixed conifer forest. New Phytologist 166:619-630.

Jones, C.G., J.H. Lawton, and M. Shachak. 1994. Organisms as ecosystem engineers. Oikos 69:373-386.

Johnson, E.A. 1992. Fire and vegetation dynamics: studies from the North American boreal forest. Cambridge University Press, Cambridge, 144 pp.

Jones, M.D., D.M. Durall, and J.W.G. Cairney. 2003. Ectomycorrhizal fungal communities in young forest stands regenerating after clearcut logging. New Phytologist 157:399-422.

Jonsson, B.G. and N. Kruys. (Eds.) 2001. Ecology of woody debris in boreal forests. Ecological

Bulletins 49, 280 pp.

Jonsson, B.G., N. Kruys, and T. Ranius. 2005. Ecology of species living on dead wood - lessons for dead wood management. Silva Fennica 39:289-309.

Jönsson, M.T., M. Edman, and B.G. Jonsson. 2008. Colonization and extinction patterns of wood-decaying fungi in a boreal old-growth Picea abies forest. Journal of Ecology 96:1065-1075.

Junninen, K. and A. Komonen. 2011. Conservation ecology of boreal polypores: a review. Biological Conservation 144:11-20.

Junninen, K., J. Kouki, and P. Renvall. 2008. Restoration of natural legacies of fire in European boreal forests: an experimental approach to the effects on wood-decaying fungi. Canadian Journal of Forest Research 38:202-215.

Junninen, K., R. Penttilä, and P. Martikainen. 2007. Fallen retention aspen trees on clear-cuts can be important habitats for red-listed polypores: a case study in Finland. Biodiversity and Conservation 16:475-490.

Junninen, K., M. Similä, J. Kouki, and H. Kotiranta. 2006. Assemblages of wood-inhabiting fungi along the gradients of succession and naturalness in boreal pine-dominated forests in Fennoscandia. Ecography 29:75-83.

Juutilainen, K., P. Halme, H. Kotiranta, and M. Mönkkönen. 2011. Size matters in studies of dead wood and wood-inhabiting fungi. Fungal Ecology 4:342-349.

Kagawa, A., A. Sugimoto, and T.C. Maximow. 2006. Seasonal course of translocation, storage and remobilization of 13C pulse-labeled photoassimilate in naturally growing Larix gmelinii saplings. New Phytologist 171:793-804.

Kaila, L., P. Martikainen, P. Punttila, and E. Yakovlev. 1994. Saproxylic beetles (Coleoptera) on dead birch trunks decayed by different polypore species. Annales Zoologici Fennici 31:97-107.

Kaila, L., P. Martikainen, and P. Punttila. 1995. Pökkelöt hakkuuaukoilla-hyötyvätkö taantuneet lahopuukuoriaiset. In: S. Hannelius S. and P. Niemelä P (Eds.), Monimuotoisuus metsien hoidossa. Vantaa Metsäntutkimuslaitoksen tiedonantoja 564, pp. 65-71.

Karjalainen, L. and T. Kuuluvainen. 2002. Amount and diversity of coarse woody debris within a boreal forest landscape dominated by Pinus sylvestris in Vienansalo wilderness, eastern Fennoscandia. Silva Fennica 36:147-167.

Karström, M. 1993 Indikatorarter som biologisk inventeringsmetod. In: Gransberg, M., M.

Karström, K. Lindahl, G.A. Olsson, and M. Williamsson M. (Eds.), Indikatorarter för identifiering av naturskogar i Norrbotten Naturvårdsverket. Rapport 4276 Solna, pp. 19- 96 (In Swedish.)

Kauserud, H., M. Lie, O. Stensrud, and M. Ohlson. 2005. Molecular characterization of airborne fungal spores in boreal forests of contrasting human disturbance. Mycologia 97:1215- 1224.

Kauserud, H. and T. Schumacher. 2003. Genetic structure of Fennoscandian populations of the threatened wood-decay fungus Fomitopsis rosea (Basidiomycota). Mycological Research 107:155-163.

Kebli, H., S. Brais, G. Kernaghan, and P. Drouin. 2012. Impact of harvesting intensity on wood- inhabiting fungi in boreal aspen forests of Eastern Canada. Forest Ecology and Management 279:45-54.

Kendrick, B. 1992. The Fifth Kingdom, 2nd ed. Mycologue Publications, Waterloo, 386 pp.

Kirk, R. 1966. The Olympic Rainforest. University of Washington Press, Seattle, Wash., 86 pp.

Kirk, T.K., and P. Fenn. 1982. Formation and action of the ligninolytic system in basidiomycetes. In: Frankland, J.C., J. N. Hedger, and M. J. Swift (Eds.), Decomposer basidiomycetes: their biology and ecology. Cambridge University Press, Cambridge, pp. 67-90.

Kneeshaw, D. 2001. Are non-fire gap disturbances important to boreal forest dynamics? Recent Developments in Ecology 1:43-58.

Kneeshaw, D. and S. Gauthier. 2003. Old growth in the boreal forest: a dynamic perspective at the stand and landscape level. Environmental Reviews 11:S99-S114.

Kohler, A., A. Kuo, L.G. Nagy, E. Morin, K.W. Barry, F. Buscot, B. Canbäck, C. Choi, N. Cichocki, A. Clum, et al. 2015. Convergent losses of decay mechanisms and rapid turnover of symbiosis genes in mycorrhizal mutualists. Nature Genetics. 47:410-415.

Koide, R.T., J.N. Sharda, J.R. Herr, and G.M. Malcolm. 2008. Ectomycorrhizal fungi and the biotrophy-saprotrophy continuum. New Phytologist 178:230-233.

Koide, R.T., D.S. Shumway, B. Xu, and J.N. Sharda. 2007. On temporal partitioning of a community of ectomycorrhizal fungi. New Phytologist 174:420-429.

Kõljalg, U., K.-H. Larsson, K. Abarenkov, R.H. Nilsson, I.J. Alexander, U. Eberhardt, S. Erland, K. Høiland, R. Kjøller, E. Larsson, et al. 2005. UNITE: a database providing web-based methods for the molecular identification of ectomycorrhizal fungi. New Phytologist 166:1063-1068.

Komonen, A., M. Jonsell, B. Økland, A. Sverdrup-Thygeson, and K. Thunes. 2004. Insect assemblage associated with the polypore Fomitopsis pinicola: a comparison across Fennoscandia. Entomologica Fennica 15:102-112.

Kotiranta, H. and T. Niemelä. 1996. Threatened polypores in Finland. 2nd revised edition. Finnish Environment Institute and Edita, Ympäristöopas 10, Helsinki, 184 pp. (In Finnish with an English summary).

Kouki, J., S. Löfman, P. Martikainen, S. Rouvinen, and A. Uotila. 2001. Forest fragmentation in Fennoscandia: linking habitat requirements of wood-associated threatened species to landscape and habitat changes. Scandinavian Journal of Forest Research 16 (suppl. 3): 27-37.

Krajick, K. 2001. Defending deadwood. Science 293:1579-1581.

Krankina, O.N., M.E. Harmon, Y.A. Kukuev, R.F. Treyfeld, N.N. Kashpor, V.G. Kresnov, V.M. Skudin, N.A. Protasov, M. Yatskov, G. Spycher, et al. 2002. Coarse woody debris in forest regions of Russia. Canadian Journal of Forest Research 32:768-778.

Kruys, N. and B.G. Jonsson. 1999. Fine woody debris is important for species richness on logs in managed boreal spruce forests of northern Sweden. Canadian Journal of Forest Research 29(8):1295-1299.

Kruys, N., C. Fries, B.G. Jonsson, T. Lämas, and G. Stahl. 1999. Wood-inhabiting cryptogams on dead Norway spruce (Picea abies) trees in managed Swedish boreal forests. Canadian Journal of Forest Research 29(2):178-186.

Küffer, N., F. Gillet, B. Senn-Irlet, M. Aragno, and D. Job. 2008. Ecological determinants of fungal diversity on deadwood in European forests. Fungal Diversity 30:83-95.

Küffer, N. and B. Senn-Irlet. 2005. Inﬂuence of forest management on the species richness and composition of wood-inhabiting basidiomycetes in Swiss forests. Biodiversity and Conservation 14:2419-2435.

Kuo, M. 2015. Polyporus varius. Retrieved from the MushroomExpert.Com Web site: http://www.mushroomexpert.com/polyporus_varius.html

Kurz, W.A., C.H. Shaw, C. Boisvenue, G. Stinson, J. Metsaranta, D. Leckie, A. Dyk, C. Smyth, and E.T. Neilson. 2013. Carbon in Canada’s boreal forest - a synthesis. Environmental Reviews 21:260-292.

Kuuluvainen, T. 2002. Natural variability of forests as a reference for restoring and managing biological diversity in boreal Fennoscandia. Silva Fennica 36:97-125.

Kuuluvainen, T. and P. Juntunen. 1998. Seedling establishment in relation to microhabitat

variation in a windthrow gap in a boreal Pinus sylvestris forest. Journal of Vegetation Science 9:551-562.

Kuusinen, M., K. Jääskeläinen, L. Kivistö, A. Kokko, and S. Lommi. 1995. Indikaattorijäkälien kartoitus Kainuussa. Metsähallituksen Luonnonsuojelujulkaisuja Sarja A 39:1-22.

Labbé, R. 2014a, May. Gymnopus confluens. Retrieved from https://www.mycoquebec.org.

Labbé, R. 2014b, May. Trichaptum fuscoviolaceum. Retrieved from https://www.mycoquebec.org.

Labbé, R. 2017, March. Cerioporus varius. Retrieved from https://www.mycoquebec.org.

Labbé, R. 2018, April. Mycena leptocephala. Retrieved from https://www.mycoquebec.org.

Labbé, R. 2019a, May. Cortinarius decipiens. Retrieved from https://www.mycoquebec.org.

Labbé, R. 2019b, August. Hydnoporia tabacina. Retrieved from https://www.mycoquebec.org.

Lafleur, B., S. Renard, C. Leroy, N.J. Fenton, M. Simard, S. Gauthier, D. Paré, A. Leduc, N. Thiffault, and Y. Bergeron. 2016. Silviculture to sustain productivity in black spruce paludified forests. Forest Ecology and Management 375:172-181.

Lampainen, J., T. Kuuluvainen, T.H. Wallenius, L. Karjalainen, and I. Vanha-Majamaa. 2004. Long-term forest structure and regeneration after wildfire in Russian Karelia. Journal of Vegetation Science 15:245-256.

Lang, G.E. 1985. Forest turnover and the dynamics of bole wood litter in subalpine balsam fir forest. Canadian Journal of Forest Research 15:262-268.

Larsson, K.H. (Ed.) 1997. Rödlistade svampar i Sverige. Artfakta, ArtDatabanken, SLU, Uppsala. (In Swedish.)

Larsson, S. and K. Danell. 2001. Science and the management of boreal forest diversity. Scandinavian Journal of Forest Research 16:S3, 5-9.

LeBarron, R.K. 1950. Silvicultural management of black spruce in Minnesota. USDA Circ. 791. U.S. Gov. Print. Off., Washington, D.C., 66 pp.

Lee, P.C., S. Crites, M. Nietfield, H. Van Guyen, and J.B. Stelfox. 1997. Characteristics and origins of deadwood material in aspen-dominated boreal forests. Ecological Applications 7:691-701.

Lepš, J. and P. Šmilauer. 2003. Multivariate analysis of ecological data using CANOCO. Cambridge University Press, Cambridge, 282 pp.

Lesica, P., B. McCune, V. Cooper, and W.S. Hong. 1991. Differences in lichen and bryophyte communities between old-growth and managed second-growth forests in the Swan Valley, Montana. Canadian Journal of Botany 69:1745-1755.

Lesieur, D., S. Gauthier, and Y. Bergeron. 2002. Fire frequency and vegetation dynamics for the south-central boreal forest of Quebec, Canada. Canadian Journal of Forest Research 32:1996-2009.

Lindahl, B. and J. Boberg. 2008. Distribution and function of litter basidiomycetes in coniferous forests. In: Boddy, L., J.C. Frankland, and P. van West (Eds.), Ecology of saprotrophic basidiomycetes. Elsevier, Amsterdam, pp. 183-209.

Lindahl, B.D., R.D. Finlay, and J.W.G. Cairney. 2005. Enzymatic activities of mycelia in mycorrhizal fungal communities. In: Dighton, J., J.F. White, and P. Oudemans (Eds.), The fungal community: its organization and role in the ecosystem, 3rd ed. CRC Press, Boca Raton, Florida, pp. 331-348.

Lindahl, B.D., K. Ihrmark, J. Boberg, S.E. Trumbore, P. Högberg, J. Stenlid and R.D. Finlay. 2006. Spatial separation of litter decomposition and mycorrhizal nitrogen uptake in a boreal forest. New Phytologist 173:611-620.

Lindahl, B.D. and S. Olsson. 2004. Fungal translocation - creating and responding to environmental heterogeneity. Mycologist 18:79-88.

Lindahl, B.D. and A.F.S. Taylor. 2004. Occurrence of N-acetylhexosaminadase encoding genes in ectomycorrhizal basidiomycetes. New Phytologist 164:193-199.

Lindblad, I. 1998. Wood-inhabiting fungi on fallen logs of Norway spruce: relations to forest management and substrate quality. Nordic Journal of Botany 18:243-255.

Lindenmayer, D.B. and J.F. Franklin. 2002. Conserving forest biodiversity: a comprehensive multiscaled approach. Island Press, Washington, D.C., 351 pp.

Linder, P., B. Elfving, and O. Zackrisson. 1997. Stand structure and successional trends in virginal boreal forest reserves in Sweden. Forest Ecology and Management 98:17-33.

Linder, P. and L. Östlund. 1998. Structural changes in three mid-boreal Swedish forest landscapes, 1885-1996. Biological Conservation 85:9-19.

Lindhe, A., N. Åsenblad, and H.-G. Toresson. 2004. Cut logs and high stumps of spruce, birch, aspen and oak - nine years of saproxylic fungi succession. Biological Conservation 119:443-454.

Lodge, D.J., J.F. Ammirati, T.E. O’Dell, and G.M. Mueller. 2004. Collecting and describing

macrofungi. In: Mueller, G.M., G.F. Bills, and M.S. Foster (Eds.), Biodiversity of fungi: inventory and monitoring methods. Elsevier Academic Press, San Diego, Calif., pp. 128- 158.

Lõhmus, A. 2011. Aspen-inhabiting Aphyllophoroid fungi in a managed forest landscape in Estonia. Scandinavian Journal of Forest Research 26:212-220.

Lonsdale, D., M. Pautasso, and O. Holdenrieder. 2008. Wood-decaying fungi in the forest: conservation needs and management options. European Journal of Forest Research 127:1-22.

Lumley, T.C., L.D. Gignac, and R.S. Currah. 2001. Microfungus communities of white spruce and trembling aspen logs at different stages of decay in disturbed and undisturbed sites in the boreal mixedwood region of Alberta. Canadian Journal of Botany 79:76-92.

Luis, P., H. Kellner, B. Zimdars, U. Langer, F. Martin, and F. Buscot. 2005. Patchiness and spatial distribution of laccase genes of ectomycorrhizal, saprotrophic and unknown basidiomycetes in the upper horizons of a mixed forest Cambisol. Microbial Ecology 50:570-579.

MacDonald, G.B. 1995. The case for boreal mixedwood management: an Ontario perspective. Forestry Chronicle 71:725-734.

Mahmood, S., R. Finlay, and S. Erland. 1999. Effects of repeated harvesting of forest residues on the ectomycorrhizal community in a Swedish spruce forest. New Phytologist 142:557- 585.

Marks, G.C. and T.T. Kozlowski. (Eds.) 1973. Ectomycorrhizae - their ecology and physiology. Academic Press, Inc., New York, 444 pp.

Mäkipää, R., T. Rajala, D. Schigel, K.T. Rinne, T. Pennanen, N. Abrego, and O. Ovaskainen. 2017. Interactions between soil- and dead wood-inhabiting fungal communities during the decay of Norway spruce logs. ISME Journal 11:1964-74.

Martikainen, P., J. Siitonen, L. Kaila, P. Punttila, and J. Rauh. 1999. Bark beetles (Coleoptera, Scolytidae) and associated beetle species in mature managed and old growth boreal forests in southern Finland. Forest Ecology and Management 116:233-245.

Martikainen, P., J. Siitonen, P. Punttila, L. Kaila, and J. Rauh. 2000. Species richness of Coleoptera in mature managed and old-growth boreal forests in southern Finland. Biological Conservation 94(2):199-209.

Martin, F. 1985. 15N-NMR studies of nitrogen assimilation and amino acid biosynthesis in the ectomycorrhizal fungus Cenococcum graniforme. FEBS Letters 182:350-354.

Maser, C. and J.M. Trappe. (Eds.) 1984. The seen and unseen world of the fallen tree, General

Technical Report PNW-164. US Department of Agriculture, Forest Service, Portland, Ore., 56 pp.

McRae, D.J., L.C. Duchesne, B. Freedman, T.J. Lynham, and S. Woodley. 2001. Comparisons between wildfire and forest harvesting and their implications in forest management. Environmental Reviews 9(4):223-260.

