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Fleshy Saprobic and Ectomycorrhizal Fungal Communities Associated with Healthy and Declining Eastern Hemlock Stands in Great Smoky Mountains National Park
Richard Baird, C. Elizabeth Stokes, Alicia Wood-Jones, Mark Alexander, Clarence Watson, Glenn Taylor, Kristine Johnson, Thomas Remaley, and Susan Diehl

Southeastern Naturalist, Volume 13, Special Issue 6 (2014): 192–218

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Southeastern Naturalist R. Baird, et al. 2014 192 Vol. 13, Special Issue 6 Fleshy Saprobic and Ectomycorrhizal Fungal Communities Associated with Healthy and Declining Eastern Hemlock Stands in Great Smoky Mountains National Park Richard Baird1,*, C. Elizabeth Stokes1, Alicia Wood-Jones1, Mark Alexander2, Clarence Watson3, Glenn Taylor4, Kristine Johnson4, Thomas Remaley4, and Susan Diehl5 Abstract - Prior to the loss of Tsuga canadensis (Eastern Hemlock) stands in the Great Smoky Mountains National Park (GRSM), we collected baseline data during 2006–2009 at two locations (Copeland Creek and Gabes Mountain) regarding macrofungi that occur under this tree species. We studied macrofungi in order to understand the current and changing ectomycorrhizal and saprobic fungal community structure associated with healthy (imidicloprid-treated) and dying Eastern Hemlock stands and to contribute data for the All Taxa Biodiversity Inventory (ATBI) in GRSM. A total of 121 taxa representing 75 ectomycorrhizal, 1 pathogenic, and 45 saprobic species, were collected from 92, 59, and 106 sampling locations in 2006, 2007, and 2008, respectively. Macrofungal species richness, diversity, and evenness (E) in Copeland Creek were significantly greater (P = 0.05) in 2006 than 2007. Eighty percent of all fungi collected were found in Copeland Creek (487 m elevation) where trees are <75 years old; the remaining 20% were collected at Gabes Mountain (1158 m elevation) where trees are >150 years old. The most common taxa found across sampling locations included Russula fragrantissima (22.0%), Amanita citrina var. lavendula (Lavender False Death Cap; 17.1%), Austroboletus gracilis (Graceful Bolete; 14.6%), Laccaria laccata (Deceiver; 9.8%), and Russula russuloides (9.8%). Amanita cinereopannosa occurred at a low frequency overall (2.4% of trees sampled) but when present had high abundance. Total macrofungi and ectomycorrhizal fungi at Copeland Creek had significantly greater species richness, density, and evenness than at the Gabes Mountain site. Ectomycorrhizal fungi evenness values following imidicloprid soil-drenching treatments were significantly less uniform for macrofungi collected from within 5 m of trees treated in subplots with full rates of the chemical than for those near untreated trees (E = 0.2 and 0.4, respectively). In addition, there was a numerical trend towards significantly less diversity and evenness at the full chemical application rate compared to the half rate or untreated control plots. There were no differences in the occurrence of saprobic fungi between chemically treated and control trees. Associated vegetation had a significant impact on the occurrence of macrofungi. Across both locations, a total of 37 plant, shrub, or tree species were identified; Acer pensylvanicum (Striped Maple) with an overall abundance of 11.0%, Pyrularia pubera (Buffalo Nut; 10.5%), Ilex opaca (American Holly; 8.2%), Calycanthus floridus (Eastern Sweetshrub; 4.8%), and Rhododendron maximum (Great Rhododendron; 4.7%), were the most common associated species. Species richness, diversity, and evenness of the associated vegetation were significantly different between locations. Evenness data for plant species abundance was not equal, but varied within and across locations. 1BCH-EPP Department, Box 9655, Mississippi State University, Mississippi State, MS 39762. 2Carbon Institute, University of Tennessee, Knoxville, TN 37996. 3University of Arkansas Division of Agriculture, 2404 North University Avenue, Little Rock, AR 72207. 4Vegetation Unit, Great Smoky Mountains National Park, 107 Park Headquarters, Gatlinburg, TN 37738. 5Forest Products, Mississippi State University, Mississippi State, MS 39762. *Corresponding author - rbaird@plantpath.msstate.edu. Manuscript Editor: Kevin Kuehn Forest Impacts and Ecosystem Effects of the Hemlock Woolly Adelgid in the Eastern US 2014 Southeastern Naturalist 13(Special Issue 6):192–218 Southeastern Naturalist 193 R. Baird, et al. 2014 Vol. 13, Special Issue 6 Introduction Tsuga canadensis (L.) Carrière (Eastern Hemlock, hereafter, Hemlock) is the most shade-tolerant tree species in the eastern US; it is able to survive in the understory at 5 percent full sunlight or greater (Baker 1949, Godman and Lancaster 1990, Graham 1954). Although reproduction can occur on bare, exposed soil, Hemlock usually becomes established under a closed canopy, and over the course of a century or more, it emerges from the subcanopy to become a dominant climax species (Tubbs 1977). The dense, evergreen canopy associated with mature hemlock forests creates a unique environment that is a critical habitat for many animal and plant species; more than 120 vertebrate species require mature Hemlock stands (DeGraaf et al. 1992, Ward et al. 2004). A wide variety of aquatic species are associated exclusively with streams sheltered by Hemlock (Snyder et al. 2004). Evans (2002) reported that Salvelinus fontinalis Mitchill (Brook Trout) populations and macroinvertebrate diversity were significantly greater in Hemlock-shaded streams, which had low summer temperatures and were less likely to become dry than in open canopy stands. Adelges tsugae Annand (Hemlock Woolly Adelgid [HWA]) defoliates and kills Hemlocks, often creating large gaps in the canopy. Canopy disturbance can catalyze a cascade of events: more sunlight reaching the soil elevates soil temperature, which increases microbial activity, and results in enhanced soil N (Sirulnik et al. 2005). Hemlock mortality associated with HWA infestation has been shown to alter soil conditions and fungal communities. Sirulnik et al. (2005) reported that higher NO3 - levels occurred in HWA-infested areas than healthy Hemlock-dominated stands, suggesting that Hemlock mortality is associated with increased N. Furthermore, examination of ectomycorrhizal roots suggested that HWA-induced defoliation may result in reduced ectomycorrhizal density and species richness. Thus, reduced macrofungal abundance and diversity, particularly of ectomycorrhizal species, can be expected after heavy HWA infestations in Hemlock forests (Lewis et al. 2008). Grime et al. (1987) suggested that mycorrhizal interactions play a role in the mediation of plant competition, and may even influence species composition. A study by van der Heijden et al. (1998) showed that higher levels of fungal diversity resulted in higher levels of plant diversity and productivity of grassland ecosystems. Conservation of mycorrhizal and saprobc fungal diversity is crucial to maintenance of plant diversity and plant-community composition in grasslands, as well as in other ecosystems such as boreal forests, where the