Mehus, H. 1986. Fruit body production of macrofungi in some North Norwegian forest types. Nordic Journal of Botany 6:679-702.

Miles, L., and V. Kapos. 2008. Reducing greenhouse gas emissions from deforestation and forest degradation: global land-use implications. Science 320:1454-1455.

Millennium Ecosystem Assessment. 2005. Millennium Ecosystem Assessment, Ecosystems and human well being. Policy Response Options. Chapter 8: wood, fuelwood and nonwood forest products. Island Press, Washington, D.C., pp. 257-293.

Miller, S.L. and B. Buyck. 2002. Molecular phylogeny of the genus Russula in Europe with a comparison of modern infrageneric classifications. Mycological Research 106:259-276.

Montreal Process, 1995. Criteria and Indicators for the Conservation and Sustainable Management of Temperate and Boreal Forests. Canadian Forest Service, Hull, QC, 27 pp.

Moore, J.C., E.L. Berlow, D.C. Coleman, P.C. de Ruiter, Q. Dong, A. Hastings, N.C. Johnson, K. McCann, K. Melville, P.J. Morin, et al. 2004. Detritus, trophic dynamics and biodiversity. Ecology Letters 7:584-600.

Moose, R.A., D. Schigel, L.J. Kirby, and M. Shumskaya. 2019. Dead wood fungi in North America: an insight into research and conservation potential. Nature Conservation 32:1- 17.

Moroni, M.T. 2006. Disturbance history affects dead wood abundance in Newfoundland boreal forests. Canadian Journal of Forest Research 36:3194-3208.

Müller-Using, S. and N. Bartsch. 2009. Decay dynamic of coarse and fine woody debris of a beech (Fagus sylvatica L.) forest in central Germany. Canadian Journal of Forest Research 218:287-296.

Natural Resources Canada, 2010. http://atlas.nrcan.gc.ca

Nehls, U., A. Bock, W. Einig, and R. Hampp. 2001. Excretion of two proteases by the ectomycorrhizal fungus Amanita muscaria. Plant, Cell and Environment 24:741-747.

Niemelä, J. 1999. Management in relation to disturbance in the boreal forest. Forest Ecology and Management 115:127-134.

Niemelä, T. 2005. Polypores, lignicolous fungi. Norrlinia 13:1-319. (In Finnish with English summary.)

Niemelä, T., P. Renvall, and R. Penttilä. 1995. Interactions of fungi at late stages of wood decomposition. Annales Botanici Fennici 32:141-152.

Nilsson, R.H., E. Kristiansson, M. Ryberg, N. Hallenberg, and K.H. Larsson. 2008. Intraspecific ITS variability in the kingdom Fungi as expressed in the international sequence databases and its implications for molecular species identification. Evolutionary Bioinformatics Online 4:193-201.

Nilsson, S.G., J. Hedin, and M. Niklasson. 2001. Biodiversity and its assessment in boreal and nemoral forests. Scandinavian Journal of Forest Research 16:S3,10-26.

Nitare, J. (Ed.) 2000. Indicator species for assessing the nature conservation value of woodland sites - a flora of selected cryptogams. The Swedish National Board of Forestry, Jönköping. (In Swedish with an English summary.)

Nitterus, K., M. Astrom, and B. Gunnarsson. 2007. Commercial harvest of logging residue in clear-cuts affects the diversity and community composition of ground beetles (Coleoptera: Carabidae). Scandinavian Journal of Forest Research 22(3):231-240.

Nordén, B. 1997. Genetic variation within and among populations of Fomitopsis pinicola (Basidiomycetes). Nordic Journal of Botany 17:319-329.

Nordén, B. and T. Appelqvist. 2001. Conceptual problems of ecological continuity and its bioindicators. Biodiversity and Conservation 10:779-791.

Nordén, B., T. Appelquist, B. Lindahl, and M. Henningsson. 1999. Cubic rot fungi - corticoid fungi in highly brown rotted spruce stumps. Mycologia Helvetica 10:13-24.

Nordén, B., M. Ryberg, F. Götmark, and B. Olausson. 2004. Relative importance of coarse and ﬁne woody debris for the diversity of wood-inhabiting fungi in temperate broadleaf forests. Biological Conservation 117:1-10.

Nordén, J., R. Penttilä, J. Siitonen, E. Tomppo, and O. Ovaskainen. 2013. Specialist species of wood-inhabiting fungi struggle while generalists thrive in fragmented forests. Journal of Ecology 101:701-712.

O’Brien, H.E., J.L. Parrent, J.A. Jackson, J.-M. Moncalvo, and R. Vilgalys. 2005. Fungal community analysis by large-scale sequencing of environmental samples. Applied and Environmental Microbiology 71:5544-5550

O’Donnell, K., N.S. Weber, S. Rehner, and G. Mills. 2003. Phylogeny and biogeography of

Morchella. In: Proceedings of the 22nd Fungal Genetics Conference, March 2003, Asilomar. Fungal Genetics Newsletter 50 (Suppl.) Abstract # 443.

Odor, P., J. Heilmann-Clausen, M. Christensen, E. Aude, K.W. Van Dort, A. Piltaver, I. Siller, M.T. Veerkamp, R. Walleyn, T. Standovar, et al. 2006. Diversity of dead wood inhabiting fungi and bryophytes in semi-natural beech forests in Europe. Biological Conservation 131:58-71.

Ohenoja, E. 1988. Behaviour of mycorrhizal fungi in fertilized soils. Karstenia 28:27-30.

Ohlson, M., L. Söderström, G. Hörnberg, O. Zackrisson, and J. Hermanssond. 1997. Habitat qualities versus long-term continuity as determinants of biodiversity in boreal old-growth swamp forests. Biological Conservation 81:221-231.

Økland, B. 1994. Mycetophilidae (Diptera), an insect group vulnerable to forestry practices? A comparison of clearcut, managed and semi-natural spruce forests in southern Norway. Biodiversity and Conservation 2:68-85.

Økland, B., A. Bakke, S. Hågvar, and T. Kvamme. 1995. What factors influence the diversity of saproxylic beetles? A multi-scaled study from a spruce forest in southern Norway. Biodiversity and Conservation 5:75-100.

Økland, B. 1996. Unlogged forests: important sites for preserving the diversity of Mycetophilids (Diptera: Sciaroidea). Biological Conservation 76:297-310.

Ontario Ministry of Natural Resources, 2001. Fire database. Unpublished manuscript. Aviation and Forest Fire Management Branch, Sault Ste. Marie, Ont.

Östlund, L. 1993. Exploitation and structural changes in the north Swedish boreal forest 1800- 1992. Dissertations in Forest Vegetation Ecology (No. 4). Swedish University of Agricultural Sciences, Umeå, 140 pp.

Östlund, L., O. Zackrisson, and A.-L. Axelsson. 1997. The history and transformation of a Scandinavian boreal forest landscape since the 19th century. Canadian Journal of Forest Research 27:1198-1206.

Paltto, H., B. Nordén, F. Götmark, and N. Franc. 2006. At which spatial and temporal scales does landscape context affect local density of Red Data Book and Indicator species? Biological Conservation 133:442-454.

Parent, S., M.-J. Simard, H. Morin, and C. Messier. 2003. Establishment and dynamics of the balsam fir seedling bank in old forests of northeastern Quebec. Canadian Journal of Forest Research 33:597-603.

Payette, S. 1992. Fire as a controlling process in the North American boreal forest. In: Shugart,

H.H., R. Leemans and G.B. Bonan. (Eds.), A systems analysis of the global boreal forest. Cambridge University Press, Cambridge, pp. 144-169.

Peay, K.G., P.G. Kennedy, and T.D. Bruns. 2008. Fungal community ecology: a hybrid beast with a molecular master. BioScience 58:799-810.

Pedlar, J.H., J.L. Pearce, L.A. Venier, and D.W. McKenney. 2002. Coarse woody debris in relation to disturbance and forest type in boreal Canada. Forest Ecology and Management 158:189-194.

Penttilä, R., M. Lindgren, O. Miettinen, H. Rita, and I. Hanski. 2006. Consequences of forest fragmentation for polyporous fungi at two spatial scales. Oikos 114:225-240.

Penttilä, R., J. Siitonen, and M. Kuusinen. 2004. Polypore diversity in managed and old-growth boreal Picea abies forests in southern Finland. Biology and Conservation 117:271-283.

Piascik, P.W. 2013. Responses of ground beetles (Coleoptera: Carabidae) to variation in woody debris supply in boreal northeastern Ontario. M.Sc. thesis. University of Toronto, 83 pp.

Pimm, S.L. and R.A. Askins. 1995. Forest losses predict bird extinctions in eastern North America. Proceedings of the National Academy of Sciences USA 92:9343-9347.

Rabinowitsch-Jokinen, R. and I. Vanha-Majamaa. 2010. Immediate effects of logging, mounding and removal of logging residues and stumps on coarse woody debris in managed boreal Norway spruce stands. Silva Fennica 44:51-62.

Rajala, T., M. Peltoniemi, T. Pennanen, and R. Mäkipää. 2012. Fungal community dynamics in relation to substrate quality of decaying Norway spruce (Picea abies [L.] Karst.) logs in boreal forests. FEMS Microbiology Ecology 81:494-505.

Rajala, T., T. Tuomivirta, T. Pennanen, and R. Mäkipää. 2015. Habitat models of wood- inhabiting fungi along a decay gradient of Norway spruce logs. Fungal Ecology 18:48- 55.

Rambaut, A. 2016. FigTree v1.4.3. Available from: http://tree.bio.ed.ac.uk/software/figtree/.

Ranius, T., P. Eliasson, and P. Johansson. 2008. Large-scale occurrence patterns of red-listed lichens and fungi on old oaks are influenced both by current and historical habitat density. Biodiversity and Conservation 17:2371-2381.

Rassi, P., I. Kanerva, and T. Mannerkoski. 2001. The 2000 Red List of Finnish Species. Ympäristöminiseriö and Suomen Ympäristökeskus, Helsinki.

Rassi, P., E. Hyvärinen, A. Juslén, and I. Mannerkoski. (Eds.) 2010. The 2010 Red List of Finnish Species. Ministry of the Environment and Finnish Environment Institute, Helsinki.

Rayner, A.D.M. and L. Boddy. 1988. Fungal decomposition of wood: its biology and ecology. Wiley, Chichester, 602 pp.

Read, D.J. and J. Perez-Moreno. 2003. Mycorrhizas and nutrient cycling in ecosystems - a journey towards relevance? New Phytologist 157:475-492.

Renvall, P. 1995. Community structure and dynamics of wood-rotting basidiomycetes on decomposing conifer trunks in northern Finland. Karstenia 35:1-51.

Riffell, S., J. Verschuyl, D. Miller, and T.B. Wigley. 2011. Biofuel harvests, coarse woody debris, and biodiversity - a meta-analysis. Forest Ecology and Management 261(4):878- 887.

Rosenzweig, M.L. 1995. Species diversity in space and time. Cambridge University Press, Cambridge, 460 pp.

Rowe, J.S. 1972. Forest Regions of Canada. J.S. Fisheries and Environment Canada, Canadian Forest Service, Headquarters, Ottawa, 172 pp.

Rudolphi, J. and L. Gustafsson. 2005. Effects of forest-fuel harvesting on the amount of deadwood on clearcuts. Scandinavian Journal of Forest Research 20:235-242.

Ruel, J.-C., R. Horvath, C.H. Ung, and A. Munson. 2004. Comparing height growth and biomass production of black spruce trees in logged and burned stands. Forest Ecology and Management 193:371-384.

Rypacek, V. 1977. Chemical composition of hemicelluloses as a factor participating in the substrate specificity of wood-destroying fungi. Wood Science and Technology 11:59-67.

Ryvarden, L. and R.L. Gilbertson. 1993. European polypores, Part 1. Synopsis Fungorum 6:1- 387.

Sato, H., S. Morimoto, and T. Hattori. 2012. A thirty-year survey reveals that ecosystem function of fungi predicts phenology of mushroom fruiting. PLoS ONE 7(11):e49777.

St. Hilaire, L.R. and D.J. Leopold. 1995. Conifer seedling distribution in relation to microsite conditions in a central New York forested minerotrophic peatland. Canadian Journal of Forest Research 25:261-269.

Ste-Marie, C. and D. Paré. 1999. Soil, pH and N availability effects on net nitrification in the forest floors of a range of boreal forest stands. Soil Biology and Biochemistry 31:1579- 1589.

Samuelsson, J., L. Gustafsson, and T. Ingelög. 1994. Dying and dead trees - a review of their importance for biodiversity. ArtDatabanken, SLU, Uppsala, 109 pp.

Sanders, F.E., B. Mosse, and P.B. Tinker. 1975. Endomycorrhizas. Academic Press, London.

Schoch, C.L., K.A. Seifert, A. Huhndorf, V. Robert, J.L. Spouge, C.A. Levesque, W. Chen, and 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 of the United States of America 109:6241-6246.

Schmit, J.P. 2005. Species-richness of tropical wood-inhabiting macrofungi provides support for species-energy theory. Mycologia 97:751-761.

Selikhovkin, A.V. 2005. Main disturbance factors in north-west Russian forests: structure and databases. Scandinavian Journal of Forest Research 20:27-32.

Sernander, R. 1936. Granskar och Fiby urskog. En studie over stormluckornas och marbuskarnas betydelse i den svenska granskogens regeneration. (The primitive forests of Granskär and Fiby: A study of the part played by storm-gaps and dwarf trees in the regeneration of the Swedish spruce forest). Acta Phytogeographica Suecica 8. Almqvist and Wiksell, Uppsala. (In Swedish.)

Setala, H. and M.A. McLean. 2004. Decomposition rate of organic substrates in relation to the species diversity of soil saprophytic fungi. Oecologia 139:98-107.

Siitonen, J. 1994. Decaying wood and saproxylic Coleoptera in two old spruce forests: a comparison based on two sampling methods. Annales Zoolgici Fennici 31:89-95.

Siitonen, J. 2001. Forest management, coarse woody debris and saproxylic organisms: Fennoscandian boreal forests as an example. Ecological Bulletins 49:11-42.

Siitonen, J., P. Martikainen, P. Punttila, and J. Rauh. 2000. Coarse woody debris and stand characteristics in mature managed and old-growth boreal mesic forests in southern Finland. Forest Ecology and Management 128:211-225.

Siitonen, P., A. Lehtinen, and M. Siitonen. 2005. Effects of forest edges on the distribution, abundance, and regional persistence of wood-rotting fungi. Conservation Biology 19:250-260.

Simard, S.W., K.J. Beiler, M.A. Bingham, J.R. Deslippe, L.J. Philip, and F.P. Teste. 2012. Mycorrhizal networks: mechanisms, ecology and modelling. Fungal Biology Reviews 26:39-60.

Similä, M., J. Kouki, M. Mönkkönen, A.L. Sippola, and E. Huhta. 2006. Co-variation and indicators of species diversity: can richness of forest-dwelling species be predicted in northern boreal forests? Ecological Indicators 6:686-700.

Sippola, A.-L., T. Lehesvirta, and P. Renvall. 2001. Effects of selective logging on coarse woody

debris and diversity of wood-decaying polypores in eastern Finland. Ecological Bulletins 49:243-254.

Sippola, A.-L., M. Mönkkönen, and P. Renvall. 2005. Polypore diversity in the herb-rich woodland key habitats of Koli national park in eastern Finland. Biological Conservation 126:260-269.

Sippola, A.-L. and P. Renvall. 1999. Wood-decomposing fungi and seed-tree cutting: a 40-year perspective. Forest Ecology and Management 115:183-201.

Sippola, A.-L., J. Siitonen, and R. Kallio. 1998. Amount and quality of coarse woody debris in natural and managed coniferous forests near the timberline in Finnish Lapland. Scandinavian Journal of Forest Research 13:204-214.

Sippola, A.-L., M. Similä, M. Mönkkönen, and J. Jokimäki. 2004. Diversity of polyporous fungi (Polyporaceae) in northern boreal forests: effects of forest site type and logging intensity. Scandinavian Journal of Forest Research 19:152-163.

Smith, M.E., G.W. Douhan, and D.M. Rizzo. 2007. Intraspecific and intrasporocarp ITS variation of ectomycorrhizal fungi as assessed by rDNA sequencing of sporocarps and pooled ectomycorrhizal roots from a Quercus woodland. Mycorrhiza 18:15-22.

Smith, S.E. and D.J. Read. 2008. Mycorrhizal symbiosis, 3rd ed. Academic Press, London, 800 pp.

Söderström, L. 1988a. Sequence of bryophytes and relation to substrate variables of decaying coniferous wood in northern Sweden. Nordic Journal of Botany 8:89-97.

Söderström, L. 1988b. The occurrence of epixylic bryophyte and lichen species in an old natural and a managed forest stand in northeastern Sweden. Biological Conservation 45:169-178.

Soil Classification Working Group. 1998. The Canadian System of Soil Classification, 3rd ed., Publication 1646. Research Branch, Agriculture and Agri-Food Canada, Ottawa, 187 pp.

Sousa, W.P. 1984. The role of disturbance in natural communities. Annual Review of Ecology, Evolution, and Systematics 15:353-391.

Speight, M.C.D. 1989. Saproxylic Invertebrates and their conservation. Council of Europe, Strasbourg, 79 pp.

Spies, T.A., J.F. Franklin, and T.B. Thomas. 1988. Coarse woody debris in Douglas-Fir forests of western Oregon and Washington. Ecology 69:1689-1702.

Steffen, K.T., M. Hofritchter, and A. Hatakka. 2002. Purification and characterization of manganese peroxidases from the litter-decomposing basidiomycetes Agrocybe praecox and Stropharia coronilla. Enzyme and Microbial Technology 30:550-555.

Stenlid, J., R. Penttilä, and A. Dahlberg. 2008. Wood-decay basidiomycetes in boreal forests: distribution and community development. In: Boddy, L., J.C. Frankland, and P. van West (Eds.), Ecology of Saprotrophic Basidiomycetes. Academic Press, U.K., pp. 239-262.

Stevens, V. 1997. The ecological role of coarse woody debris: an overview of the ecological importance of CWD in B.C. forests, Working Paper 30. Research Branch, BC Ministry of Forests, Victoria, BC, 21 pp.

Stocks, B.J. 1991. The extent and impact of forest fires in northern circumpolar countries. In: Levine, J.S. (Ed.), Global biomass burning: atmospheric, climatic, and biospheric implications, MIT Press, Cambridge, Mass., pp. 197-202.

Stocks, B.J., B.M. Wotton, M.D. Flannigan, M.A. Fosberg, D.R. Cahoon, and J.G. Goldammer. 2001. Boreal forest fire regimes and climate change. In: Beniston, M. and M.M. Verstraete (Eds.), Remote sensing and climate modeling: synergies and limitations. Kluwer Academic, Dordrecht, pp. 233-246.

Stokland, J.N., J. Siitonen, and B.G. Jonsson. 2012. Biodiversity in Dead Wood. Cambridge University Press, New York, N.Y., 524 pp.

Stokland, J.N. and H. Kauserud. 2004. Phellinus nigrolimitatus - a wood-decomposing fungus highly influenced by forestry. Forest Ecology and Management 187:333-343.

Stokland, J.N. and K.-H. Larsson. 2011. Legacies from natural forest dynamics: different effects of forest management on wood-inhabiting fungi in pine and spruce forests. Forest Ecology and Management 261:1707-1721.

Straatsma, G., F. Ayer, and S. Egli. 2001. Species richness, abundance, and phenology of fungal fruit bodies over 21 years in a Swiss forest plot. Mycological Research 105:515-523.

Straatsma, G. and I. Krisai-Greilhuber. 2003. Assemblage structure, species richness, abundance, and distribution of fungal fruit bodies in a seven year plot-based survey near Vienna. Mycological Research 107:632-640.

Sturtevant, B.R., J.A. Bissonette, J.N. Long, and D.W. Roberts. 1997. Coarse woody debris as a function of age, stand structure, and disturbance in boreal Newfoundland. Ecological Applications 7:702-712.

Sullivan, B.W., T.E. Kolb, S.C. Hart, J.P. Kaye, B.A. Hungate, S. Dore, and M. Montes-Helu. 2011. Wildfire reduces carbon dioxide efflux and increases methane uptake in ponderosa pine forest soils of the southwestern USA. Biogeochemistry 104:251-265.

Sverdrup-Thygeson, A. and D.B. Lindenmayer. 2003. Ecological continuity and assumed indicator fungi in boreal forest: the importance of the landscape matrix. Forest Ecology and Management 174:353-363.

Sverdrup-Thygeson, A. and F. Midtgaard. 1998. Fungus infected trees as islands in boreal forest: spatial distribution of the fungivorous beetles Bolitophagus reticulatus (Coleoptera, Tenebrionidae). Ecoscience 5:486-493.

Takahashi, K. 1994. Effect of size structure, forest floor type and disturbance regime on tree species composition in a coniferous forest in Japan. Journal of Ecology 82:769-773.

Talbot, J.M, S.D. Allison, and K.K. Treseder. 2008. Decomposers in disguise: mycorrhizal fungi as regulators of soil C dynamics in ecosystems under global change. Functional Ecology 22:955-963.

Tamura, K., G. Stecher, D. Peterson, A. Filipski, and S. Kumar. 2013. MEGA 6: molecular evolutionary genetics analysis version 6.0. Molecular Biology and Evolution 30:2725- 2729 .

Tarasov, M.E. 1999. Role of coarse woody debris in carbon balance of forest ecosystems of Leningrad Oblast. Ph.D. Thesis. St. Petersburg Forestry Research Institute, St. Petersburg, 21 pp. (In Russian.)

Taylor, A.F.S. 2002. Fungal diversity in ectomycorrhizal communities: sampling effort and species detection. Plant and Soil 244:19-28.

Tedersoo, L., M. Bahram, S. Põlme, U. Kõljalg, N.S. Yorou, R. Wijesundera, L.V. Ruiz, A.M. Vasco-Palacios, T.P. Quang, A. Suija et al. 2014. Global diversity and geography of soil fungi. Science 346(6213):1256688.

Tedersoo, L., T. Jairus, B.M. Horton, K. Abarenkov, T. Suvi, I. Saar, and U. Kõljalg. 2008. Strong host preference of ectomycorrhizal fungi in a Tasmanian wet sclerophyll forest as revealed by DNA barcoding and taxon-specific primers. New Phytologist 180:479-490.

Tedersoo, L., U. Kõljalg, N. Hallenberg, and K.-H. Larsson. 2003. Fine scale distribution of ectomycorrhizal fungi and roots across substrate layers including coarse woody debris in a mixed forest. New Phytologist 159:153-165.

Tedersoo, L. and M.E. Smith. 2013. Lineages of ectomycorrhizal fungi revisited: foraging strategies and novel lineages revealed by sequences from belowground. Fungal Biology Reviews 27:83-99.

Ter Braak, C.J.F. and P. Šmilauer. 2002. CANOCO reference manual and Cano Draw for Windows user’s guide: software for canonical community ordination (version 4.5). Microcomputer Power, Ithaca, N.Y.

Tikkanen, O.-P., P. Martikainen, E. Hyvärinen, K. Junninen, and J. Kouki. 2006. Red-listed boreal forest species of Finland: associations with forest structure, tree species, and decaying wood. Annales Zoolgici Fennici 43:373-383.

Tikkanen, O.-P., P. Punttila, and R. Heikkilä. 2009. Species-area relationships of redlisted species in old boreal forests: a large-scale data analysis. Diversity and Distributions 15:852-862.

Tofts, R.J. and P.D. Orton. 1998. The species accumulation curve for agarics and boleti from a Caledonian pinewood. Mycologist 12:98-102.

Toivanen, T., A. Markkanen, J.S. Kotiaho, and P. Halme. 2012. The effect of forest fuel harvesting on the fungal diversity of clear-cuts. Biomass and Bioenergy 39:84-93.

Tyrrell, L.E. and T.R. Crow. 1994. Structural characteristics of old-growth hemlock-hardwood forests in relation to age. Ecology 75:370-386.

United Nations International Year of Forests. 2011. [http://www.un.org/en/events/iyof2011/]

Uotila, A., J. Kouki, H. Kontkanen, and P. Pulkkinen. 2002. Assessing the naturalness of boreal forests in eastern Fennoscandia. Forest Ecology and Management 161:257-277.

Uotila, A., M. Maltamo, J. Uuttera, and A. Isomäki. 2001. Stand structure in semi-natural and managed forests in eastern Finland and Russian Karelia. Ecological Bulletins 49:149- 158.

Vanderwel, M.C., J.R. Malcolm, J.P. Caspersen, and M.A. Newman. 2010. Finescale habitat associations of red-backed voles in boreal mixedwood stands. Journal of Wildlife Management 74(7):1492-1501.

Van Wagner, C.E. 1968. The line intercept method in forest fuel sampling. Forest Science 14:20- 26.

Veblen, T.T. 1989. Tree regeneration responses to gaps along a transandean gradient. Ecology 70:541-543.

Vincent, J.S. and L. Hardy. 1977. L’évolution et l’extension des lacs glaciaires Barlow et Ojibway en territoire québécois. Géographie physique et Quaternaire, 31(3-4):357-372.

Virkkala, R. & Toivonen, H. 1999. Maintaining biological diversity in Finnish forests. Finnish Environment Institute, Helsinki, 56 pp.

Vogt, K.A., D.J. Vogt, H. Asbjornsen, and R.A. Dahlgren. 1995. Roots, nutrients and their relationship to spatial patterns. Plant and Soil 168-169:113-123.

Wardle, D.A. 2002. Communities and ecosystems: linking the aboveground and belowground components. Princeton University Press, Princeton, N.J., 400 pp.

Waring, R.H. and S.W. Running. 1998. Forest ecosystems: analysis at multiple scales. Academic

Press, San Diego, 370 pp.

Wästerlund, I. and T. Ingelög. 1981. Fruit body production of larger fungi in some young Swedish forests with special reference to logging waste. Forest Ecology and Management 3:269-294.

Watling, R. 1995. Assessment of fungal diversity: macromycetes, the problems. Canadian Journal of Botany 73:S15-S24.

Weber, M.G. and B.J. Stocks. 1998. Forest ﬁres in the boreal forests of Canada. In: Moreno, J.M. (Ed.), Large forest fires. Backhuys Publishers, Leiden, Netherlands, pp. 215-233.

Woodall, C.W. and M.S. Williams. 2005. Sampling, estimation, and analysis procedures for the down woody materials indicator. USDA Forest Service, General Technical Report, NC- 256, St. Paul, Minn.

Yatskov, M.A., O.N. Krankina, and M.E. Harmon. 2003. A chronosequence of wood decomposition in the boreal forests of Russia. Canadian Journal of Forest Research 33:1211-1226.

Deciduous Shrub Plot name Eastinga Northinga Treatmentb Year of origin (type) Ptc Abc Bpc Pbc Pac Pmc % DCA1d DCA2d densitye HL12CO 390543 5468620 CO 1943 (clearcut horse) 12 4 0 6 0 1 77 0.72 0.46 1.28 HL12FR 390962 5469129 FR 1943 (clearcut horse) 8 9 1 5 1 0 56 0.84 0.30 1.30 HL12HR 390350 5468634 HR 1943 (clearcut horse) 10 4 0 4 1 1 69 0.75 0.34 2.03 HL14CO 388848 5467906 CO 1951 (clearcut horse) 1 4 6 1 0 4 32 0.41 0.00 1.39 HL14FR 389361 5468216 FR 1951 (clearcut horse) 13 6 3 1 3 0 64 1.45 0.99 1.82 HL14HR 388180 5467152 HR 1959 (clearcut horse) 10 15 1 1 1 1 39 0.52 0.00 1.13 HL17CO 403826 5482174 CO 1944 (clearcut horse) 5 7 4 6 3 2 54 2.03 1.09 1.90 HL17FR 403866 5482431 FR 1944 (clearcut horse) 5 10 5 6 5 2 47 1.89 1.13 1.70 HL17HR 403810 5482654 HR 1944 (clearcut horse) 9 4 3 3 4 2 56 2.14 1.16 2.18 HL20CO 408325 5481274 CO 1955 (clearcut horse) 1 6 2 11 1 8 49 1.28 1.10 0.97 HL20FR 408486 5481479 FR 1955 (clearcut horse) 7 7 5 7 2 1 65 1.46 1.01 1.96 HL20HR 408545 5481773 HR 1955 (clearcut horse) 3 9 6 1 4 2 43 1.98 1.18 2.08 HL24CO 401500 5463773 CO 1946 (clearcut horse) 7 8 5 3 1 1 62 1.10 0.45 1.54 HL24FR 400923 5462448 FR 1946 (clearcut horse) 4 12 7 1 3 7 36 1.04 0.62 0.87 HL24HR 400645 5462325 HR 1946 (clearcut horse) 6 10 5 1 1 2 49 0.81 0.34 0.92 ML04CO 393633 5431015 CO 1971 (clearcut skidder) 11 13 4 2 2 0 54 1.95 1.16 1.32 ML04FR 394160 5432666 FR 1975 (clearcut skidder) 9 11 3 5 4 0 53 2.07 1.19 1.29 ML04HR 394093 5431927 HR 1975 (clearcut skidder) 9 10 5 1 6 1 48 1.72 1.16 1.57 ML10CO 358379 5440608 CO 1973 (clearcut skidder) 18 3 0 0 2 2 74 0.98 1.25 1.90 ML10FR 358816 5440093 FR 1973 (clearcut skidder) 7 4 2 6 0 1 73 0.37 1.92 1.43 ML10HR 358675 5440234 HR 1973 (clearcut skidder) 17 2 2 3 0 0 89 0.51 1.73 1.41 ML11CO 385823 5465286 CO 1967 (clearcut skidder) 2 7 1 1 1 9 20 0.84 0.82 1.39 ML11FR 385598 5465203 FR 1967 (clearcut skidder) 2 14 1 2 7 2 19 1.58 0.90 0.75 ML11HR 385308 5464992 HR 1967 (clearcut skidder) 7 5 3 2 0 5 60 1.50 0.82 1.17 ML29CO 402676 5441380 CO 1972 (clearcut skidder) 5 3 1 1 1 8 36 0.00 1.24 1.18 ML29FR 403160 5441727 FR 1972 (clearcut skidder) 3 10 3 2 1 5 37 1.67 1.09 1.68 ML29HR 402961 5441494 HR 1972 (clearcut skidder) 4 6 1 2 1 4 43 0.69 1.77 2.16 UL01G1 373690 5401181 UL 1851 (fire) 5 4 5 1 6 0 46 2.12 1.06 2.07 UL01PG 373497 5401560 UL 1851 (fire) 8 3 3 1 1 4 48 2.17 1.02 2.00 UL03G1 373831 5399387 UL 1891 (fire) 10 4 7 0 6 3 56 1.61 0.75 1.62 UL03PG 373618 5399704 UL 1891 (fire) 3 4 7 2 0 8 49 1.83 0.79 1.33 UL06G1 385034 5395626 UL 1891 (fire) 3 6 12 1 4 0 46 2.28 1.21 1.87 UL06PG 384770 5396058 UL 1851 (fire) 0 2 6 4 1 2 34 2.20 0.82 1.18 UL07G1 384446 5395205 UL 1851 (fire) 4 4 7 8 4 0 55 2.21 1.17 1.18 UL07PG 384680 5394833 UL 1866 (fire) 8 11 7 1 0 0 57 2.11 1.24 2.99 UL20G1 403545 5486624 UL 1924 (fire) 14 4 0 0 7 2 53 1.19 0.56 1.06 UL20PG 403372 5486743 UL 1924 (fire) 17 5 2 0 8 0 58 2.06 0.89 1.34 a Datum = NAD83 b CO = logged, control; FR =logged, full coarse woody debris removal; HR = logged, one-half coarse woody debris removal; UL = unlogged. c Basal areas in m2/ha for trees ≥7 cm diameter at breast height (Pt = Populus tremoloides, Ab = Abies balsamea, Bp = Betula papyrifera, Pb = Populus balsamea, Pa = Picea albicans, Pm = Picea mariana). d Axes one and two scores from detrended correspondence analysis of shrub communities. e Number of stems per m2.

Sampling period 1 2 3

(June 20-31) (July 26-31) (Aug 19-28) First canonical axis 0.0009 n.s. n.s. All canoncial axes 0.0003 0.0001 n.s. Small, early decay, coniferous CWD n.s. n.s. n.s. Small, late decay, coniferous CWD n.s. n.s. n.s. Large, early decay, coniferous CWD 0.041 n.s. n.s. Large, late decay, coniferous CWD 0.0158 n.s. n.s. Small, early decay, deciduous CWD n.s. n.s. n.s. Small, late decay, deciduous CWD 0.0035 n.s. n.s. Large, early decay, deciduous CWD 0.0002 0.0253 n.s. Large, late decay, deciduous CWD 0.0028 0.0007 n.s. Total CWD 0.0001 n.s. 0.0032 DCA1 0.0028 0.0232 0.0002 DCA2 0.005 n.s n.s. Percent deciduous 0.0009 0.0447 0.0124 Shrub density n.s. n.s. n.s.

Table 3. Mean (± SE) species richness in each of four ecological guilds and the total number of morphospecies across treatment plots during three sampling periods in mixedwood forests of boreal northeastern Ontario.