fungal community is known to influence allocation of resources between plant species (Read 1991, Simard 1997). Because of the apparent ecological importance of ectomycorrhizal fungi, it is critical to determine their associates in healthy Hemlock stands, and to preserve these fungi for reintroduction if HWA control measures are effective and allow Hemlock reforestation. Previous work has shown that the neonicotinoid insecticide imidacloprid is effective in controlling HWA in landscape environments (Cowles et al. 2004, Doccola et al. 2003, Steward and Horner 1994, Webb et al. 2003). However, the secondary impacts of the chemical on other insects and arthropod species have only recently been examined (Reynolds 2008). Moore et al. (1988) suggested that arthropods Southeastern Naturalist R. Baird, et al. 2014 194 Vol. 13, Special Issue 6 play a major role in regulating micro- and mesobiota in below-ground detrital food webs. Many fungi are consumed by mycophagous Collembolans, oribatids, enchytraeids, and Dipterans (Moore et al. 1988; Newell 1984a, b; Shaw 1992). Many Coleopterans (beetles), Hymenopterans (ants), and Isopterans (termites) are fungivores (Martin 1987, Shaw 1992), and are often fungi-species–specific or selective feeders (Bills et al. 2004). Van der Drift and Jansen (1977) suggested that fungal activity might be stimulated by grazing, whereas Hanlon and Anderson (1979) showed that grazing resulted in a decrease in fungal populations with a subsequent increase in bacterial numbers. The soil-drench method of imidacloprid application impacts insect communities in the litter and soil (Mullin et al. 2005). Recent research in a Hemlock forest demonstrated that imidacloprid killed Collembolan species throughout the drench zone following treatment (Reynolds 2008). The imidacloprid- induced removal of fungivores from the Hemlock rhizosphere could result in a variety of effects on soil fungi. Negative effects could include the proliferation of plant pathogenic fungi in the absence of fungivorous insects. Furthermore, nutrient cycling could be disrupted with the removal of litter-degrading insects, as simplified substrates become unavailable for use by beneficial fungi (Ingham and Thies 1996). Ectomycorrhizal fungal populations would likely be affected by the imidaclopridinduced mortality of soil invertebrates. Disturbances that change forest tree species composition or completely remove a dominant species from an area can have large and lasting impacts on ectomycorrhizal abundance and diversity (Durall et al. 1999, Visser 1995). A recent study carried out over 12 months reported that ectomycorrhizal hyphal abundance in soils was significantly reduced following experimental girdling of Notholithocarpus densiflorus (Hook & Arn.) Manos, Cannon, and S.H. Oh (Tan Oak) (Bergemann et al. 2013). The authors concluded that lower hyphal levels in soil might have been due to the reduced carbon availability caused by tree mortality (Kaiser et al. 2010, Pena et al. 2010). Therefore, loss of Hemlock in the eastern US could affect species richness, diversity and evenness of both saprobic and ectomycorrhizal fungal communities in these forest ecosystems. Little is known about the character and function of Hemlock-associated fungi, and until now, no study has been conducted on the assemblage of soilborne fungi associated with healthy pure stands of mature Hemlock. Without baseline data on pre-HWA-infested, pre-imidacloprid-treated Hemlock, there would be no reference for comparison with the forest following Hemlock decline. Therefore, the purpose of this study was to obtain baseline data on the species composition, richness, and diversity of fleshy saprobic and ectomycorrhizal fungi that formed basidiomata associated with uninfested, old-growth Hemlock and to determine if imidacloprid treatments impact fungal communities. Methods Study area We conducted our study April 2006–September 2008 at 2 locations in Great Smoky Mountains National Park (GRSM): Copeland Creek (elevation 487 m) and Gabes Mountain (elevation 1158 m). Both sites consisted of pure stands of mature Southeastern Naturalist 195 R. Baird, et al. 2014 Vol. 13, Special Issue 6 Hemlock and were the same ones used in a previous study of Hemlock rhizosphere microbial communities (Baird et al. 2014 [this issue]). Sampling methodology We determined plot sizes and shapes based on available healthy Hemlock stands (see below). At each site, we established one 48.8 x 48.8-m plot which we divided into sixty-four 6.1 x 6.1-m subplots in an 8 x 8 grid to more precisely define macrofungal sporophore abundance and diversity. We counted all Hemlock trees within the subplots, measured their diameter at breast height (dbh), and determined tree heights using a Suunto® clinometer (http://www.suunto.com). All other tree species and lesser woody vegetation were inventoried and characterized within the subplots (Baird et al. 2014 [this issue]). We recorded tree-crown health-indicator data for each Hemlock tree within each subplot at both locations. Crown-condition classification included vigor class, uncompact live-crown ratio, crown light exposure, crown position, crown density, crown dieback, and foliage transparency (Schomaker et al. 2007). These indicators of crown health were based on pure stands of Hemlock rated at vigor 1 or >35% crown ratio, which indicated high vigor or healthy sites. Tree-climbers in the area provided us with data they had collected the month before the study, which indicated that these sites had approximately 30% foliage infestation at the beginning of the study, but needles remained intact and the crowns healthy. We also obtained vigor data at the end of the third year of the study, which included insect infestation levels based on tree-climber data. We selected 20 Hemlock trees with dbh >20 cm. All chemical application-rates and treatments followed those previously described by Baird et al. (2014 [this issue]). Selected trees were at least 10 m apart to avoid interactions between roots of nearby trees during treatments. To experimentally assess the effect of imidacloprid on soil fungi, we subjected each of the 20 selected trees per plot to 1 of 2 imidacloprid treatments or a non-treatment control. We replicated each treatment on a minimum of 6 randomly selected trees per location and applied all chemical treatments in September 2006. Treatments included: 1) a single imidacloprid (Merit® SP a.i. 75) application at rate of 11.8 ml/1 cm dbh, 2) a single application at 5.9 ml/1 cm dbh, and 3) a non-treatment control. We sampled all macrofungal sporocarps each month from April–September in 2006–2008 within each subplot at both locations. We collected all macrofungi present on the forest floor (terrestrial) or on woody debris within the subplot and noted the associated tree species when present. We transported specimens to laboratory facilities on the day of collection and stored them at 4 °C for up to 12 h. For each specimen, we recorded macromorphological characteristics of fresh basidiomata (Largent et al. 1977). We photographed each fresh specimen in the field and laboratory for future reference and