Treatment FR HR CO UL p-value Sample 1 ECM 0.44 + 0.24a 0.11 + 0.11a 0.44 + 0.24a 0.3 + 0.15a n.s. 2 WS 9.22 + 1.57a 11.56 + 1.04a 14.11 + 0.98a 14.5 + 1.82a 0.04591 LS 3.33 + 0.78a 4.44 + 0.77a,b 3.89 + 0.77a 8.00 + 1.35b 0.00671 PARA 0.67 + 0.29 1.11 + 0.35 1.00 + 0.24 1.30 + 0.42 n.s. 2 Total 11.00 + 2.03a 14.00 + 1.29a,b 16.33 + 1.08a,b 19.22 + 2.30b 0.01791 Sample 2 ECM 6.00 + 1.63 6.22 + 1.28 5.22 + 0.85 3.90 + 0.75 n.s. 1 WS 12.89 + 1.81a 16.11 + 2.05a,b 13.56 + 1.54a,b 19.80 + 1.82b 0.04081 LS 14.33 + 2.20 13.33 + 2.35 13.11 + 2.26 14.70 + 1.78 n.s. 1 PARA 1.33 + 0.29 1.56 + 0.50 1.11 + 0.35 2.30 + 0.67 n.s. 2 Total 30.78 + 4.93 30.78 + 4.62 29.78 + 3.60 36.30 + 3.39 n.s. 1 Sample 3 ECM 21.78 + 3.95 18.67 + 3.08 18.00 + 2.57 21.00 + 1.98 n.s. 1 WS 14.89 + 2.12a 15.33 + 2.09a 17.56 + 1.43a,b 25.60 + 2.92b 0.00531 LS 15.67 + 3.16a 17.78 + 2.60a 11.89 + 1.37a 30.40 + 4.66b 0.00181 PARA 2.44 + 0.44 1.67 + 0.44 3.00 + 0.53 3.20 + 0.63 n.s. 1 Total 49.67 + 7.95a,b 49.44 + 6.22a,b 45.11 + 3.89a 71.20 + 7.69b 0.03521

1 Results from ANOVAs; n.s. = not significant (p > 0.05). 2 Results from Median tests; n.s. = not significant (p > 0.05). 3 FR = full-removal, HR = half-removal, CO = control, UL = unlogged 4ECM = ectomycorrhizal; WS = wood saprotrophs; LS = litter saprotrophs; PARA = parasites; Total = total richness 5 Letters in common indicate no significant difference between treatment means (Tukey’s HSD, α=0.05).

Figure 1. Canonical correspondence analyses conducted on macrofungal species assemblages in unlogged and logged plots in mixedwood forests of northeastern Ontario from sampling periods 1 (upper), 2 (mid) and 3 (lower). Asterisks denote environmental variables with significant effects in constraining community composition (* = p<0.05, ** = p<0.01, *** = p<0.005, **** = p<0.001, ***** = p<0.0005). vtot = total CWD volume; vlc_p_u1 = large-diameter, early-decay conifer CWD; vlc_p_u3 = large-diameter, late-decay conifer CWD; vld_p_u1 = large- diameter, early-decay deciduous CWD; vld_p_u3 = large-diameter, late-decay deciduous CWD; vsc_p_u1 = small- diameter, early-decay conifer CWD; vsc_p_u3 = small-diameter, late-decay conifer CWD; vsd_p_u1 = small-diameter, early-decay deciduous CWD; vsd_p_u3 = small-diameter, late-decay deciduous CWD; perc_dec = percent basal area of deciduous trees; shrub_st = shrub stem density; DCA1 = light exposure and conifer composition gradient (low end = shade-intolerant, high end = shade-tolerant); DCA2 = soil moisture gradient (low end = hydric conditions, high end = mesic conditions).

100

101

a a a,b

Figure 3. Box plot showing average species richness of wood saprotrophs in the four treatments (FR = full CWD removal; HR = half CWD removal; CO = control (no CWD removed); UL = unlogged) in mixedwood forests of northeastern Ontario sampled in the first sampling period. F- value and p-value (upper left) are from an analysis of variance; letters in common indicate no significant difference between treatment means (Tukey’s HSD, a=0.05).

102

Figure 4. Redundancy analyses conducted on macrofungal ecological guilds in unlogged and logged plots in mixedwood forests of northeastern Ontario from sampling periods 1 (upper), 2 (mid) and 3 (lower). ECM = ectomycorrhizal morphospecies, litter_sap = litter saprotrophs, para = parasites, total_species = total number of morphospecies, wood_sap = wood saprotrophs. Asterisks denote environmental variables with significant effects in constraining community composition (* = p<0.05, ** = p<0.01, *** = p<0.005, **** = p<0.001, ***** = p<0.0005). vtot = total CWD volume; vlc_p_u1 = large-diameter, early-decay conifer CWD; vlc_p_u3 = large-diameter, late-decay conifer CWD; vld_p_u1 = large- diameter, early-decay deciduous CWD; vld_p_u3 = large-diameter, late-decay deciduous CWD; vsc_p_u1 = small- diameter, early-decay conifer CWD; vsc_p_u3 = small-diameter, late-decay conifer CWD; vsd_p_u1 = small-diameter, early-decay deciduous CWD; vsd_p_u3 = small-diameter, late-decay deciduous CWD; perc_dec = percent basal area of deciduous trees; shrub_st = shrub stem density; DCA1 = light exposure and conifer composition gradient (low end = shade-intolerant, high end = shade-tolerant); DCA2 = soil moisture gradient (low end = hydric conditions, high end = mesic conditions).

103

Appendix I. ITS sequences of fruiting bodies sampled from unlogged and logged plots in boreal mixedwood forests near Kapuskasing, Ontario. The collection number, phylum and morphospecies name are listed, as well as the top BLAST hit from GenBank or UNITE, its accession number, species name and country of origin.

Collection # Morphospecies Max Accession # Top GenBank or UNITE hit Top hit country identity %

Ascomycota

J36 Byssonectria terrestris 100 KJ619952.1 Byssonectria terrestris Sweden M148 Hymenoscyphus sp.1 93 NR_154907.1 Hymenoscyphus aurantiacus China X146 Lachnellula cf. calyciformis(1) 99 KY777393.1 Lachnellula calyciformis USA J341 Lachnellula cf. calyciformis(2) 99 KY777393.1 Lachnellula calyciformis USA W75 Otidea cf. mirabilis 98 KM010095.1 Otidea mirabilis Finland R734 Otidea cf. tuomikoskii 99 JN942776.1 Otidea tuomikoskii Norway X1182 Rutstroemiaceae sp.1 95 KF588371.1 Rutstroemia echinophila Spain

Basidiomycota

R111_ITS1F Agaricus cf. friesianus 99 KF447907.1 Agaricus friesianus Sweden J198 Agrocybe cf. praecox 99 UDB023779 Agrocybe elatella Estonia R480 Amanita cf. sinicoflava 100 KJ638263.1 Amanita sinicoflava Canada B545 Aphroditeola olida 99 KM248880.1 Aphroditeola olida Canada X1215 Armillaria aff. sinapina 90 FJ664609.1 Armillaria sinapina USA R510 Armillaria cf. ostoyae(1) 99 KT822292.1 Armillaria ostoyae China L130 Armillaria cf. ostoyae(2) 99 KT822292.1 Armillaria ostoyae China X379 Arrhenia aff. epichysium 97 KR673630.1 Arrhenia sp. South Korea M32 Baeospora myriadophylla 100 UDB015456 Baeospora myriadophylla Estonia J79 Bolbitiaceae sp.1 80 HQ840656.1 Bolbitius viscosus USA J252 Cerioporus cf. varius(1) 98 KX524509.1 Polyporus varius Serbia F97 Cerioporus cf. varius(2) 98 KX524509.1 Polyporus varius Serbia B667 Cerioporus cf. varius(3) 100 MG748571.1 Polyporus varius USA X228 Cerrena unicolor 100 JQ031127.1 Cerrena unicolor Sweden C668 Cheimonophyllum candidissimum(1) 98 FJ596811.1 Cheimonophyllum sp. USA R880 Cheimonophyllum candidissimum(2) 99 FJ596811.1 Cheimonophyllum sp. USA C196 Clavariadelphus cf. sachalinensis 99 EU834196.1 Clavariadelphus sachalinensis USA W55 Clavulina cf. coralloides 99 MH542542.1 Clavulina reae Mexico W49 Clavulina cf. rugosa(1) 99 EU862210.1 Clavulina cf. rugosa Finland B772U Clavulina cf. rugosa(2) 100 KM248920.1 Clavulina rugosa Canada B91 Clavulinopsis cf. corniculata 96 UDB011577 Clavulinopsis corniculata Estonia R772 Clitocybe odora(1) 99 UDB011154 Clitocybe odora Estonia R919 Clitocybe odora(2) 100 UDB011154 Clitocybe odora Estonia X431U Clitocybe odora(3) 100 UDB011154 Clitocybe odora Estonia X710 Clitocybula aff. familia 99 KX897414.1 Clitocybula sp. Canada B49 Collybia cookei(1) 99 DQ830806.1 Collybia cookei USA B479 Collybia cookei(2) 100 DQ830806.1 Collybia cookei USA F139 Collybia tuberosa(1) 100 UDB015205 Collybia tuberosa Estonia L17 Collybia tuberosa(2) 99 UDB015205 Collybia tuberosa Estonia V438 Collybia tuberosa(3) 100 DQ830807.1 Collybia tuberosa USA X928 Coltricia perennis 99 UDB014110 Coltricia perennis Estonia L115 Conocybe cf. siennophylla 97 JX968240.1 Conocybe cylindracea Italy R11 Coprinellus cf. radians 100 KU761146.1 Coprinellus radians Canada X1153 Cortinarius aff. barbarorum 95 NR_130238.1 Cortinarius olympianus USA X67 Cortinarius aff. boreidionysae(1) 98 KF732489.1 Cortinarius boreidionysae Finland X80 Cortinarius aff. boreidionysae(2) 98 KF732489.1 Cortinarius boreidionysae Finland X457 Cortinarius aff. delibutus 99 KP406542.1 Cortinarius sp. Canada R583 Cortinarius aff. dolabratus(1) 96 LC373241.1 Cortinarius sp. Japan

104

Collection # Morphospecies Max Accession # Top GenBank or UNITE hit Top hit country identity %

R45 Cortinarius aff. dolabratus(2) 96 LC373241.1 Cortinarius sp. Japan R360 Cortinarius aff. dolabratus(3) 96 LC373241.1 Cortinarius sp. Japan F145 Cortinarius aff. glaucopus 99 KJ421201.1 Cortinarius subfoetens Germany R699 Cortinarius aff. helvelloides 99 AY669684.1 Cortinarius helvelloides Germany R724 Cortinarius aff. laetissimus 98 GQ159898.1 Cortinarius laetissimus Canada R551 Cortinarius aff. privignipallens 98 KP165569.1 Cortinarius privignipallens Sweden R584 Cortinarius aff. xanthocephalus 99 DQ097877.1 Cortinarius alboviolaceus Canada B68 Cortinarius albocyaneus 99 KX302206.1 Cortinarius albocyaneus Sweden W33 Cortinarius argenteolilacinus 99 MH923077.1 Cortinarius argenteolilacinus var. dovrensis Norway W46 Cortinarius centrirufus 100 KP165578.1 Cortinarius centrirufus Finland R568A Cortinarius cf. balaustinus 98 UDB018299 Cortinarius saturatus France F50 Cortinarius cf. caesioarmeniacus(1) 99 MF139753.1 Cortinarius caesioarmeniacus Sweden R261 Cortinarius cf. caesioarmeniacus(2) 99 MF139753.1 Cortinarius caesioarmeniacus Sweden R927 Cortinarius cf. casimiri(1) 99 HQ604719.1 Cortinarius casimiri Canada X817 Cortinarius cf. casimiri(2) 100 HQ604719.1 Cortinarius casimiri Canada X116 Cortinarius cf. comptulus(1) 99 KR019806.1 Cortinarius sp. Latvia X160 Cortinarius cf. comptulus(2) 99 KR019806.1 Cortinarius sp. Latvia R655 Cortinarius cf. decipiens(1) 99 GQ159796.1 Cortinarius alnetorum Canada X501 Cortinarius cf. decipiens(2) 99 GQ159796.1 Cortinarius alnetorum Canada X1197 Cortinarius cf. decipiens(3) 99 GQ159796.1 Cortinarius alnetorum Canada X1231 Cortinarius cf. decipiens(4) 100 GQ159796.1 Cortinarius alnetorum Canada X1219 Cortinarius cf. decipiens(5) 100 GQ159796.1 Cortinarius alnetorum Canada R43 Cortinarius cf. decipiens(6) 99 GQ159796.1 Cortinarius alnetorum Canada X963 Cortinarius cf. depressus 99 UDB002292 Cortinarius depressus Sweden B111 Cortinarius cf. diasemospermus 100 JQ724020.1 Cortinarius diasemospermus Sweden X41 Cortinarius cf. fulvescens 99 UDB018657 Cortinarius fulvescens Estonia R509 Cortinarius cf. impennoides 100 KT591604.1 Cortinarius impennoides Finland R578 Cortinarius cf. infractus(1) 100 KJ421115.1 Cortinarius infractus Germany R361 Cortinarius cf. infractus(2) 99 KJ421115.1 Cortinarius infractus Germany R123 Cortinarius cf. lucorum 99 AY695794.1 Cortinarius lucorum USA W1 Cortinarius cf. malicorius 99 JX045668.1 Cortinarius malicorius Finland R247 Cortinarius cf. obtusus 99 GQ159780.1 Cortinarius aff. rigens Canada P67 Cortinarius cf. paragaudis 100 HQ845156.1 Cortinarius paragaudis France R183 Cortinarius cf. polaris 99 KX355537.1 Cortinarius croceus Poland P71 Cortinarius cf. renidens(1) 98 UDB002178 Cortinarius renidens Sweden X326 Cortinarius cf. renidens(2) 99 UDB002178 Cortinarius renidens Sweden R529_ITS4 Cortinarius cf. rusticus(1) 100 UDB036993 Cortinarius rusticus Norway R558 Cortinarius cf. rusticus(2) 100 UDB036993 Cortinarius rusticus Norway X1024 Cortinarius cf. saniosus 99 DQ102681.1 Cortinarius saniosus Sweden B360 Cortinarius cf. scandens 99 GQ159849.1 Cortinarius scandens Canada X314 Cortinarius cf. subbalaustinus 99 JX407335.1 Cortinarius subbalaustinus Finland X9 Cortinarius cf. subcroceofolius(1) 99 MK069541.1 Cortinarius sp. Canada X27 Cortinarius cf. subcroceofolius(2) 99 MK069541.1 Cortinarius sp. Canada L324 Cortinarius cf. tabularis 99 UDB034771 Cortinarius sp. Estonia X336 Cortinarius cf. trivialis(1) 99 DQ295109.1 Cortinarius trivialis Austria R198 Cortinarius cf. trivialis(2) 99 UDB011737 Cortinarius trivialis Estonia R181 Cortinarius cf. trivialis(3) 99 UDB011737 Cortinarius trivialis Estonia X961 Cortinarius cyanites 100 UDB001154 Cortinarius cyanites Sweden M718 Cortinarius rubellus(1) 100 AY669595.1 Cortinarius rubellus Sweden X86 Cortinarius rubellus(2) 100 AY669595.1 Cortinarius rubellus Sweden W13 Cortinarius sphagnophilus 100 KJ421145.1 Cortinarius sphagnophilus Germany X62 Cortinarius spilomeus(1) 99 NR_153075.1 Cortinarius spilomeus Sweden X1185 Cortinarius spilomeus(2) 99 NR_153075.1 Cortinarius spilomeus Sweden

105

Collection # Morphospecies Max Accession # Top GenBank or UNITE hit Accession # identity country %

R678 Cortinarius vibratilis 100 EU821696.1 Cortinarius vibratilis Canada W43 Crepidotus aff. cesatii(1) 100 MF461345.1 Crepidotus sp. China X342 Crepidotus aff. cesatii(2) 99 MF461345.1 Crepidotus sp. China R283 Crepidotus aff. luteolus(1) 94 JF907961.1 Crepidotus subverrucisporus Italy B638 Crepidotus aff. luteolus(2) 95 MF461345.1 Crepidotus sp. China J11 Crepidotus calolepis(1) 100 UDB019553 Crepidotus calolepis Estonia J164 Crepidotus calolepis(2) 100 UDB019553 Crepidotus calolepis Estonia B695 Crepidotus cf. versutus(1) 97 JF907961.1 Crepidotus subverrucisporus Italy M389 Crepidotus cf. versutus(2) 97 JF907961.1 Crepidotus subverrucisporus Italy X256 Crepidotus cf. versutus(3) 97 JF907961.1 Crepidotus subverrucisporus Italy C572 Crepidotus cf. versutus(4) 97 JF907961.1 Crepidotus subverrucisporus Italy X844 Crepidotus cf. versutus(5) 96 JF907961.1 Crepidotus subverrucisporus Italy B413 Crinipellis campanella(1) 86 DQ486708.1 Gymnopus contrarius USA P450 Crinipellis campanella(2) 86 DQ486708.1 Gymnopus contrarius USA C298 Crinipellis piceae 99 UDB018815 Crinipellis setipes Canada B483 Cyclocybe cf. erebia(1) 99 UDB011453 Agrocybe erebia Estonia B674 Cyclocybe cf. erebia(2) 99 UDB011453 Agrocybe erebia Estonia W71 Cyclocybe cf. erebia(3) 99 UDB011453 Agrocybe erebia Estonia B812U Cyptotrama cf. chrysopepla 99 MH910552.1 Cyptotrama chrysopepla USA W82 Dermoloma sp.1 96 KU058497.1 Dermoloma sp. Slovakia R100 Entoloma aff. llimonae(1) 98 JX454832.1 Entoloma llimonae Spain P193 Entoloma aff. llimonae(2) 98 JN021022.1 Entoloma sp. Canada B35 Entoloma aff. serrulatum 97 KC581341.1 Leptonia serrulata Canada B166U Entoloma cf. dysthales(1) 99 LN850553.1 Entoloma dysthales Finland B78U Entoloma cf. dysthales(2) 99 LN850553.1 Entoloma dysthales Finland L197 Entoloma cf. dysthales(3) 99 LN850553.1 Entoloma dysthales Finland M341 Entoloma cf. elodes 86 UDB000927 Entoloma sp. Iceland X935U Entoloma cf. griseum 99 KJ705163.1 Entoloma griseum Canada B648 Entoloma cf. minutum(1) 98 JX454829.1 Entoloma aff. minutum Spain P265 Entoloma cf. minutum(2) 98 JX454829.1 Entoloma aff. minutum Spain P267 Entoloma cf. minutum(3) 98 JX454829.1 Entoloma aff. minutum Spain V243 Entoloma cf. olivaceotinctum(1) 86 KP191922.1 Richoniella sp. Australia X1206 Entoloma cf. olivaceotinctum(2) 87 KJ001429.1 Entoloma aff. iodiolens Spain B159 Entoloma sp.1(1) 98 JN021024.1 Entolomataceae sp. Canada B745 Entoloma sp.1(2) 98 JN021024.1 Entolomataceae sp. Canada X611 Entoloma sp.1(3) 98 JN021024.1 Entolomataceae sp. Canada X686 Entoloma sp.1(4) 98 JN021024.1 Entolomataceae sp. Canada X601 Entoloma sp.2 92 MF773595.1 Entoloma sp. USA M542 Flammulaster cf. granulosus 93 MF954833.1 Flammulaster sp. Canada L6 Flammulaster cf. rhombosporus(1) 93 MF954833.1 Flammulaster sp. Canada R457 Flammulaster cf. rhombosporus(2) 91 MF954833.1 Flammulaster sp. Canada M31 Fomitiporia cf. punctata 99 LC333732.1 Fomitiporia punctata Japan X1179A Galerina aff. stylifera 98 MF954860.1 Galerina cf. camerina Canada L247 Galerina cf. mniophila 100 AJ585459.1 Galerina mniophila Norway W21 Galerina cf. triscopa 99 KY744148.1 Galerina triscopa USA L110 Galerina cf. vittiformis 99 KY230504.1 Galerina sp. China J110 Gloeophyllum sepiarium 100 UDB018824 Gloeophyllum sepiarium Canada X729 Gymnopilus cf. bellulus 99 MF039254.1 Gymnopilus bellulus Germany M262 Gymnopus aff. westii 99 KY026691.1 Gymnopus sp. USA X806 Gymnopus cf. alkalivirens 99 DQ480112.1 Gymnopus alkalivirens Greenland B450 Gymnopus cf. androsaceus 99 KY026749.1 Gymnopus androsaceus Canada J54 Gymnopus cf. contrarius(1) 99 DQ486708.1 Gymnopus contrarius USA R675 Gymnopus cf. contrarius(2) 99 DQ486708.1 Gymnopus contrarius USA J210 Gymnopus cf. dryophilus 99 KT874997.1 Gymnopus dryophilus Mexico

106

Collection # Morphospecies Max Accession # Top GenBank or UNITE hit Accession # identity country %

X36U Gymnopus cf. perforans 99 KY026743.1 Gymnopus perforans subsp. transatlanticus Canada M15 Gymnopus cf. subsulphureus(1) 99 AY256693.1 Gymnopus junquilleus USA J298_ITS1F Gymnopus cf. subsulphureus(2) 100 KF007938.1 Gymnopus junquilleus USA J295 Gymnopus cf. subsulphureus(3) 99 AY256693.1 Gymnopus junquilleus USA M246 Gymnopus cf. subsulphureus(4) 99 AY256693.1 Gymnopus junquilleus USA C495 Gymnopus confluens(1) 99 KP710272.1 Gymnopus confluens USA B652 Gymnopus confluens(2) 99 KP710272.1 Gymnopus confluens USA B602 Gymnopus confluens(3) 100 KP710272.1 Gymnopus confluens USA P294 Hapalopilus rutilans 100 KX752622.1 Hapalopilus rutilans Finland F112 Hebeloma sp.1 99 UDB037449 Hebeloma sordescens Norway X264_ITS4 Hebeloma sp.2 100 KT217496.1 Hebeloma ingratum Finland J149 Hemimycena aff. gracilis 92 MK169368.1 Hemimycena albicolor USA R449 Hemimycena cf. mairei 93 MH856249.1 Hemimycena mairei France B478 Hemimycena cf. pseudolactea(1) 99 UDB017907 Hemimycena pseudolactea Estonia W47 Hemimycena cf. pseudolactea(2) 99 UDB017907 Hemimycena pseudolactea Estonia L202 Hemimycena cf. pseudolactea(3) 99 UDB017907 Hemimycena pseudolactea Estonia X811 Hemimycena cf. pseudolactea(4) 98 UDB017907 Hemimycena pseudolactea Estonia X935L Hemimycena cf. pseudolactea(5) 98 UDB017907 Hemimycena pseudolactea Estonia M64 Heteroradulum cf. kmetii 99 KX262109.1 Heteroradulum kmetii Canada X372_ITS1F Homophron cf. camptopodum 99 UDB015582 Homophron camptopodum Estonia J215 Homophron cf. spadiceum 99 MG773815.1 Homophron spadiceum USA R6 Hydnopolyporus cf. fimbriatus(1) 97 MG231801.1 Porotheleum fimbriatum China R122 Hydnopolyporus cf. fimbriatus(2) 99 UDB031596 Porotheleum fimbriatum Finland X590 Hydnum cf. albertense 98 NR_158492.1 Hydnum albertense Canada W25 Hydnum cf. melitosarx 100 KX388686.1 Hydnum cf. melitosarx Canada L185 Hygrocybe cf. cantharellus(1) 99 KM248890.1 Hygrocybe cantharellus Canada M534 Hygrocybe cf. cantharellus(2) 99 MF156249.1 Hygrocybe calciphila Mexico X1210 Hygrocybe cf. cantharellus(3) 99 KM248890.1 Hygrocybe cantharellus Canada R39 Hygrocybe cf. parvula 99 KY744189.1 Hygrocybe parvula USA M61 Hypholoma capnoides(1) 100 UDB023650 Hypholoma capnoides Estonia M8 Hypholoma capnoides(2) 100 UDB023650 Hypholoma capnoides Estonia B456 Infundibulicybe sp.1(1) 99 UDB019582 Clitocybe metachroa Estonia B473 Infundibulicybe sp.1(2) 99 UDB019582 Clitocybe metachroa Estonia F136 Inocybe aff. proximella 99 UDB023652 Inocybe proximella Estonia X317 Inocybe aff. rimosa 99 MH024853.1 Inocybe holoxantha Canada X757 Inocybe cf. calamistrata(1) 98 AM882947.2 Inocybe calamistrata Norway P744 Inocybe cf. calamistrata(2) 98 UDB015568 Inocybe calamistrata Estonia R199U Inocybe cf. flocculosa 99 JF908124.1 Inocybe flocculosa Italy W81 Inocybe cf. griseolilacina 99 MH578030.1 Inocybe griseolilacina USA V315 Inocybe cf. jacobi(1) 98 HQ604804.1 Inocybe jacobi Canada W18AU Inocybe cf. jacobi(2) 98 HQ604804.1 Inocybe jacobi Canada W38 Inocybe cf. jacobi(3) 98 HQ604804.1 Inocybe jacobi Canada R811 Inocybe cf. jacobi(4) 98 HQ604804.1 Inocybe jacobi Canada R177 Inocybe cf. lanuginosa(1) 97 HQ604316.1 Inocybe lanuginosa var. lanuginosa Canada B873 Inocybe cf. lanuginosa(2) 99 HQ604307.1 Inocybe lanuginosa var. ovatocystis Canada R321 Inocybe cf. lanuginosa(3) 99 HQ604307.1 Inocybe lanuginosa var. ovatocystis Canada X110 Inocybe cf. lanuginosa(4) 99 JF908137.1 Inocybe leptophylla Italy X52A Inocybe cf. leptocystis 99 MF797857.1 Inocybe leptocystis USA X232 Inocybe cf. maculata 100 JX030262.1 Inocybe maculata USA L253 Inocybe cf. mixtilis(1) 100 MH578018.1 Inocybe sp. Mexico X1111 Inocybe cf. mixtilis(2) 99 KX290817.1 Inocybe occulta Germany X101 Inocybe cf. mixtilis(3) 99 KX290822.1 Inocybe occulta Austria X10 Inocybe cf. nitidiuscula(1) 99 HQ604087.1 Inocybe cf. pruinosa Canada

107

Collection # Morphospecies Max Accession # Top GenBank or UNITE hit Accession # identity country %

X20 Inocybe cf. nitidiuscula(2) 99 UDB015345 Inocybe sp. Estonia R517 Inocybe cf. nitidiuscula(3) 99 HQ604149.1 Inocybe flocculosa var. flocculosa Canada X1033 Inocybe cf. ochroalba 99 HQ604527.1 Inocybe splendens var. phaeoleuca Canada R239 Inocybe cf. pallidicremea 99 MG429701.1 Inocybe pallidicremea USA X234 Inocybe cf. pseudodestricta(1) 99 HQ604514.1 Inocybe glabrescens Canada X968 Inocybe cf. pseudodestricta(2) 99 HQ604514.1 Inocybe glabrescens Canada R723 Inocybe cf. pseudodestricta(3) 99 HQ604514.1 Inocybe glabrescens Canada X46 Inocybe cf. squarrosoides(1) 100 MH578003.1 Inocybe minima Canada X688 Inocybe cf. squarrosoides(2) 99 MH578003.1 Inocybe minima Canada X218 Inocybe cf. whitei 99 UDB017935 Inocybe whitei Estonia R348 Inocybe sp.1(1) 92 KY990544.1 Inocybe aff. geophylla USA R882 Inocybe sp.1(2) 86 KP406546.1 Inocybe geophylla Canada L196 Inocybe sp.2 91 MF804316.1 Inocybe ionochlora Germany M70 Kuehneromyces cf. lignicola 99 UDB036255 Kuehneromyces lignicola Norway R304 Laccaria cf. laccata(1) 100 JN942782.1 Laccaria cf. laccata Japan R610 Laccaria cf. laccata(2) 100 JN942782.1 Laccaria cf. laccata Japan M543 Laccaria cf. laccata(3) 100 JN942782.1 Laccaria cf. laccata Japan X193 Laccaria cf. striatula 99 MH979278.1 Laccaria striatula USA B704 Laccaria nobilis(1) 100 UDB018822 Laccaria nobilis Canada L34 Laccaria nobilis(2) 100 UDB018822 Laccaria nobilis Canada X933 Laccaria sp.1 99 KM067827.1 Laccaria sp. USA B224 Laccaria sp.2(1) 99 UDB001466 Laccaria laccata Denmark X399 Laccaria sp.2(2) 99 UDB001466 Laccaria laccata Denmark X427 Lactarius aff. hysginus 99 MH985012.1 Lactarius sp. South Korea X175 Lactarius aff. nitidus 99 GQ415318.1 Lactarius sp. USA R138 Lactarius aff. uvidus(1) 99 KR090925.1 Lactarius montanus USA X1002U Lactarius aff. uvidus(2) 99 KR090925.1 Lactarius montanus USA R256 Lactarius badiosanguineus 99 KX394281.1 Lactarius badiosanguineus USA R685U Lactarius cf. affinis(1) 99 KJ705208.1 Lactarius affinis Canada X415 Lactarius cf. affinis(2) 99 KJ705208.1 Lactarius affinis Canada L37 Lactarius cf. camphoratus 99 UDB000885 Lactarius camphoratus Sweden P1010 Lactarius cf. picinus(1) 99 JQ446132.1 Lactarius picinus Norway R523 Lactarius cf. picinus(2) 98 JQ446132.1 Lactarius picinus Norway X960 Lactarius cf. picinus(3) 99 JQ446132.1 Lactarius picinus Norway X103 Lactarius cf. rufus 100 KT165277.1 Lactarius rufus Russia X396 Lactarius cf. tabidus(1) 99 MK131494.1 Lactarius tabidus Canada X50 Lactarius cf. tabidus(2) 99 MK131494.1 Lactarius tabidus Canada V467_ITS1F Lentaria byssiseda(1) 99 UDB018199 Lentaria byssiseda Estonia X12_ITS1F Lentaria byssiseda(2) 99 UDB018199 Lentaria byssiseda Estonia M394 Lentinellus cf. cystidiosus(1) 99 NR_119502.1 Lentinellus cystidiosus USA X470 Lentinellus cf. cystidiosus(2) 99 NR_119502.1 Lentinellus cystidiosus USA F2 Lentinellus micheneri 99 AY513160.1 Lentinellus micheneri Greenland X153 Lentinus brumalis 100 KU761244.1 Polyporus brumalis Canada R79 Lepiota castanea 99 AY176463.1 Lepiota castanea Netherlands R466 Lepiota cf. cristata 99 AF391051.1 Lepiota cristata USA R242 Lepiota cf. fuscosquamea 99 UDB015543 Lepiota cortinarius Estonia R904_ITS1F Lepiota cf. magnispora 99 UDB015845 Lepiota magnispora Estonia R738 Lepiota cf. pseudolilacea 99 U85330.1 Lepiota felina USA R26 Lepiota clypeolaria(1) 99 AF390999.1 Lepiota clypeolaria USA R453 Lepiota clypeolaria(2) 99 AF390999.1 Lepiota clypeolaria USA X480 Lepiota sp.1 96 MH979433.1 Lepiota castanea Canada X740 Lepista cf. nebularis 100 UDB023648 Lepista nebularis Estonia X939_ITS1F Leucocybe cf. candicans(1) 99 JN021000.1 Clitocybe dealbata Canada

108

Collection # Morphospecies Max Accession # Top GenBank or UNITE hit Accession # identity country %

B751 Leucocybe cf. candicans(2) 99 JN021000.1 Clitocybe dealbata Canada X1234 Leucocybe cf. candicans(3) 99 KJ681027.1 Leucocybe candicans Spain X1208 Lyophyllum cf. mycenoides 99 KP192561.1 Lyophyllum cf. mycenoides France P1021 Marasmiaceae sp.1 89 MF100962.1 Trogia aff. furcata Sao Tome X1006 Marasmius cf. cohaerens 99 KF774175.1 Marasmius cohaerens Canada C314 Marasmius cf. wettsteinii(1) 99 UDB018814 Marasmius wettsteinii Canada P474 Marasmius cf. wettsteinii(2) 99 UDB018814 Marasmius wettsteinii Canada B476 Marasmius epiphyllus 99 UDB015277 Marasmius epiphyllus Estonia C322A Mycena cf. acicula(1) 97 JF908384.1 Mycena acicula Italy M353 Mycena cf. acicula(2) 99 JF908384.1 Mycena acicula Italy X21 Mycena cf. amicta(1) 99 MH857183.1 Mycena amicta France B782B Mycena cf. amicta(2) 99 MH857183.1 Mycena amicta France B113 Mycena cf. amicta(3) 99 MH857183.1 Mycena amicta France X375 Mycena cf. borealis 93 UDB015432 Mycena polygramma Estonia B102 Mycena cf. capillaripes(1) 97 UDB019514 Mycena capillaripes Estonia P1003 Mycena cf. capillaripes(2) 97 UDB019514 Mycena capillaripes Estonia X113 Mycena cf. citrinomarginata(1) 99 KT149164.1 Mycena sp. China M222 Mycena cf. citrinomarginata(2) 100 GU234150.1 Mycena citrinomarginata Svalbard V390 Mycena cf. citrinomarginata(3) 99 GU234150.1 Mycena citrinomarginata Svalbard J103 Mycena cf. clavicularis 99 JF908467.1 Mycena clavicularis Spain B412 Mycena cf. epipterygia var. lignicola 100 MH396633.1 Mycena epipterygia China B191 Mycena cf. filopes 100 HQ604769.1 Mycena filopes China R801 Mycena cf. floridula(1) 99 KJ705189.1 Mycena adonis var. adonis Canada V225 Mycena cf. floridula(2) 99 KJ705189.1 Mycena adonis var. adonis Canada P969 Mycena cf. floridula(3) 99 KJ705189.1 Mycena adonis var. adonis Canada R760U Mycena cf. floridula(4) 98 KJ705189.1 Mycena adonis var. adonis Canada P957 Mycena cf. floridula(5) 98 KJ705189.1 Mycena adonis var. adonis Canada C501 Mycena cf. floridula(6) 99 KJ705189.1 Mycena adonis var. adonis Canada B460 Mycena cf. flos-nivium 98 MH857186.1 Mycena flos-nivium France M3 Mycena cf. leptocephala(1) 99 UDB019723 Mycena sp. Estonia M43 Mycena cf. leptocephala(2) 99 UDB019723 Mycena sp. Estonia X1205 Mycena cf. metata 99 MH396636.1 Mycena metata China B590 Mycena cf. praelonga 98 JF908049.1 Hydropus trichoderma Italy B738 Mycena cf. pura(1) 99 KF007948.1 Mycena pura USA C246 Mycena cf. pura(2) 99 JN021065.2 Mycena cf. purpureofusca Canada B437_ITS4 Mycena cf. robusta(1) 99 KJ705179.1 Mycena robusta Canada L205 Mycena cf. robusta(2) 99 KJ705179.1 Mycena robusta Canada X425 Mycena cf. tenax(1) 99 EU669224.1 Mycena tenax USA B359 Mycena cf. tenax(2) 99 EU846251.1 Mycena tenax USA M319 Mycena purpureofusca 99 MG654742.1 Mycena purpureofusca China M185 Mycena silvae-nigrae(1) 99 JF908452.1 Mycena silvae-nigrae Italy M4 Mycena silvae-nigrae(2) 99 KF359604.1 Mycena silvae-nigrae USA J44 Mycena silvae-nigrae(3) 98 KF359604.1 Mycena silvae-nigrae USA M65 Mycena silvae-nigrae(4) 98 KF359604.1 Mycena silvae-nigrae USA P611 Mycena sp.1 96 JF908391.1 Mycena renati Italy P340U Mycena subcaerulea(1) 89 MH857183.1 Mycena amicta France P332 Mycena subcaerulea(2) 90 DQ490645.1 Mycena amicta USA M899 Mycena urania(1) 94 UDB015412 Mycena metata Estonia M184 Mycena urania(2) 94 UDB015412 Mycena metata Estonia M912 Mycena urania(3) 94 UDB015412 Mycena metata Estonia B408U Mycena vulgaris 99 UDB017891 Mycena vulgaris Estonia M520 Mycenella lasiosperma 99 UDB017867 Mycenella lasiosperma Estonia