to aid in identification. We determined color for each specimen using the Methuen Handbook of Colour (Kornerup and Wanscher 1978). Following collection data from fresh specimens, we preserved basidiomata by warm-air drying and stored them for later microscopic data collection (Largent et al. 1977). We rehydrated dried tissues Southeastern Naturalist R. Baird, et al. 2014 196 Vol. 13, Special Issue 6 with 10% KOH or 95% ETOH and then rinsed them in distilled water. We used a compound microscope to determine micromorphological details including characteristics of basidia, hyphae, cystidia, and spores. Voucher collections from the study are housed at MSU herbarium (R. Baird, MSU collections H 1-197). We obtained voucher specimens from TENN (University of Tennessee Herbarium, Knoxville, TN) to confirm our identifications. We obtained monthly precipitation totals recorded at Elkmont (TN11) in the GRSM and reported by the National Atmospheric Deposition Program (http://nadp.sws.uiuc.edu/). Statistical analyses We randomly assigned the two insecticide treatments and the untreated control treatment to trees within each location, and we analyzed the data as a completely randomized design within each location. All data were analyzed using analysis of variance (ANOVA). Fisher’s protected least significant difference test (P < 0.05) was performed to compare means. Pearson correlation coefficients were estimated using Proc CORR- of SAS® (SAS Institute 1999) to evaluate the association among fungal diversity values and tree-health indicators within respective subplots. A Bonferroni correction was applied to adjust for multiple correlative comparisons. We calculated species richness and diversity indices, and coefficient of community or beta diversity (Dyke 2003) for ectomycorrhizal and saprobic taxa based on our counts of the sporocarps that we collected and identified. We used the total number of basidiomata for each species within the plots to calculate biodiversity. The biodiversity indices calculated included species richness (n), Shannon-Weaver species diversity index (H'), Shannon species evenness index (E), and coefficient of community (CC) (Price 1997; Stephenson 1989; Stephenson et al. 2004). The Shannon diversity index is highly sensitive to species evenness and the presence of rare species. It is also the most widely used measurement of biodiversity, which allowed us to compare our results with those from previous studies (Magurran 2004). We also calculated relative frequency of fungal occurrence. Where appropriate, data were further analyzed using one-way analysis of variance (ANOVA) using the general linear models procedure (Proc GLM) of the Statistical Analyses System Software (SAS® Institute, Inc. 1999). Results We collected a total of 121 species of macrofungi representing 66 genera during the 3-year study (Appendix A), which comprised 45 saprobic, 1 parasitic, and 75 ectomycorrhizal species. At both study sites, a total of 77.6% of the collections were located <5 m from the Hemlock trees selected for chemical treatment or within control plots. Of the total species identified, 80.0% were collected from Copeland Creek and 20.0% from Gabes Mountain. The macrofungi collected included 92, 59, and 106 taxa obtained from 2006, 2007, and 2008, respectively. The dominant species, accounting for 27.5% of collections, included Russula fragrantissima Romagn. (at 22.0% of trees sampled), Amanita citrina var. lavendula (Coker) Sartory & Maire (Lavender False Death Cap; 17.1%), Austroboletus gracilis (Peck) Wolfe Southeastern Naturalist 197 R. Baird, et al. 2014 Vol. 13, Special Issue 6 (Graceful Bolete; 14.6%), Laccaria laccata (Scop.) Cooke (Deceiver; 9.8%), and Gymnopus dryophilus (Bull.) Murrill; 9.8%). Amanita cinereopannosa Bas had low frequency (2.4% of trees sampled), but when present it was abundant: it accounted for 96% of all taxa collected. However, the majority of macrofungi taxa occurred at low abundance and frequency. For example, 64 taxa were represented by only a single collection date over the 3-year study. Sporophore formation is highly influenced by precipitation throughout a growing season. March–April precipitation totals measured at the GRSM Tremont station in 2006, 2007, and 2008 were 17.0, 19.1, and 14.2 cm, respectively (Fig. 1). Monthly rainfall was highly variable but generally declined later in the season. Notably, May–August 2007 rainfall was extremely low. At Copeland Creek, macrofungi and ectomycorrhizal species had significantly greater richness, diversity, and evenness than at those Gabes Mountain (Table 1), whereas analysis of saprobic fungi data showed similar values at both locations. Ectomycorrhizal species evenness had significantly lower uniformity for the full-rate imidacloprid application (E = 0.2) than for the untreated control (E = 0.4). In addition, there was a trend toward less diversity and evenness in plots that received the full chemical rate treatment compared to those that received the half-rate treatment or untreated control trees. Among saprobic fungi, there were no significant differences between sites, chemical rates or years (data not shown). Fungal diversity and evenness values were significantly greater in 2006 than in 2007 for both total and ectomycorrhizal macrofungi at Copeland Creek (Table 2). In 2006 and 2008, total and ectomycorrhizal fungi at Copeland Creek had significantly greater richness, diversity, and evenness than at Gabes Mountain. At Copeland Creek, richness, diversity, and evenness values in 2006 were significantly greater than in 2007 for both total and Figure 1. Monthly precipitation totals recorded by the National Atmospheric Deposition Program, Great Smoky Mountains National Park, Sevier County, TN 2006–2008. Southeastern Naturalist R. Baird, et al. 2014 198 Vol. 13, Special Issue 6 ectomycorrhizal fungi. In contrast to the ectomycorrhizal species, the results for saprobic macrofungi were similar during the 3 years of the study (data not shown). We compared coefficient of community values for ectomycorrhizal, saprobic, and total macrofungi and derived an average CC value of 0.34. A total of 19.4% of all fungi identified, 15.2% of ectomycorrhizal fungi, and 26.9% of saprobic macrofungi occurred at both sites. We compared the species occurrence data by group (ectomycrrhizal fungi and saprobic fungi) and total fungi by imidicloprid Table 1. Species richness, diversity, and evenness of total and ectomycorrhizal macrofungi associated with Eastern Hemlock by site, rate, and year, Great Smoky Mountains National Park 2006–2008. Copeland Creek is a second-growth mature Hemlock forest, 487 m elevation. Gabes Mountain is a virgin old-growth forest at 1158 m elevation. n = species richness, H' = Shannon diversity index, and E = Shannon evenness index across two treatments and control. Rate indicates imidacloprid soil-drench application at full dosage (11.8 ml/cm dbh), half (5.9 ml/cm dbh) dosage, and untreated control rates. Within each of the data grouping, means followed by the same letter in the same column are not significantly different using Fisher’s LSD test (P > 0.05). n H' E Site Total