109

Collection # Morphospecies Max Accession # Top GenBank or UNITE hit Accession # identity country %

R313 Naucoria aff. silvaenovae 99 JN943941.1 Alnicola silvaenovae Estonia V80 Neofavolus cf. alveolaris 100 MH979293.1 Neofavolus sp. USA P309A Neofavolus sp.1(1) 99 KP283502.1 Neofavolus sp. Canada B89 Neofavolus sp.1(2) 99 KP283505.1 Neofavolus sp. USA R856 Onnia tomentosa 99 UDB01184 Onnia tomentosa Estonia C426 Peniophora cf. erikssonii 100 UDB031589 Peniophora erikssonii Finland W69 Phaeoclavulina aff. flaccida(1) 96 JQ408227.1 Phaeoclavulina argentea USA B627 Phaeoclavulina aff. flaccida(2) 96 JQ408227.1 Phaeoclavulina argentea USA V429 Phaeoclavulina cf. abietina 97 AJ408383.1 Ramaria abietina Spain P489 Phaeomarasmius proximans 99 DQ404381.1 Phaeomarasmius proximans USA M9 Phanerochaete sp.1 99 KP134995.1 Phanerochaete sp. USA M865 Phloeomana cf. speirea 98 UDB017946 Mycena speirea Estonia X123_ITS1F Pholiota cf. spumosa(1) 99 UDB015486 Pholiota spumosa Estonia R98 Pholiota cf. spumosa(2) 100 UDB015486 Pholiota spumosa Estonia W11 Pholiota cf. spumosa(3) 99 UDB015486 Pholiota spumosa Estonia W16 Pholiota cf. spumosa(4) 99 UDB015486 Pholiota spumosa Estonia C590 Pholiota cf. subsulphurea 100 FJ596817.1 Pholiota sp. USA M387 Picipes sp.1 97 KY038480.1 Polyporus sp. South Korea M279 Pleurotus cf. abieticola 99 AF345656.1 Pleurotus abieticola Russia J213 Pleurotus populinus 100 KP026249.1 Pleurotus populinus USA X453 Plicatura nivea 99 UDB016382 Plicaturopsis crispa Estonia W52 Pluteus aff. hispidulus 94 FJ774083.1 Pluteus exiguus Russia M149 Pluteus aff. tomentosulus 99 KR022009.1 Pluteus sp. USA R5 Pluteus brunneidiscus 100 HM562217.1 Pluteus brunneidiscus USA X154 Pluteus cf. rangifer 100 KJ009652.1 Pluteus rangifer Russia B66 Postia cf. guttulata 99 UDB031613 Postia guttulata Finland X349 Postia cf. subcaesia 99 JQ673071.1 Posta subcaesia USA X402 Ramaria cf. apiculata(1) 99 FJ627035.1 Ramaria abietina Canada B161 Ramaria cf. apiculata(2) 99 KT307859.1 Ramaria abietina Mexico B128 Ramaria cf. apiculata(3) 99 KT307859.1 Ramaria abietina Mexico B577 Ramaria cf. apiculata(4) 99 JQ408241.1 Ramaria sp. USA X383 Ramaria cf. apiculata(5) 99 KT307859.1 Ramaria abietina Mexico L21 Ramaria cf. rubella 96 KC346860.1 Ramaria rubella USA X423 Ramaria sp.1(1) 99 DQ365605.1 Ramaria sp. USA X1019U_ITS1F Ramaria sp.1(2) 99 UDB032771 Ramaria eumorpha Canada M1 Rhizocybe aff. vermicularis 97 KY030726.1 Rhizocybe alba China L285U Rhodocollybia maculata 100 KY777395.1 Rhodocollybia maculata USA J31 Rhodofomes cajanderi 99 KU863095.1 Fomitopsis cajanderi N/A L112 Rhodophana cf. nitellina(1) 88 UDB015654 Rhodocybe nitellina Estonia R623 Rhodophana cf. nitellina(2) 88 UDB015654 Rhodocybe nitellina Estonia X835 Rhodophana cf. nitellina(3) 88 UDB015654 Rhodocybe nitellina Estonia B369 Rhodophana cf. nitellina(4) 96 UDB015654 Rhodocybe nitellina Estonia F28 Rhodophana cf. nitellina(5) 88 UDB015654 Rhodocybe nitellina Estonia R713 Rhodophana cf. nitellina(6) 96 UDB015654 Rhodocybe nitellina Estonia R228 Rhodophana cf. nitellina(7) 89 UDB015654 Rhodocybe nitellina Estonia M181_ITS1F Rickenella cf. fibula 90 MG982555.1 Rickenella fibula USA B523 Ripartites cf. metrodii 99 JF908749.1 Ripartites tricholoma Italy P1001 Roridomyces roridus(1) 99 FJ596760.1 Mycena rorida USA V392 Roridomyces roridus(2) 99 FJ596760.1 Mycena rorida USA X279 Russula acetolens(1) 99 HQ604848.1 Russula lutea Canada R530 Russula acetolens(2) 99 HQ604848.1 Russula lutea Canada X74_ITS1F Russula aff. firmula 99 JF834342.1 Russula aff. firmula USA R525 Russula aff. renidens 96 UDB015975 Russula renidens Estonia

110

Collection # Morphospecies Max Accession # Top GenBank or UNITE hit Accession # identity country %

X124 Russula aff. sphagnophila 99 JX425400.1 Russula sp. China X827 Russula cf. fragilis 99 HQ650737.1 Russula fragilis Canada R71 Russula cf. fragrantissima(1) 99 KF245487.1 Russula foetens USA R76 Russula cf. fragrantissima(2) 100 KF245487.1 Russula foetens USA R120 Russula cf. globispora(1) 99 UDB016122 Russula globispora Estonia F99U Russula cf. globispora(2) 99 UDB016122 Russula globispora Estonia R514U Russula cf. hydrophila(1) 98 KX812853.1 Russula montana Canada W51 Russula cf. hydrophila(2) 99 UDB024024 Russula paludosa Canada R179 Russula cf. hydrophila(3) 99 KX579802.1 Russula montana Canada F147U Russula cf. integriformis 99 UDB031529 Russula integriformis Canada X56 Russula cf. nauseosa 99 KT933985.1 Russula nauseosa Germany W70 Russula cf. olivina 99 UDB016260 Russula olivina Finland X421 Russula cf. turci(1) 100 JQ711969.1 Russula turci Canada X441 Russula cf. turci(2) 100 JQ711969.1 Russula turci Canada R137 Russula cf. versicolor 99 KP177204.1 Russula odorata China F19 Russula sp.1 99 UDB024064 Russula puellaris Canada B635 Simocybe cf. centunculus 92 KT715796.1 Simocybe reducta Italy B552 Simocybe cf. reducta 99 KT715796.1 Simocybe reducta Italy M546 Simocybe cf. sumptuosa 99 MF153085.1 Simocybe serrulata USA L14 Sistotrema cf. confluens 99 UDB017974 Sistotrema confluens Estonia X1287 Steccherinum ochraceum 99 KJ140599.1 Steccherinum ochraceum USA M404 Stereum cf. sanguinolentum(1) 99 MH856746.1 Stereum sanguinolentum Canada W85 Stereum cf. sanguinolentum(2) 99 MH856746.1 Stereum sanguinolentum Canada B268 Strobilurus albipilatus(1) 100 GQ892818.1 Strobilurus albipilatus USA R224 Strobilurus albipilatus(2) 99 GQ892818.1 Strobilurus albipilatus USA B708 Tephrocybe cf. striipilea(1) 99 KP192644.1 Tephrocybe striaepilea France B712_ITS1F Tephrocybe cf. striipilea(2) 99 KP192647.1 Tephrocybe striaepilea France B492 Tephrocybe cf. striipilea(3) 99 KP192647.1 Tephrocybe striaepilea France X859_ITS1F Tephrocybe cf. striipilea(4) 97 KP192647.1 Tephrocybe striaepilea France R448 Tephrocybe cf. striipilea(5) 94 KP192647.1 Tephrocybe striaepilea France X1096 Tephrocybe cf. striipilea(6) 98 KP192647.1 Tephrocybe striaepilea France R146 Tephrocybe cf. striipilea(7) 99 KP192647.1 Tephrocybe striaepilea France R226 Tephrocybe sp.1 96 KP192559.1 Tephrocybe cf. ochraceobrunnea France W92 Thelephora cf. anthocephala(1) 99 MF926570.1 Thelephora cf. anthocephala Russia X1032 Thelephora cf. anthocephala(2) 99 MF926570.1 Thelephora cf. anthocephala Russia F49 Thelephora sp.1 100 MH038094.1 Thelephora sp. USA M47 Trichaptum cf. abietinum 99 MF381025.1 Trichaptum abietinum USA X304 Trichaptum cf. biforme 100 MH862562.1 Trichaptum biforme USA M44U Trichaptum cf. fuscoviolaceum(1) 99 MH211725.1 Trichaptum sp. USA F119L Trichaptum cf. fuscoviolaceum(2) 99 MF319120.1 Trichaptum fuscoviolaceum Finland X1229 Tricholoma cf. fulvum 99 LT000171.1 Tricholoma fulvum Sweden F33_ITS1F Tricholoma cf. myomyces(1) 100 JN389291.1 Tricholoma myomyces USA F23 Tricholoma cf. myomyces(2) 100 JN389291.1 Tricholoma myomyces USA X1159_ITS1F Tricholoma inamoenum 100 JN021105.1 Tricholoma inamoenum Canada F135U Tricholomopsis cf. flammula(1) 99 FN554893.1 Tricholomopsis flammula Czech Republic M966U Tricholomopsis cf. flammula(2) 98 FN554896.1 Tricholomopsis flammula Czech Republic M395_ITS1F Tubaria cf. confragosa 98 DQ267126.1 Tubaria confragosa USA L264 Tubaria cf. furfuracea(1) 97 MH173775.1 Tubaria romagnesiana USA M67 Tubaria cf. furfuracea(2) 99 MH173775.1 Tubaria romagnesiana USA P497 Tubaria cf. furfuracea(3) 99 MH173775.1 Tubaria romagnesiana USA X920 Tubaria cf. furfuracea(4) 99 MH173775.1 Tubaria romagnesiana USA M191 Typhrasa aff. gossypina(1) 91 KC992946.1 Psathyrella gossypina Germany M205 Typhrasa aff. gossypina(2) 91 KC992946.1 Psathyrella gossypina Germany

111

Collection # Morphospecies Max Accession # Top GenBank or UNITE hit Accession # identity country %

M48 Xeromphalina cf. campanella(1) 99 KM024537.1 Xeromphalina campanella USA X451 Xeromphalina cf. campanella(2) 99 KM024520.1 Xeromphalina campanella USA J16 Xeromphalina cf. enigmatica 99 KM024591.1 Xeromphalina sp. USA

112

Appendix II. Morphospecies of fruiting bodies sampled from unlogged and logged plots in mixedwood forests near Kapuskasing, Ontario, together with the total number of occurrences recorded and ecological guild assignments. Ones indicate assignment to the respective guild.

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Ascomycota Akanthomyces pistillariiformis 1 0 0 0 1 Ascomycota Bisporella citrina 6 0 1 0 0 Ascomycota Byssonectria terrestris 1 0 0 1 0 Ascomycota Chlorociboria aeruginascens 8 0 1 0 0 Ascomycota Chlorosplenium cf. chlora 1 0 1 0 0 Ascomycota Ciboria rufofusca 1 0 0 1 0 Ascomycota Cordyceps militaris 5 0 0 0 1 Ascomycota Cudonia circinans 11 0 1 1 0 Ascomycota Daldinia childiae 5 0 1 0 0 Ascomycota Discina ancilis 4 0 1 1 0 Ascomycota Geopyxis cf. carbonaria 1 0 0 1 0 Ascomycota Gibellula leiopus 4 0 0 0 1 Ascomycota Helvella atra 1 1 0 1 0 Ascomycota Helvella costifera 1 1 0 0 0 Ascomycota Helvella crispa 1 1 0 0 0 Ascomycota Helvella lacunosa 2 1 0 0 0 Ascomycota Helvella macropus 6 1 0 0 0 Ascomycota Humaria hemisphaerica 18 1 0 0 0 Ascomycota Hymenoscyphus cf. calyculus 2 0 1 0 0 Ascomycota Hymenoscyphus cf. monticola 2 0 1 0 0 Ascomycota Hymenoscyphus sp.1 12 0 1 0 0 Ascomycota Hymenoscyphus vernus 2 0 1 0 0 Ascomycota Hypoxylon cf. perforatum 7 0 1 0 0 Ascomycota Hypoxylon fragiforme 7 0 1 0 0 Ascomycota Hypoxylon sp.1 2 0 1 0 0 Ascomycota Hypoxylon sp.2 1 0 1 0 0 Ascomycota Jackrogersella cf. multiformis 38 0 1 0 0 Ascomycota Lachnellula cf. calyciformis 14 0 1 0 0 Ascomycota Leotia lubrica 11 0 1 1 0 Ascomycota Morchella cf. angusticeps 5 1 0 0 0 Ascomycota Neocudoniella albiceps 4 0 1 0 0 Ascomycota Ophiocordyceps gracilis 4 0 0 0 1 Ascomycota Ophiocordyceps variabilis 1 0 0 0 1 Ascomycota Otidea cf. bufonia 2 0 0 1 0 Ascomycota Otidea cf. mirabilis 2 0 0 1 0 Ascomycota Otidea cf. tuomikoskii 1 0 0 1 0 Ascomycota Pachyella cf. clypeata 3 0 1 0 0 Ascomycota Peziza brunneoatra 1 0 0 1 0 Ascomycota Peziza cf. badia 1 0 0 1 0

113

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Ascomycota Peziza cf. michelii 4 0 0 1 0 Ascomycota Peziza cf. varia 10 0 1 0 0 Ascomycota Peziza phyllogena 1 0 0 1 0 Ascomycota Peziza succosa 4 0 0 1 0 Ascomycota Pseudoplectania nigrella 8 0 0 1 0 Ascomycota Pseudorhiza sphaerospora 3 0 1 0 0 Ascomycota Rutstroemia sp.1 1 0 1 0 0 Ascomycota Rutstroemiaceae sp.1 2 0 1 1 0 Ascomycota Sarcosoma globosum 2 0 0 1 0 Ascomycota Scutellinia scutellata 14 0 1 0 0 Ascomycota Spathularia flavida 4 0 0 1 0 Ascomycota Spathulariopsis velutipes 4 0 1 1 0 Ascomycota Tarzetta cf. cupularis 3 1 0 0 0 Ascomycota Tolypocladium ophioglossoides 2 0 0 0 1 Ascomycota Trichoglossum alutaceum 2 0 0 1 0 Ascomycota Trichoglossum cf. farlowii 1 0 1 1 0 Ascomycota Xylaria hypoxylon 5 0 1 0 0 Basidiomycota Agaricus cf. abruptibulbus 2 0 0 1 0 Basidiomycota Agaricus cf. friesianus 1 0 0 1 0 Basidiomycota Agaricus cf. langei 1 0 0 1 0 Basidiomycota Agaricus cf. silvicola 3 0 0 1 0 Basidiomycota Agrocybe cf. pediades 1 0 0 1 0 Basidiomycota Agrocybe cf. praecox 1 0 1 1 0 Basidiomycota Agrocybe firma 2 0 1 1 0 Basidiomycota Aleurodiscus amorphus 3 0 1 0 0 Basidiomycota Amanita albiceps 1 1 0 0 0 Basidiomycota Amanita cf. fulva 1 1 0 0 0 Basidiomycota Amanita cf. rhacopus 1 1 0 0 0 Basidiomycota Amanita cf. sinicoflava 9 1 0 0 0 Basidiomycota Ampulloclitocybe cf. clavipes 2 0 0 1 0 Basidiomycota Aphroditeola olida 1 0 0 1 0 Basidiomycota Armillaria aff. sinapina 19 0 1 0 1 Basidiomycota Armillaria cf. gallica 12 0 1 0 0 Basidiomycota Armillaria cf. ostoyae 19 0 1 0 1 Basidiomycota Arrhenia aff. epichysium 9 0 1 0 0 Basidiomycota Artomyces pyxidatus 5 0 1 0 0 Basidiomycota Atractosporocybe cf. inornata 2 0 0 1 0 Basidiomycota Auricularia americana 8 0 1 0 0 Basidiomycota Auriscalpium vulgare 3 0 0 1 0 Basidiomycota Baeospora myriadophylla 20 0 1 0 0 Basidiomycota Bjerkandera adusta 12 0 1 0 0 Basidiomycota Bolbitiaceae sp.1 1 0 1 0 0 Basidiomycota Calocera cornea 2 0 1 0 0