macrofungi predictors Copeland Creek 3.1a 0.8a 0.6a Gabes Mountain 0.8b 0.2b 0.3b LSD 0.6 0.2 0.1 Ectomycorrhizal Macrofungi Predictors Copeland Creek 2.4a 0.6a 0.5a Gabes Mountain 0.2b <0.1b <0.1b LSD 0.5 0.2 0.1 Rate Total macrofungi predictors 11.8 ml/cm dbh 1.8a 0.5a 0.4a 5.9 ml/cm dbh 2.2a 0.6a 0.5a Untreated control 1.8a 0.6a 0.4a LSD 0.8 0.2 0.2 Ectomycorrhizal Macrofungi Predictors 11.8 ml/cm dbh 1.2a 0.3a 0.2a 5.9 ml/cm dbh 1.5a 0.4a 0.3ab Untreated control 1.3a 0.4a 0.4b LSD 0.7 0.2 0.1 Year Total macrofungi predictors 2006 1.8a 0.5a 0.5a 2007 1.1a 0.2b 0.3b 2008 1.3a 0.4ab 0.4ab LSD 0.9 0.2 0.2 Ectomycorrhizal Macrofungi Predictors 2006 1.3a 0.4a 0.3a 2007 0.6a 0.1b 0.1b 2008 0.9a 0.3ab 0.3a LSD 0.7 0.2 0.2 Southeastern Naturalist 199 R. Baird, et al. 2014 Vol. 13, Special Issue 6 half-rate vs. full rate treatments, imidicloprid full-rate treatment vs. control, and imidicloprid half-rate treatment vs. control. The CC values were 0–0.518 and averaged 0.34. Species overlaps were 21.6% for the full-rate vs. control comparisons, 33.3% for the full-rate vs. half rate comparisons, and 48.7% for the half-rate vs. control comparisons. Macrofungal species frequencies by treatment (full, half, and control) were 35.6%, 34.5%, and 27.5%, respectively for ectomycorrhizal taxa and 22.2%, 31.6%, and 27.6%, respectively, for saprobic fungi. We observed no other consistent trends for richness, diversity, or evenness of fungal taxa. Pearson correlations were significant between fungal (richness, diversity and evenness of total ectomycorrhizal and saprobic fungi) and plant (abundance, richness, diversity and evenness) biodiversity measures (Table 3). Table 2. Richness, diversity, and evenness of total and ectomycorrhizal macrofungi associated with Eastern Hemlock, site by year interaction at two locations in Great Smoky Mountains National Park 2006–2008. Copeland Creek is a second-growth mature Hemlock forest, 487 m elevation. Gabes Mountain is a virgin old-growth forest, 1158 m elevation. Species richness = mean number of unique taxa, species diversity = Shannon diversity index, and evenness = Shannon evenness index. Means followed by the same lowercase letter in the same column for each data grouping, or by the same uppercase letter in the same row (in parentheses) are not significantly different using Fisher’s LSD test (P > 0.05). Site 2006 2007 2008 LSD Total macrofungi predictor Species richness Copeland Creek 3.0 a (A) 1.3 a (B) 2.0 a (AB) 1.1 Gabes Mountain 0.5 b (A) 0.8 a (A) 0.6 b (A) 0.6 LSD (P < 0.05) 1.1 0.7 1.0 Species diversity Copeland Creek 0.9 a (A) 0.3 a (B) 0.6 a (AB) 0.3 Gabes Mountain 0.1 b (A) 0.2 a (A) 0.2 b (A) 0.2 LSD (P < 0.05) 0.3 0.3 0.3 Species evenness Copeland Creek 0.8 a (A) 0.3 a (AB) 0.6 a (B) 0.3 Gabes Mountain 0.2 b (A) 0.2 a (A) 0.2 b (A) 0.3 LSD (P < 0.05) 0.2 0.3 0.3 Ectomycorrhizal macrofungi predictor Species Richness Copeland Creek 2.4 a (A) 1.0 a (B) 1.5 a (AB) 1.0 Gabes Mountain 0.1 b (A) 0.2 b (A) 0.2 b (A) 0.3 LSD (P < 0.05) 1.0 0.6 0.7 Species diversity Copeland Creek 0.7 a (A) 0.2 a (B) 0.5 a (AB) 0.3 Gabes Mountain 0.0 b (A) 0.0 a (A) 0.0 b (A) 0.1 LSD (P < 0.05) 0.3 0.2 0.2 Species evenness Copeland Creek 0.6 a (A) 0.2 a (B) 0.5 a (AB) 0.3 Gabes Mountain 0.0 b (A) 0.0 b (A) 0.1 b (A) 0.1 LSD (P < 0.05) 0.2 0.2 0.2 Southeastern Naturalist R. Baird, et al. 2014 200 Vol. 13, Special Issue 6 Biodiversity measures for total and ectomycorrhizal macrofungi were significantly correlated with tree health (Table 4). Richness, diversity, and evenness were positively correlated with live-crown ratio and crown density, but negatively correlated Table 4. Correlation between macrofungi associated with Eastern Hemlock and tree health at two locations in Great Smoky Mountains National Park, 2006–2007. Tree health parameters measured according to protocols of The Forest Inventory and Analysis Program of the US Department of Agriculture– Forest Service (Schomaker et al. 2007). Abundance = number of macrofungi collections, richness = number of fungal taxa, diversity = Shannon diversity index, evenness = Shannon evenness index, and * = Pearson correlation coefficient (r-value) is significantly different from zero (P < 0.05). Tree health Live-crown Crown Crown Branch Tree ratio Density transparency dieback diameter Total macrofungi predictor Abundance 0.3* 0.2* -0.2* -0.2 -0.3* Richness 0.3* 0.2* -0.2* -0.2* -0.3* Diversity 0.3* 0.2* -0.2* -0.2* -0.3* Evenness 0.3* 0.2* -0.1* -0.1 -0.3* Ectomycorrhizal macrofungi predictor Abundance 0.3* 0.2* -0.2* -0.2 -0.3* Richness 0.4* 0.2* -0.3* -0.2 -0.3* Diversity 0.4* 0.2* -0.3* -0.2* -0.3* Evenness 0.4* 0.2* -0.3* -0.3* -0.4* Saprobic macrofungi predictor Abundance 0.1 0.1 -0.1 <-0.1 -0.1 Richness 0.1 0.1 -0.1 <-0.1 -0.1 Diversity 0.1 0.2* -0.2 <-0.1 -0.1 Evenness 0.1 0.2* -0.2 <0.1 -0.1 Table 3. Correlations among predictor variables for macrofungi associated with Eastern Hemlock at two locations in Great Smoky Mountains National Park, Sevier Co., TN, 2006–2008. Abundance = number of individual macrofungi collections, richness = number of fungal taxa, diversity = Shannon diversity index, Evenness = Shannon evenness index, and * = Pearson correlation coefficient (r-value) is significantly different from zero (P < 0.05). Richness Diversity Evenness Total macrofungi predictor Abundance 1.0* 1.0* 0.7* Richness 0.9* 0.7* Diversity 0.9* Ectomycorrhizae predictor Abundance 1.0* 1.0* 0.7* Richness 1.0* 0.8* Diversity 0.9* Saprobe predictor Abundance 1.0* 0.9* 0.8* Richness 0.9* 0.8* Diversity 1.0* Southeastern Naturalist 201 R. Baird, et al. 2014 Vol. 13, Special Issue 6 with crown transparency and diameter. Diversity and evenness of saprobic macrofungi were positively correlated with crown density and transparency. As in the previous study by Baird et al. (2014 [this issue]) at the same sites, we identified 37 species of woody plants or shrubs and trees. The most frequently occurring species included Acer pensylvanicum L. (Striped Maple; 11.0%); Pyrularia pubera Michx. (Buffalo Nut; 10.5%); Ilex opaca Aiton (American Holly; 8.2%); Calycanthus floridus L. (Eastern Sweetshrub; 4.8%); and Rhododendron maximum L. (Great Rhododendron; 4.7%). Species abundance, richness, and diversity were significantly higher at the Copeland Creek site (39.3, 10.3, and 1.8, respectively) than at Gabes Mountain (22.0, 5.6, and 1.4, respectively). CC values for woody vegetation comparisons between sites indicated 52% species similarity. When biodiversity measures for total and ectomycorrhizal macrofungi were compared with those for woody vegetation, all comparisons were statistically significant at α = 0.05 (Table 5). However, biodiversity measures for saprobic macrofungi had no significant correlation with those for woody vegetation, with CC values of -0.1 for abundance or richness and <0.1 for diversity (data not shown). Each pair-wise comparision of biodiversity measures for woody vegetation—richness, diversity, and evenness—showed a positive correlation (Table 6). Discussion This research was the first major survey in GRSM to obtain baseline data on the species composition, richness, and diversity of fleshy saprobic and ectomycorrhizal fungi that form basidiomata associated with Hemlock unifested with HWA and to determine if imidacloprid treatments impact fungal communities. We also attempted to identify important and rare endemic fungal symbionts before habitat changes due Table 5. Correlations between total and ectomycorrhizal macrofungi associated with Eastern Hemlock and associated woody vegetation at two locations in Great Smoky Mountains National Park, Sevier Co., TN, September 2006–2008. Associated woody vegetation inventoried in sixty-four 6 x 6-m subplots per site. Abundance = number of macrofungi collections and number of individual woody plants. richness = number of fungal taxa and number of woody plant species, diversity = Shannon diversity index, evenness = Shannon evenness index, and * = Pearson correlation coefficient (r-value) is significantly different from zero (P < 0.05). Associated woody vegetation Abundance Richness Diversity Evenness Total macrofungi predictor Abundance 0.3* 0.3* 0.3* 0.6* Richness 0.3* 0.3* 0.3* 0.8* Diversity 0.4* 0.3* 0.3* 0.9* Evenness 0.3* 0.3* 0.3* 0.9* Ectomycorrhizal macrofungi predictor Abundance 0.4* 0.3* 0.3* 0.6* Richness 0.4* 0.4* 0.3* 0.8* Diversity 0.5* 0.4* 0.3* 0.9* Evenness 0.5* 0.4* 0.3* 0.9* Southeastern Naturalist R. Baird, et al. 2014 202 Vol. 13, Special Issue 6 to HWA infestations or other causes occur. However, almost all fungi identified in the list are present in numerous habitats in the southern Appalachian Mountains or are not specifically associated with Hemlocks, but are found with other tree species within or bordering Hemlock-dominated stands (Baird et al. 2014 [this issue]). The majority of the macrofungi that we identified occur in the eastern United States and exhibit a broad geographical range. Even though 71 of the 121 species we collected were found only once during the study, we cannot assume that they were rare. Throughout the study, we observed many of these rarely collected taxa just outside the plots (R. Baird, pers. observ.). We suspect that they will be impacted by the same environmental parameters affecting trees in our study plots; apparently age of associated trees can influence basidiomata formation. It is important to collect baseline data on saprobic and ectomycorrhizal fungi for the ATBI in GRSM because many of the taxa that occur in Eastern Hemlock ecosystems are becoming regionally or globally extinct (e.g., stipitate Hydnum spp.) and their absence may impact natural reforestation in the future (Baird et al. 2013). Similar surveys have been conducted in regenerating Hemlock forests at the northern edge of their range (McLenon-Porter 2008), on Quercus spp. (oak) seedlings in the southern Appalachians (Walker et al. 2005), and in other forest types (Burke et al. 2009, DeBellis et al. 2006, Dickie et al. 2009), including mature stands of Tsuga heterophylla Raf. Sarg. (Western Hemlock) in the Pacific Northwest (Wright et al. 2009). These and other floristic studies, while conducted across a variety of host species and forest types, provide some basis for comparison with the results of the current study, as discussed below. Our study sites were different in age and floristic composition. We selected the Copeland Creek and Gabes Mountain sites based on previous National Park Service surveys that identified mature Hemlock stands with limited presence of HWA damage and a tree vigor rating of 1 (see Schomaker et al. 2007). We wanted to conduct our study in healthy Hemlock stands without prior imidicloprid application, low insect-infestation levels, and similar age; few other sites in the GRSM were available or represented greater age uniformity prior to establishment of this study. The Copeland Creek site was composed of mature second-growth Hemlock forest with intermittent mixed hardwoods. Additionally, a component of large Pinus strobus L. (White Pine) at Copeland Creek, recently killed by Dendroctonus frontalis Zimmermann (Southern Pine Bark Beetle) infestation, resulted in diverse Table 6. Correlations among predictor variables for Eastern Hemlock-associated woody vegetation from two locations in Great Smoky Mountains National Park, September 2006. Associated woody vegetation inventoried in sixty-four 8 x 8-m subplots per site. Abundance = number of individual woody plants, richness = number of plant species, diversity = Shannon diversity index, evenness = Shannon evenness index, and * Pearson correlation coefficient (r-value) is significantly different from zero (P < 0.05). Richness Diversity Evenness Abundance 0.802* 0.606* 0.752* Richness 0.901* 0.533* Diversity 0.662* Southeastern Naturalist 203 R. Baird, et al. 2014 Vol. 13, Special Issue 6 woody plant regeneration in the understory and abundant coarse woody debris on the forest floor (R. Baird, pers. observ.). The Gabes Mountain site was a highelevation virgin Hemlock forest with minor components of large Halesia carolina L. (Carolina Silverbell) and Tilia americana L. (American Basswood) and a sparse understory dominated by Striped Maple and Great Rhododendron. In addition, this location is included in a large tract of old-growth Hemlock that is being managed as a long-term ecological and genetic conservation stand by the National Park Service (Johnson et al. 2008; T. Remaley, pers. comm.). Vigor ratings for both sites at the end of the study recorded trees ranging from vigor = 3–5 (Schomaker et al. 2007); some dead trees were present, and the insect infestation level was 95%. The chemically treated trees in our study plots tended to have the higher vigor ratings. Differences in tree vigor between the start and end of the study may have impacted fungal community data such as species richness, diversity, and evenness, but we could not make comparisons or conduct statistical analyses without monthly vigor rating data. We identified a total of 121 species of macrofungi during the 3 years of monthly collections. The majority of fungal taxa identified were listed as mycorrhizal mutualists in previous reports (Appendix A). The importance of mycorrhizal fungi to individual plants and plant communities is well established (Smith and Read 1997). Mycorrhizal fungi, which included 113 of the taxa we collected, were high in richness and diversity. In comparison to 14 previous studies compiled by Horton and Bruns (2001), greater species richness was found only in a mature Pseudotsuga menziesii (Mirb.) Franco (Douglas-fir)/Western Hemlock forest where 200 taxa were recorded (Luoma et al. 1997). The results of our study suggest that mature Hemlocks in the southern Appalachians host some of the richest and most diverse fungal communities observed in any temperate forest. Based on data from the TENN Fungal Herbarium and other sources, we determined that most of the fungal species identified in this study had been previously collected in GRSM (Appendix A). However, none of these taxa were reported specifically as components of Hemlock forests in GRSM. From the viewpoint of conservation biology, the fungal richness and diversity we documented at our study sites are important findings and underscore the value of these forests as