114

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Cantharellus cf. cibarius 5 1 0 0 0 Basidiomycota Cerioporus cf. varius 57 0 1 0 0 Basidiomycota Cerioporus mollis 2 0 1 0 0 Basidiomycota Cerrena unicolor 3 0 1 0 0 Basidiomycota Chalciporus piperatus 1 0 0 0 1 Basidiomycota Cheimonophyllum candidissimum 47 0 1 0 0 Basidiomycota Chromosera cyanophylla 2 0 1 0 0 Basidiomycota Chroogomphus rutilus 1 1 0 0 0 Basidiomycota Clavariadelphus cf. sachalinensis 3 0 0 1 0 Basidiomycota Clavulina cf. coralloides 2 1 0 0 0 Basidiomycota Clavulina cf. rugosa 4 1 0 0 0 Basidiomycota Clavulinopsis cf. corniculata 2 0 1 1 0 Basidiomycota Clavulinopsis laeticolor 6 0 1 1 0 Basidiomycota Clitocybe cf. coniferophila 2 0 0 1 0 Basidiomycota Clitocybe cf. diatreta 1 0 0 1 0 Basidiomycota Clitocybe cf. gibba 10 0 0 1 0 Basidiomycota Clitocybe cf. subclavipes 1 0 0 1 0 Basidiomycota Clitocybe cf. truncicola 3 0 1 0 0 Basidiomycota Clitocybe cf. vibecina 3 0 0 1 0 Basidiomycota Clitocybula aff. familia 1 0 1 0 0 Basidiomycota Clitocybula odora 5 0 0 1 0 Basidiomycota Clitopilus prunulus 3 0 0 1 0 Basidiomycota Collybia cookei 10 0 1 1 0 Basidiomycota Collybia tuberosa 25 0 0 1 0 Basidiomycota Coltricia perennis 1 1 0 1 0 Basidiomycota Conocybe cf. apala 1 0 0 1 0 Basidiomycota Conocybe cf. siennophylla 2 0 1 1 0 Basidiomycota Coprinellus cf. radians 3 0 1 0 0 Basidiomycota Coprinellus sp.1 2 0 1 0 0 Basidiomycota Corticium roseum 1 0 1 0 0 Basidiomycota Cortinarius aff. barbarorum 1 1 0 0 0 Basidiomycota Cortinarius aff. boreidionysae 1 1 0 0 0 Basidiomycota Cortinarius aff. caesioarmeniacus 1 1 0 0 0 Basidiomycota Cortinarius aff. delibutus 8 1 0 0 0 Basidiomycota Cortinarius aff. dolabratus 5 1 0 0 0 Basidiomycota Cortinarius aff. glaucopus 1 1 0 0 0 Basidiomycota Cortinarius aff. helvelloides 2 1 0 0 0 Basidiomycota Cortinarius aff. laetissimus 1 1 0 0 0 Basidiomycota Cortinarius aff. privignipallens 1 1 0 0 0 Basidiomycota Cortinarius aff. xanthocephalus 2 1 0 0 0 Basidiomycota Cortinarius albocyaneus 1 1 0 0 0 Basidiomycota Cortinarius alboviolaceus 3 1 0 0 0 Basidiomycota Cortinarius biformis 6 1 0 0 0

115

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Cortinarius bolaris 1 1 0 0 0 Basidiomycota Cortinarius caperatus 1 1 0 0 0 Basidiomycota Cortinarius centrirufus 1 1 0 0 0 Basidiomycota Cortinarius cf. anomalochrascens 1 1 0 0 0 Basidiomycota Cortinarius cf. argenteolilacinus 1 1 0 0 0 Basidiomycota Cortinarius cf. aureovelatus 1 1 0 0 0 Basidiomycota Cortinarius cf. balaustinus 10 1 0 0 0 Basidiomycota Cortinarius cf. betuletorum 2 1 0 0 0 Basidiomycota Cortinarius cf. brunneus 10 1 0 0 0 Basidiomycota Cortinarius cf. caninus 1 1 0 0 0 Basidiomycota Cortinarius cf. casimiri 3 1 0 0 0 Basidiomycota Cortinarius cf. comptulus 5 1 0 0 0 Basidiomycota Cortinarius cf. decipiens 25 1 0 0 0 Basidiomycota Cortinarius cf. depressus 1 1 0 0 0 Basidiomycota Cortinarius cf. diasemospermus 3 1 0 0 0 Basidiomycota Cortinarius cf. flexipes 13 1 0 0 0 Basidiomycota Cortinarius cf. fulvescens 2 1 0 0 0 Basidiomycota Cortinarius cf. glandicolor 4 1 0 0 0 Basidiomycota Cortinarius cf. impennoides 2 1 0 0 0 Basidiomycota Cortinarius cf. infractus 3 1 0 0 0 Basidiomycota Cortinarius cf. lucorum 3 1 0 0 0 Basidiomycota Cortinarius cf. malachius 4 1 0 0 0 Basidiomycota Cortinarius cf. malicorius 12 1 0 0 0 Basidiomycota Cortinarius cf. obtusus 5 1 0 0 0 Basidiomycota Cortinarius cf. paragaudis 1 1 0 0 0 Basidiomycota Cortinarius cf. polaris 1 1 0 0 0 Basidiomycota Cortinarius cf. renidens 3 1 0 0 0 Basidiomycota Cortinarius cf. rusticus 4 1 0 0 0 Basidiomycota Cortinarius cf. sanguineus 9 1 0 0 0 Basidiomycota Cortinarius cf. saniosus 1 1 0 0 0 Basidiomycota Cortinarius cf. scandens 2 1 0 0 0 Basidiomycota Cortinarius cf. subcroceofolius 1 1 0 0 0 Basidiomycota Cortinarius cf. tabularis 2 1 0 0 0 Basidiomycota Cortinarius cf. trivialis 8 1 0 0 0 Basidiomycota Cortinarius croceus 2 1 0 0 0 Basidiomycota Cortinarius cyanites 1 1 0 0 0 Basidiomycota Cortinarius flabellus 1 1 0 0 0 Basidiomycota Cortinarius rubellus 3 1 0 0 0 Basidiomycota Cortinarius sp.1 1 1 0 0 0 Basidiomycota Cortinarius sphagnophilus 5 1 0 0 0 Basidiomycota Cortinarius spilomeus 7 1 0 0 0 Basidiomycota Cortinarius subbalaustinus 7 1 0 0 0 Basidiomycota Cortinarius traganus 1 1 0 0 0

116

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Cortinarius vibratilis 15 1 0 0 0 Basidiomycota Cortinarius violaceus 11 1 0 0 0 Basidiomycota Craterellus fallax 1 1 0 0 0 Basidiomycota Craterellus tubaeformis 1 1 0 0 0 Basidiomycota Crepidotus aff. cesatii 29 0 1 0 0 Basidiomycota Crepidotus aff. luteolus 34 0 1 0 0 Basidiomycota Crepidotus calolepis 36 0 1 0 0 Basidiomycota Crepidotus cf. applanatus 1 0 1 0 0 Basidiomycota Crepidotus cf. conchatus 1 0 1 0 0 Basidiomycota Crepidotus cf. occidentalis 2 0 1 0 0 Basidiomycota Crepidotus cf. versutus 8 0 1 0 0 Basidiomycota Crepidotus cf. vulgaris 5 0 1 0 0 Basidiomycota Crepidotus malachius 11 0 1 0 0 Basidiomycota Crepidotus nyssicola 1 0 1 0 0 Basidiomycota Crinipellis campanella 2 0 1 0 0 Basidiomycota Crinipellis piceae 22 0 0 1 0 Basidiomycota Cryptoporus volvatus 2 0 1 0 0 Basidiomycota Cyclocybe cf. erebia 3 0 0 1 0 Basidiomycota Cyptotrama cf. chrysopepla 1 0 1 0 0 Basidiomycota Cystoderma amianthinum 1 0 0 1 0 Basidiomycota Dacrymyces chrysospermus 10 0 1 0 0 Basidiomycota Dacrymyces stillatus 1 0 1 0 0 Basidiomycota Dacryopinax spathularia 6 0 1 0 0 Basidiomycota Daedaleopsis confragosa 9 0 1 0 0 Basidiomycota Datroniella scutellata 2 0 1 0 0 Basidiomycota Dermoloma sp.1 1 0 0 1 0 Basidiomycota Ductifera pululahuana 2 0 1 0 0 Basidiomycota Entoloma aff. llimonae 3 0 0 1 0 Basidiomycota Entoloma aff. minutum 2 0 1 0 0 Basidiomycota Entoloma aff. serrulatum 2 0 1 1 0 Basidiomycota Entoloma caeruleogriseum 1 0 1 0 0 Basidiomycota Entoloma cf. abortivum 1 0 1 1 1 Basidiomycota Entoloma cf. anatinum 2 0 0 1 0 Basidiomycota Entoloma cf. asprellum 9 0 0 1 0 Basidiomycota Entoloma cf. bicolor 1 0 0 1 0 Basidiomycota Entoloma cf. dysthales 7 0 0 1 0 Basidiomycota Entoloma cf. elodes 1 0 0 1 0 Basidiomycota Entoloma cf. exile 1 0 1 1 0 Basidiomycota Entoloma cf. formosum 3 0 1 1 0 Basidiomycota Entoloma cf. fragrans 1 0 0 1 0 Basidiomycota Entoloma cf. griseum 31 0 0 1 0 Basidiomycota Entoloma cf. olivaceotinctum 7 0 0 1 0 Basidiomycota Entoloma cf. papillatum 2 0 0 1 0

117

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Entoloma cf. rhodopolium 23 1 0 0 0 Basidiomycota Entoloma cf. rubrobasis 6 1 0 0 0 Basidiomycota Entoloma melenosmum 2 1 0 0 0 Basidiomycota Entoloma sericellum 2 0 1 1 0 Basidiomycota Entoloma sp.1 4 0 1 1 0 Basidiomycota Entoloma sp.2 2 0 1 1 0 Basidiomycota Exidia recisa 7 0 1 0 0 Basidiomycota Exidia repanda 11 0 1 0 0 Basidiomycota Flammulaster cf. granulosus 7 0 0 1 0 Basidiomycota Flammulaster cf. rhombosporus 16 0 0 1 0 Basidiomycota Fomes fomentarius 27 0 1 0 1 Basidiomycota Fomitiporia cf. punctata 6 0 1 0 0 Basidiomycota Fomitopsis betulina 9 0 1 0 1 Basidiomycota Fomitopsis pinicola 33 0 1 0 1 Basidiomycota Galerina aff. stylifera 8 0 1 0 0 Basidiomycota Galerina cf. hypnorum 2 0 1 1 0 Basidiomycota Galerina cf. mniophila 1 0 0 1 0 Basidiomycota Galerina cf. triscopa 13 0 1 0 0 Basidiomycota Galerina cf. vittiformis 11 0 1 0 0 Basidiomycota Ganoderma applanatum 1 0 1 0 1 Basidiomycota Ganoderma tsugae 1 0 1 0 1 Basidiomycota Geastrum cf. saccatum 5 0 0 1 0 Basidiomycota Geastrum quadrifidum 3 0 0 1 0 Basidiomycota Gliophorus laetus 3 0 0 1 0 Basidiomycota Gliophorus psittacinus 1 0 0 1 0 Basidiomycota Gloeophyllum cf. sepiarium 2 0 1 0 0 Basidiomycota Gomphidius borealis 1 1 0 0 0 Basidiomycota Guepinia helvelloides 2 0 1 0 0 Basidiomycota Guepiniopsis alpina 1 0 1 0 0 Basidiomycota Gymnopilus cf. bellulus 5 0 1 0 0 Basidiomycota Gymnopus aff. westii 15 0 0 1 0 Basidiomycota Gymnopus cf. alkalivirens 8 0 0 1 0 Basidiomycota Gymnopus cf. androsaceus 14 0 0 1 0 Basidiomycota Gymnopus cf. contrarius 48 0 0 1 0 Basidiomycota Gymnopus cf. dryophilus 47 0 1 1 0 Basidiomycota Gymnopus cf. impudicus 4 0 0 1 0 Basidiomycota Gymnopus cf. perforans 51 0 0 1 0 Basidiomycota Gymnopus cf. putillus 1 0 0 1 0 Basidiomycota Gymnopus cf. subsulphureus 15 0 1 1 0 Basidiomycota Gymnopus confluens 52 0 0 1 0 Basidiomycota Gymnopus foetidus 1 0 1 0 0 Basidiomycota Hapalopilus rutilans 2 0 1 0 0 Basidiomycota Hebeloma cf. excedens 2 1 0 0 0

118

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Hebeloma cf. fastibile 2 1 0 0 0 Basidiomycota Hebeloma cf. fusisporum 3 1 0 0 0 Basidiomycota Hebeloma cf. incarnatulum 2 1 0 0 0 Basidiomycota Hebeloma cf. mesophaeum 1 1 0 0 0 Basidiomycota Hebeloma sp.1 1 1 0 0 0 Basidiomycota Hebeloma sp.2 3 1 0 0 0 Basidiomycota Hemimycena aff. gracilis 53 0 0 1 0 Basidiomycota Hemimycena cf. pseudolactea 28 0 0 1 0 Basidiomycota Hemimycena mairei 1 0 0 1 0 Basidiomycota Hemimycena mauretanica 2 0 0 1 0 Basidiomycota Hericium americanum 1 0 1 0 0 Basidiomycota Hericium corraloides 2 0 1 0 0 Basidiomycota Heteroradulum cf. kmetii 15 0 1 0 0 Basidiomycota Hohenbuehelia sp.1 1 0 1 0 0 Basidiomycota Homophron cf. camptopodum 1 0 1 0 0 Basidiomycota Homophron cf. spadiceum 4 0 0 1 0 Basidiomycota Hydnopolyporus fimbriatus 2 0 1 0 0 Basidiomycota Hydnum cf. albertense 7 1 0 0 0 Basidiomycota Hydnum cf. melitosarx 5 1 0 0 0 Basidiomycota Hydropus marginellus 5 0 1 0 0 Basidiomycota Hygrocybe cf. cantharellus 31 0 0 0 1 Basidiomycota Hygrocybe cf. parvula 3 0 0 0 1 Basidiomycota Hygrocybe cf. vitellina 1 0 0 0 1 Basidiomycota Hygrocybe conica 1 0 0 0 1 Basidiomycota Hygrophoropsis cf. macrospora 1 0 0 1 0 Basidiomycota Hymenochaetopsis tabacina 61 0 1 0 0 Basidiomycota Hypholoma capnoides 2 0 1 0 0 Basidiomycota Hypholoma sp.1 1 0 1 0 0 Basidiomycota Infundibulicybe sp.1 5 0 0 1 0 Basidiomycota Inocybe aff. proximella 5 1 0 0 0 Basidiomycota Inocybe aff. rimosa 2 1 0 0 0 Basidiomycota Inocybe cf. calamistrata 3 1 0 0 0 Basidiomycota Inocybe cf. flocculosa 16 1 0 0 0 Basidiomycota Inocybe cf. griseolilacina 8 1 0 0 0 Basidiomycota Inocybe cf. jacobi 23 1 0 0 0 Basidiomycota Inocybe cf. lacera 7 1 0 0 0 Basidiomycota Inocybe cf. lanuginosa 12 1 0 0 0 Basidiomycota Inocybe cf. leptocystis 1 1 0 0 0 Basidiomycota Inocybe cf. maculata 10 1 0 0 0 Basidiomycota Inocybe cf. mixtilis 13 1 0 0 0 Basidiomycota Inocybe cf. nitidiuscula 37 1 0 0 0 Basidiomycota Inocybe cf. ochroalba 1 1 0 0 0 Basidiomycota Inocybe cf. pallidicremea 1 1 0 0 0