important areas of biological diversity. As previously discussed, host-tree health, as measured by canopy defoliation, was the most significant parameter affecting fungal communities. Richness, diversity, and evenness of ectomycorrhizal and total macrofungi were significantly correlated with Hemlock tree health (Table 3). Trees with greater canopy density and live-crown ratio were consistently associated with richer, more diverse fungal communities. Defoliation increased over the course of the study by an average of 12.6% at Copeland Creek and 27.8% at Gabes Mountain. As canopy defoliation advanced, associated macrofungal communities were significantly reduced in richness, diversity, and evenness. Saprobic macrofungi appeared to be less sensitive to canopy defoliation, but diversity and evenness of these species were still significantly reduced. The reduction in macrofungi associated with host tree defoliation could be due to decreased availability of carbohydrates from host trees (Lewis et Southeastern Naturalist R. Baird, et al. 2014 204 Vol. 13, Special Issue 6 al. 2008), as well as microsite changes at the forest floor (Tedersoo et al. 2008). Microenvironmental variation within and among different microhabitats is known to affect fungal species richness and diversity (Baird et al. 2007, Stephenson 1989). Hemlock defoliation affects forest-floor microenvironments by increasing light and temperature and decreasing soil moisture. In the current investigation, we observed seasonal patterns among monthly collections of macrofungi. We collected taxa such as Amanita, Clitocybe, and several Russula species only in the spring and early summer, while Tricholoma, Cortinarius, and hydnaceaous species occurred later in the season. A number of studies have linked intra-annual variability in ectomycorrhizal colonization to climatic variation (Harvey et al. 1978, Rastin et al. 1990, Swaty et al. 1998, Vogt and Edmonds 1980, Walker et al. 2008). It is notable, however, that fungal diversity followed a decreasing trend in 2008 that did not reflect rainfall patterns and suggests other causes. Furthermore, saprobic fungi were evidently not affected by yearly differences in precipitation. The decrease in ectomycorrhizal diversity over the course of the study may be attributable to aforementioned causal factors relating to host-tree health and the impact of HWA. Evenness is an important estimator of diversity and measures the degree of equality among the relative species abundances (Drobner et al. 1998). The low overall evenness of macrofungi observed at Copeland Creek and Gabes Mountain (E = 0.6 and 0.3, respectively) was due to high numbers of rarely observed species and a few dominant taxa. Of 121 total macrofungi taxa, 71 occurred only once during the study (Appendix A). Further, 9 taxa accounted for 36.6% of all macrofungi collections. These results must be interpreted with caution due to the highly irregular and ephemeral nature of ectomycorrhizal sporophore production, and because high numbers of rare species can reduce the power of statistical tests (Krebs 1989). Molecular data of root-tissue-sequenced fungi did not show a similar effect on evenness (Baird et al. 2013). Because ectomycorrhizal fungi require photosynthetically derived carbohydrates from their host trees for growth and sporophore production (Smith and Read 2008), differences in tree health and vigor almost certainly affect ectomycorrhizal communities. The forest at Copeland Creek had numerous standing dead White Pines, killed by Southern Pine Bark Beetle. This recent disturbance effectively released the canopy-enclosed Hemlocks from competition, allowing increased photosynthesis and growth. Fine-root mass increases on vigorously growing trees and provides niches for diverse ectomycorrhizal fungi (Wright et al. 2009). The increased diversity of macrofungi observed at Copeland Creek may be partially attributable to increased vigor of host Hemlock trees and the resulting increased nutrient and niche availability. This hypothesis is supported by the lack of significant differences in saprobic macrofungi between study sites, because saprobes are less dependent upon host tree vigor. Because of their important role in plant health, ectomycorrhizal fungi have been the subject of many studies using DNA-based methods in forest ecosystems, but the diversity of saprobic fungi has been given little attention (Lynch and Thorn 2006, Southeastern Naturalist 205 R. Baird, et al. 2014 Vol. 13, Special Issue 6 Porter et al. 2008). Our assessment of saprobic fungi richness and diversity was based on only 60 basidiomata collected over 3 years; these collections represented 45 taxa. In fact, we observed no significant correlations between saprobe occurrences and any measured parameter with the exception of tree health. Further, a greater proportion of saprobic macrofungi was found at both sites compared with ectomycrrhizal fungi (26.9% and 15.2%, respectively; Baird et al. 2013). Perhaps the most critical factor affecting both Hemlock tree vigor and nutrient availability, and thus fungal community structure, is canopy defoliation by HWA. As previously discussed, we selected study sites based on locations with limited impact by HWA at the beginning of the study. However, as with much of the southern Appalachians, both sites became increasingly infested during the 3-year study. At Copeland Creek, defoliation levels averaged 36.0% at the onset of the study, and 48.6% at the conclusion of the study. Defoliation by HWA was more severe at the Gabes Mountain site, with initial and final defoliation at 32.0% and 69.8% respectively, and with 15% tree mortality observed by the end of the third year. Numerous studies have shown that mycorrhizal fungal communities may be altered by defoliation of host trees (Rossow et al. 1997, Saikkonen et al. 1999). Ectomycorrhizal fungi use as much as 25% of host carbohydrate production (Hobbie 2006). Decreased carbohydrate supply to roots has been shown to reduce mycorrhizal root-tip abundance (Lewis and Strain 1994) and alter fungal and plant community compositions (Lewis et al. 2008, Rygiewicz et al. 2000). Pearson correlation results showed richness, diversity, and evenness of total, ectomycorrhizal, and saprobic macrofungi were closely correlated with woody plant abundance, richness, diversity, and evenness. Other studies have shown that woody vegetation can have an impact on fungal communities. A recent study by Burke et al. (2009) indicated that plant distribution was strongly correlated with root-associated fungi in a mature Fagus spp. (beech)-Acer spp. (maple) forest, and that both woody and herbaceous plants affected tree-root fungal communities. Many ectomycorrhizal fungi can colonize a wide range of plant species, and can host diverse tree species (Trappe 1977), especially those in the families Russulaceae and Thelephoraceae (Horton and Bruns 2001, Izzo et al. 2005, Tedersoo et al. 2008). However, some ectomycorrhizal fungi are specific to certain tree species (Dickie et al. 2009). Herbaceous plants could also affect the distribution of root-associated fungi by influencing the nutrient cycling, including N, P, and K within forests (Gilliam 2007). Another study by DeBellis et al. (2006) indicated that the distributions of ectomycorrhizal fungi are influenced by the relative proportions of host-tree species within the community. These influences have been attributed to patterns of ectomycorrhizal fungal host preference (Massicotte et al. 1999), brought about by differences in root and litter inputs (Tuininga and Dighton 2004), or to differences in patterns of belowground resource allocation (Bauhus and Messier 1999). Imidacloprid has been effective and extensively used in the GRSM for HWA control since 2006. Through January 2008, over 75,000 Hemlock trees covering approximately 890 ha have been systemically treated with imidacloprid (Johnson Southeastern Naturalist R. Baird, et al. 2014 206 Vol. 13, Special Issue 6 et al. 2008). This study provides the first known report on the in situ effects of imidacloprid soil-drenching on Hemlock-associated fungal communities. Imidacloprid is a nitrogen-containing nicotinoid chemical (Soloway et al. 1978). As mentioned previously, soil nitrogen can significantly alter the distribution and community structure of ectomycorrhizal fungi (Burke et al. 2009), and the response of macrofungi to elevated nitrogen availability has received much attention (Kranabetter et al. 2009). Evidence from studies using nitrogen fertilizer applications suggest immediate declines in ectomycorrhizal sporophore production and species richness with increased nitrogen availability, but with some positive responses in abundance for a subset of nitrophilic species (Avis et al. 2003, Edwards et al. 2004). In a study conducted in Hemlock stands, Kernaghan et al. (1995) noted a trend toward reduction in the proportion of Cenococcum (Ascomycota) and other mycorrhizal types lacking a mantle when granulated urea (46% N) was applied. Further, a study by Singh and Singh (2005) showed that imidacloprid directly suppressed fungal growth in microcosms. In the current study, mineralization or decomposition of imidacloprid by soil microbes leading to increased soil nitrogen and other chemical deposition mechanisms could possibly contribute to occurrences of some ectomycorrhizal fungi, thereby decreasing species evenness. Imidacloprid applications may directly or indirectly impact fungivorous insect populations in the soil thereby reducing the consumption of mycorrhizal fungi and indirectly disrupting symbiosis and the carbohydrate supply (Gehring and Whitham 1994). The observed effect of imidacloprid could also be an indirect result of the removal of fungivorous insects from the root zone. Soil arthropods play a major role in the regulation of below-ground detrital food webs (Moore et al. 1988). In particular, many Collembolans are fungal species-specific in their feeding choices or feed selectively on multiple species of ectomycorrhizal fungi (Bills et al. 2004). Selective grazing by Collembolans has been shown to alter the outcome of competition between fungal species (Newell 1984b, Parkinson et al. 1979). Arthropods have also been shown to facillitate the dispersal of fungi, especially ectomycorrhizal fungi, and thus aid in natural reforestation (Lilleskov and Bruns 2005). Recent studies in the GRSM have shown Collembolans to be highly susceptible to imidacloprid soil-drenching (Reynolds 2008). It is possible that the removal of these fungivorous insects allowed for increased sporophore production in certain dominant species resulting in the observed decrease in species evenness, but this is only conjecture. Regardless of the mechanism, results of this study indicate that imidacloprid has no deleterious effects on fungal community structure in terms of species richness and diversity. Treatments may in fact enhance the stress tolerance and stability of fungal communities by preferentially enhancing dominant Hemlock-associated taxa indirectly as an effect of Collembolans loss. Soil drenches of imidicloprid applied to the litter and topsoil layers around Hemlock trees could cause ectomycorrhizal fungi to be in more direct contact with the chemical than saprobic species. However, differences in impacts of these two groups may be minimal because both groups of fungi occur in and on soil and litter. For while decay fungal biomass occurs on branches and larger wood above the Southeastern Naturalist 207 R. Baird, et al. 2014 Vol. 13, Special Issue 6 ground or on standing dead trees because sporophores are associated with those plant tissues, in most cases, those same fungi can be tracked back to soil surface. Consequently, impacts by the nitrogen release following chemical degradation and negative losses to the arthropod fungivous community would be similar for both groups of fungi, thus affecting their dissemination and life cycles. Macrofungal communities were unaffected by imidicloprid treatments, as measured by species richness and diversity. However, although community structure was unaffected by treatment, species evenness was reduced. This finding indicates that some dominant ectomycorrhizal taxa increased in relative abundance. The long-term effects of imidacloprid on Hemlock-associated ectomycorrhizal communities require further study. However, our findings suggest that any potentially negative effects of imidacloprid on fungal communities in Hemlock forests are far outweighed by the potentially devastating effects of hemlock defoliation and death due to HWA. In conclusion, macrofungal diversity was lower at the high-elevation virgin Hemlock forest site than in the lower second-growth forest site. Macrofungal communities were influenced by associated vegetation, tree health, and microsite variation and not necessarily elevation alone. The extremely high diversity of beneficial fungi in the Southern Appalachian Mountains has important implications for conservation of biodiversity in Hemlock forests. Further research is needed to determine the ecological requirements and roles of rare, difficult-to-identify, and host-specific fungi. Host-tree characteristics such as size, age, and vigor may have worked in combination with environmental factors related to elevation to affect ectomycorrhizal diversity. However, repeated sampling of multiple sites at high and low elevations in the Appalachian Mountains would be necessary to confirm the generality of these patterns and to more fully test relationships between the environment and mycorrhizal communities (Walker et al. 2005). The results of this study will help to illuminate the ecology of Hemlock-associated fungal communities and illustrate their sensitivity to environmental changes. In addition, we evaluated the effects of imidacloprid application, and found no significant effect on the structure of fungal communities. While community structure was unaffected by treatments, community composition of ectomycorrhizal macrofungi was altered, as evidenced by reduced species evenness. This result indicates that certain dominant ectomycorrhizal taxa increased in relative abundance. These findings suggest that use of imidacloprid to suppress HWA is beneficial to Hemlock-associated fungal communities to the extent that canopy defoliation is avoided and host-tree health is maintained. Acknowledgments Appreciation is extended to Highlands Biological Station for financial support as grants-in-aid during 2007 and 2008. Thanks are due to GSRM (National Park Service, US Department of the Interior) for logistical and technical support of the project, David Pratt for use of laboratory and housing facilities at the University of Tennessee Field Station, and Mississippi State University (MAFES publication number 12343) for use of laboratory facilities and for supplies not covered by grants. Southeastern Naturalist R. Baird, et al. 2014 208 Vol. 13, Special Issue 6 Literature Cited Avis, P.G., D.J. McLaughlin, B.C. Dentinger, and P.B. Reich. 