119

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Inocybe cf. pseudodestricta 9 1 0 0 0 Basidiomycota Inocybe cf. squarrosoides 10 1 0 0 0 Basidiomycota Inocybe cf. whitei 5 1 0 0 0 Basidiomycota Inocybe cincinnata 5 1 0 0 0 Basidiomycota Inocybe sp.1 15 1 0 0 0 Basidiomycota Inocybe sp.2 1 1 0 0 0 Basidiomycota Irpex lacteus 5 0 1 0 0 Basidiomycota Kuehneromyces cf. lignicola 18 0 1 0 0 Basidiomycota Laccaria cf. bicolor 17 1 0 0 0 Basidiomycota Laccaria cf. laccata 14 1 1 0 0 Basidiomycota Laccaria cf. striatula 13 1 0 0 0 Basidiomycota Laccaria nobilis 9 1 0 0 0 Basidiomycota Laccaria sp.1 8 1 0 0 0 Basidiomycota Laccaria sp.2 2 1 0 0 0 Basidiomycota Lactarius aff. hysginus 1 1 0 0 0 Basidiomycota Lactarius aff. nitidus 8 1 0 0 0 Basidiomycota Lactarius aff. uvidis 21 1 0 0 0 Basidiomycota Lactarius badiosanguineus 7 1 0 0 0 Basidiomycota Lactarius cf. affinis 5 1 0 0 0 Basidiomycota Lactarius cf. camphoratus 13 1 0 0 0 Basidiomycota Lactarius cf. picinus 20 1 0 0 0 Basidiomycota Lactarius cf. resimus 1 1 0 0 0 Basidiomycota Lactarius cf. trivialis 3 1 0 0 0 Basidiomycota Lactarius deterrimus 6 1 0 0 0 Basidiomycota Lactarius rufus 3 1 0 0 0 Basidiomycota Lactarius scrobiculatus 2 1 0 0 0 Basidiomycota Lactarius tabidus 6 1 0 0 0 Basidiomycota Lactarius thyinos 1 1 0 0 0 Basidiomycota Lactarius torminosus 2 1 0 0 0 Basidiomycota Leccinum cf. aurantiacum 8 1 0 0 0 Basidiomycota Lentaria byssiseda 2 0 1 0 0 Basidiomycota Lentinellus cf. cystidiosus 16 0 1 0 0 Basidiomycota Lentinellus micheneri 2 0 1 0 0 Basidiomycota Lentinellus ursinus 1 0 1 0 0 Basidiomycota Lentinus brumalis 7 0 1 0 0 Basidiomycota Lepiota castanea 1 0 0 1 0 Basidiomycota Lepiota cf. boudieri 1 0 1 1 0 Basidiomycota Lepiota cf. cortinarius 3 0 0 1 0 Basidiomycota Lepiota cf. cristata 2 0 1 1 0 Basidiomycota Lepiota cf. fuscosquamea 7 0 0 1 0 Basidiomycota Lepiota cf. magnispora 4 0 0 1 0 Basidiomycota Lepiota cf. pseudolilacea 4 0 0 1 0 Basidiomycota Lepiota clypeolaria 9 0 0 1 0

120

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Lepiota sp.1 1 0 0 1 0 Basidiomycota Lepista cf. nebularis 3 0 0 1 0 Basidiomycota Lepista cf. nuda 2 0 0 1 0 Basidiomycota Leucocybe cf. candicans 13 0 0 1 0 Basidiomycota Limacella cf. delicata 12 0 0 1 0 Basidiomycota Lycoperdon americanum 2 0 1 1 0 Basidiomycota Lycoperdon cf. molle 2 0 1 1 0 Basidiomycota Lycoperdon excipuliforme 9 0 0 1 0 Basidiomycota Lycoperdon perlatum 12 0 0 1 0 Basidiomycota Lycoperdon pyriforme 24 0 1 1 0 Basidiomycota Lycoperdon umbrinum 1 0 0 1 0 Basidiomycota Lyophyllum cf. mycenoides 1 0 0 1 0 Basidiomycota Marasmiaceae sp.1 4 0 0 1 0 Basidiomycota Marasmius cf. cohaerens 4 0 0 1 0 Basidiomycota Marasmius cf. wettsteinii 6 0 0 1 0 Basidiomycota Marasmius epiphyllus 26 0 0 1 0 Basidiomycota Megacollybia rodmanii 1 0 0 1 0 Basidiomycota Melanoleuca cf. melaleuca 1 0 0 1 0 Basidiomycota Mycena aff. hiemalis 2 0 1 0 0 Basidiomycota Mycena amicta 14 0 1 1 0 Basidiomycota Mycena cf. abramsii 1 0 1 0 0 Basidiomycota Mycena cf. acicula 20 0 0 1 0 Basidiomycota Mycena cf. alcalina 8 0 1 1 0 Basidiomycota Mycena cf. borealis 1 0 1 1 0 Basidiomycota Mycena cf. capillaripes 26 0 0 1 0 Basidiomycota Mycena cf. citrinomarginata 39 0 0 1 0 Basidiomycota Mycena cf. clavicularis 22 0 0 1 0 Basidiomycota Mycena cf. epipterygia 8 0 1 0 0 Basidiomycota Mycena cf. filopes 42 0 0 1 0 Basidiomycota Mycena cf. floridula 18 0 0 1 0 Basidiomycota Mycena cf. flos-nivium 8 0 0 1 0 Basidiomycota Mycena cf. leptocephala 39 0 1 1 0 Basidiomycota Mycena cf. maculata 2 0 1 1 0 Basidiomycota Mycena cf. metata 10 0 0 1 0 Basidiomycota Mycena cf. odorifera 5 0 0 1 0 Basidiomycota Mycena cf. praelonga 1 0 0 1 0 Basidiomycota Mycena cf. pura 49 0 0 1 0 Basidiomycota Mycena cf. robusta 21 0 0 1 0 Basidiomycota Mycena cf. sanguinolenta 33 0 1 1 0 Basidiomycota Mycena cf. tenax 3 0 0 1 0 Basidiomycota Mycena cinerella 5 0 0 1 0 Basidiomycota Mycena galopus 17 0 0 1 0 Basidiomycota Mycena haematopus 6 0 1 0 0

121

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Mycena leucogala 5 0 1 1 0 Basidiomycota Mycena polygramma 1 0 0 1 0 Basidiomycota Mycena purpureofusca 29 0 1 0 0 Basidiomycota Mycena silvae-nigrae 21 0 1 1 0 Basidiomycota Mycena sp.1 1 0 0 1 0 Basidiomycota Mycena stipata 2 0 1 0 0 Basidiomycota Mycena subcaerulea 18 0 1 0 0 Basidiomycota Mycena urania 6 0 0 1 0 Basidiomycota Mycena vulgaris 11 0 0 1 0 Basidiomycota Mycenella lasiosperma 1 0 1 0 0 Basidiomycota Mycorrhaphium adustulum 3 0 1 0 0 Basidiomycota Mythicomyces corneipes 1 0 0 1 0 Basidiomycota Myxarium cf. nucleatum 7 0 1 0 0 Basidiomycota Naucoria aff. silvaenovae 2 1 0 0 0 Basidiomycota Naucoria cf. bohemica 7 1 0 0 0 Basidiomycota Naucoria cf. escharoides 1 1 0 0 0 Basidiomycota Naucoria cf. paludosa 2 1 0 0 0 Basidiomycota Neofavolus cf. alveolaris 6 0 1 0 0 Basidiomycota Neofavolus sp.1 2 0 1 0 0 Basidiomycota Neofavolus sp.2 1 0 1 0 0 Basidiomycota Onnia tomentosa 1 0 1 0 1 Basidiomycota Panellus ringens 3 0 1 0 0 Basidiomycota Paxillus cf. vernalis 1 1 0 1 0 Basidiomycota Paxillus involutus 4 1 0 1 0 Basidiomycota Peniophora cf. erikssonii 3 0 1 0 0 Basidiomycota Peniophora rufa 17 0 1 0 0 Basidiomycota Peniophora sp.1 1 0 1 0 0 Basidiomycota Peniophora violaceolivida 4 0 1 0 0 Basidiomycota Perenniporia subacida 2 0 1 0 0 Basidiomycota Phaeoclavulina aff. flaccida 9 0 0 1 0 Basidiomycota Phaeoclavulina cf. abietina 10 0 0 1 0 Basidiomycota Phaeomarasmius cf. proximans 3 0 1 0 0 Basidiomycota Phanerochaete sp.1 1 0 1 0 0 Basidiomycota Phellinus betulinus 9 0 1 0 0 Basidiomycota Phellinus cf. alni 17 0 1 0 0 Basidiomycota Phellinus tremulae 5 0 1 0 1 Basidiomycota Phlebia radiata 1 0 1 0 0 Basidiomycota Phlebia tremellosa 1 0 1 0 0 Basidiomycota Phloeomana cf. speirea 15 0 1 0 0 Basidiomycota Pholiota cf. spumosa 7 0 1 0 0 Basidiomycota Pholiota cf. subsulphurea 8 0 1 0 0 Basidiomycota Pholiota flammans 1 0 1 0 0 Basidiomycota Picipes cf. melanopus 1 0 1 0 0

122

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Picipes sp.1 1 0 1 0 0 Basidiomycota Pleurocybella cf. porrigens 2 0 1 0 0 Basidiomycota Pleurotus cf. abieticola 1 0 1 0 0 Basidiomycota Pleurotus populinus 3 0 1 0 0 Basidiomycota Plicatura nivea 13 0 1 0 0 Basidiomycota Plicaturopsis crispa 11 0 1 0 0 Basidiomycota Pluteus aff. hispidulus 1 0 1 1 0 Basidiomycota Pluteus aff. tomentosulus 4 0 1 0 0 Basidiomycota Pluteus brunneidiscus 1 0 1 0 0 Basidiomycota Pluteus cf. luctuosus 9 0 1 0 0 Basidiomycota Pluteus cf. plautus 1 0 1 0 0 Basidiomycota Pluteus cf. rangifer 1 0 1 0 0 Basidiomycota Pluteus romellii 1 0 1 0 0 Basidiomycota Porodaedalea pini 3 0 1 0 0 Basidiomycota Porostereum crassum 1 0 1 0 0 Basidiomycota Postia cf. alni 20 0 1 0 0 Basidiomycota Postia cf. guttulata 1 0 1 0 0 Basidiomycota Postia cf. subcaesia 6 0 1 0 0 Basidiomycota Postia fragilis 1 0 1 0 0 Basidiomycota Protostropharia cf. semiglobata 1 0 0 1 0 Basidiomycota Psathyrella subamara 1 0 1 0 0 Basidiomycota Pseudoclitocybe cf. cyathiformis 1 0 1 0 0 Basidiomycota Pseudohydnum gelatinosum 15 0 1 0 0 Basidiomycota Punctularia strigozonata 3 0 1 0 0 Basidiomycota Radulomyces cf. confluens 1 0 1 0 0 Basidiomycota Ramaria cf. apiculata 4 1 1 0 0 Basidiomycota Ramaria cf. obtuissima 1 1 0 0 0 Basidiomycota Ramaria cf. rubella 2 1 1 0 0 Basidiomycota Ramaria cf. spinulosa 7 1 0 0 0 Basidiomycota Ramaria sp.1 2 1 0 0 0 Basidiomycota Ramaria stricta 1 1 1 0 0 Basidiomycota Resupinatus cf. applicatus 2 0 1 0 0 Basidiomycota Rhizocybe aff. vermicularis 17 0 1 1 0 Basidiomycota Rhodocollybia butyracea 3 0 0 1 0 Basidiomycota Rhodocollybia maculata 1 0 1 0 0 Basidiomycota Rhodofomes cajanderi 8 0 1 0 0 Basidiomycota Rhodofomes roseus 6 0 1 0 0 Basidiomycota Rhodophana cf. nitellina 20 0 0 1 0 Basidiomycota Rickenella cf. fibula 7 0 1 0 1 Basidiomycota Rickenella cf. mellea 3 0 0 0 1 Basidiomycota Rickenella swartzii 8 0 0 0 1 Basidiomycota Ripartites cf. metrodii 8 0 0 1 0 Basidiomycota Ripartites tricholoma 3 0 0 1 0

123

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Roridomyces roridus 25 0 0 1 0 Basidiomycota Russula acetolens 4 1 0 0 0 Basidiomycota Russula aff. firmula 3 1 0 0 0 Basidiomycota Russula aff. renidens 1 1 0 0 0 Basidiomycota Russula aff. sphagnophila 1 1 0 0 0 Basidiomycota Russula albonigra 1 1 0 0 0 Basidiomycota Russula cf. abietina 18 1 0 0 0 Basidiomycota Russula cf. adusta 3 1 0 0 0 Basidiomycota Russula cf. brevipes 2 1 0 0 0 Basidiomycota Russula cf. brunneola 12 1 0 0 0 Basidiomycota Russula cf. claroflava 3 1 0 0 0 Basidiomycota Russula cf. foetentula 4 1 0 0 0 Basidiomycota Russula cf. fragilis 31 1 0 0 0 Basidiomycota Russula cf. fragrantissima 8 1 0 0 0 Basidiomycota Russula cf. globispora 2 1 0 0 0 Basidiomycota Russula cf. hydrophila 40 1 0 0 0 Basidiomycota Russula cf. illota 1 1 0 0 0 Basidiomycota Russula cf. integriformis 6 1 0 0 0 Basidiomycota Russula cf. nauseosa 3 1 0 0 0 Basidiomycota Russula cf. olivina 4 1 0 0 0 Basidiomycota Russula cf. paludosa 19 1 0 0 0 Basidiomycota Russula cf. puellaris 3 1 0 0 0 Basidiomycota Russula cf. turci 2 1 0 0 0 Basidiomycota Russula cf. versicolor 4 1 0 0 0 Basidiomycota Russula sp.1 1 1 0 0 0 Basidiomycota Sarcodon cf. imbricatus 1 1 0 0 0 Basidiomycota Schizopora cf. paradoxa 1 0 1 0 0 Basidiomycota Simocybe cf. centunculus 3 0 1 0 0 Basidiomycota Simocybe cf. reducta 2 0 1 0 0 Basidiomycota Simocybe cf. sumptuosa 1 0 1 0 0 Basidiomycota Singerocybe cf. adirondackensis 13 0 0 1 0 Basidiomycota Sistotrema cf. confluens 2 0 1 1 0 Basidiomycota Sphaerobolus stellatus 1 0 1 0 0 Basidiomycota Steccherinum fimbriatum 3 0 1 0 0 Basidiomycota Steccherinum ochraceum 5 0 1 0 0 Basidiomycota Stereum cf. sanguinolentum 21 0 1 0 0 Basidiomycota Stereum striatum 5 0 1 0 0 Basidiomycota Stereum subtomentosum 6 0 1 0 0 Basidiomycota Strobilurus albipilatus 5 0 1 1 0 Basidiomycota Stropharia cf. hardii 1 0 1 1 0 Basidiomycota Suillus clintonianus 2 1 0 0 0 Basidiomycota Suillus tomentosus 1 1 0 0 0 Basidiomycota Tapinella panuoides 1 0 1 0 0

124

Phylum Morphospecies Occur.a ECMa WoodSapa LitterSapa Parasitica

Basidiomycota Tephrocybe cf. striaepilea 19 0 0 1 0 Basidiomycota Tephrocybe sp.1 2 0 0 1 0 Basidiomycota Thelephora cf. anthocephala 4 1 0 0 0 Basidiomycota Thelephora sp.1 5 1 0 0 0 Basidiomycota Tomentella testaceogilva 3 0 1 0 0 Basidiomycota Trametes betulina 4 0 1 0 0 Basidiomycota Trametes hirsuta 4 0 1 0 0 Basidiomycota Trametes ochracea 2 0 1 0 0 Basidiomycota Trametes pubescens 36 0 1 0 0 Basidiomycota Tremella cf. fuciformis 2 0 1 0 0 Basidiomycota Tremella mesenterica 5 0 1 0 1 Basidiomycota Trichaptum cf. abietinum 21 0 1 0 0 Basidiomycota Trichaptum cf. biforme 13 0 1 0 0 Basidiomycota Trichaptum cf. fuscoviolaceum 45 0 1 0 0 Basidiomycota Tricholoma cf. fulvum 1 1 0 0 0 Basidiomycota Tricholoma cf. myomyces 1 1 0 0 0 Basidiomycota Tricholoma cf. saponaceum 3 1 0 0 0 Basidiomycota Tricholoma inamoenum 4 1 0 0 0 Basidiomycota Tricholoma subluteum 1 1 0 0 0 Basidiomycota Tricholomopsis cf. flammula 4 0 1 0 0 Basidiomycota Tubaria cf. confragosa 2 0 1 0 0 Basidiomycota Tubaria cf. furfuracea 23 0 1 1 0 Basidiomycota Tylopilus cf. felleus 1 1 0 0 0 Basidiomycota Typhrasa aff. gossypina 7 0 1 0 0 Basidiomycota Xerocomus cf. ferrugineus 1 1 0 0 0 Basidiomycota Xeromphalina cf. campanella 27 0 1 0 0 Basidiomycota Xeromphalina cf. cauticinalis 5 0 0 1 0 Basidiomycota Xeromphalina cf. enigmatica 4 0 1 0 0 Basidiomycota Xeromphalina cf. kauffmanii 2 0 1 0 0 a Occur. = number of occurrences, ECM = ectomycorrhizal, WoodSap = wood saprotroph, LitterSap = litter saprotroph.