2003. Long-term increase in nitrogen supply alters above- and below-ground ectomycorrhizal communities and increases dominance of Russula spp. in a temperate oak savanna. New Phytologist 160:239–253. Baird, R.E., C.E. Watson, and S. Woolfolk. 2007. Microfungi from bark of healthy and damaged American Beech, Fraser Fir, and Eastern Hemlock trees during an All Taxa Biodiversity Inventory in forests of Great Smoky Mountains National Park. Southeastern Naturalist 6:67–82. Baird, R., L. Wallace, G. Baker, and M. Scruggs. 2013. Stipitate hydnoid fungi of the temperate southeastern United States. Fungal Diversity 62:41–114 Baird, R.E., E. Stokes, A. Wood-Jones, C. Watson, M. Alexander, G. Taylor, K. Johnson, P. Threadgill, and S. Diehl. 2014. A molecular clone and culture inventory of the root fungal community associated with Eastern Hemlock in Great Smoky Mountains National Park. Southeastern Naturalist Special Issue 6:219–237. Baker, F.S. 1949. A revised tolerance table. Journal of Forestry 47:179–181. Bauhaus, J., and C. Messier. 1999. Soil-exploitation strategies of fine roots in different tree species of the southern boreal forest of eastern Canada. Canadian Journal of Forest Research 29:260–273. Bergemann, S.E., N.C. Kordesch, W. Van Sant-Glass, M. Garbelotto, and T.A. Metz. 2013. Implications of Tan Oak decline in forests impacted by Phytophthora ramorum: Girdling decreases the soil hyphal abundance of ectomycorrhizal fungi associated with Notholithocarpus densiflorus. Madrono 60:95–106. Bills, G.F., M. Christensen, M. Powell, and G. Thorn. 2004. Saprophytic soil fungi. Pp. 303–316, In G.M. Mueller, G.F. Bills, and M.F. Foster (Eds.). Biodiversity of Fungi: Inventory and Monitoring Methods. Elsevier Academic Press, San Diego, CA. Burke, D.J., J.C. López-Gutierrez, K.A.S. Memo, and C.R. Chan. 2009. Vegetation and soil environment influence the spatial distribution of root-associated fungi in a mature beech-maple forest. Applied Environmental Microbiology 75:7639–7648. Cowles, R.S., C. Cheah, and M.E. Montgomery. 2004. Effect of imidacloprid application technique on Hemlock Woolly Adelgid mortality and insecticide concentration in sap. In B. Onken and D. Souto. (Eds.). Hemlock Woolly Adelgid newsletter. USDA Forest Service, Northeastern Area State and Private Forestry, Forest Health Protection. Issue 6. Available online at http://www.fs.fed.us/na/morgantown/fhp/hwa/news6/news6.html. Accessed January 2004. De Bellis, T., G. Kernaghan, R. Bradley, and P. Widden. 2006. Relationships between stand composition and ectomycorrhizal community structure in boreal mixed-wood forests. Microbial Ecology 99:114–126. DeGraaf, R.M., M. Yamasaki, W.B. Leak, and J.W. Lanier. 1992. New England wildlife: Management of forested habitats. General Technical Report NE-144. US Department of Agriculture, Forest Service, Northeastern Research Station Radnor [now Newtown Square], PA. 271 pp. Dickie, I.A., B. Dentinger, P. Avis, and D.P. McLaughlin. 2009. Ectomycorrhizal fungal communities of oak savanna are distinct from forest communities. Mycologia 101:473–483. Doccola, J.J., P.M. Wild, I. Ramasamy, P. Castillo, and C. Taylor. 2003. Efficacy of Arborjet VIPER microinjections in the management of Hemlock Woolly Adelgid. Journal of Arboriculture 29:327–330. Southeastern Naturalist 209 R. Baird, et al. 2014 Vol. 13, Special Issue 6 Drobner, U., J. Biddy, B. Smith, and J.B. Wilson. 1998. The relation between community biomass and evenness: What does community theory predict and can these predictions be tested? Oikos 82:295–302. Durall, D.M., M.D. Jones, E.F. Wright, P. Kroeger, and K.D. Coates. 1999. Species richness of ectomycorrhizal fungi in cutblocks of different sizes in the interior cedar–hemlock forests of northwestern British Columbia: Sporocarps and ectomycorrhizae. Canadian Journal of Forest Research 29:1322–1333. Dyke, F.V. 2003. Conservation Biology: Foundations, Concepts, Applications. McGraw- Hill, New York, NY. 414 pp. Edwards, I.P., J.L. Cripliver, A.R. Gillespie, K.H. Johnsen, J.M. Scholler, and R.F. Turco. 2004. Nitrogen availability alters macrofungal basidiomycete community structure in optimally fertilized Loblolly Pine forests. New Phytologist 162:755–770. Evans, R.A. 2002. An ecosystem unraveling? Pp. 23–33, In B. Onken, R. Reardon, and J. Lashomb (Eds.) Proceedings, Hemlock Woolly Adelgid in the Eastern United States Symposium. 5–7 February 2002. Rutgers University, East Brunswick, NJ. Gehring, C.A., and T.G. Whitham. 1994. Comparisons of ectomycorrhizae on Pinyon Pine (Pinus edulis, Pinaceae) across extremes of soil type and herbivory. American Journal of Botany 81:1509–1516. Gilliam, F.S. 2007. The ecological significance of the herbaceous layer in temperate forest ecosystems. Bioscience 57:845–858. Godman, R.M., and K. Lancaster. 1990. Tsuga canadensis (L.) Carr. Eastern Hemlock. Pp. 604–612, In R.M. Burns and B.H. Honkala (Eds.). Silvics of North America: Vol. 1. Conifers. Agriculture Handbook 654. USDA Forest Service. Washington, DC. Graham, S.A. 1954. Scoring tolerance of forest trees. University of Michigan Research Note 4. Ann Arbor, MI. 2 pp. Grime, J.P., J.M.L. Mackey, S.H. Hillier, and D.J. Read. 1987. Floristic diversity in a model system using experimental microcosms. Nature 328:420–422. Hanlon, R.D., and J.M. Anderson. 1979. The effects of Collembola grazing on microbial activity in decomposing leaf litter. Oecologia 38:93–99. Harvey, A.E., M.F. Jurgensen, and M.J. Larsen. 1978. Seasonal distribution of ectomycorrhizae in a mature Douglas-fir/larch soil in western Montana. Forest Science 24:203–208. Hobbie, E.A. 2006. Carbon allocation to ectomycorrhizal fungi correlates with belowground allocation in culture studies. Ecology 87:563–569. Horton, T.R., and T.D. Bruns. 2001. The molecular revolution in ecomycorrhizal ecology: Peeking into the black-box. Molecular Ecology 10:1855–1871. Ingham, E.R., and W. Thies. 1996. Response of soil food-web organisms in the first year following clear-cutting and application of chloropicrin to control laminated root rot. Applied Soil Ecology 3:35–47. Izzo, A., J. Agbowo, and T.D. Bruns. 2005. Detection of plot-level changes in ectomycorrhizal communities across years in old-growth mixed-conifer forest. New Phytologist 166:619–630. Johnson, K., T. Remaley, and G. Taylor. 2008. Managing Hemlock Woolly Adelgid at Great Smoky Mountains National Park: Situation and response. Pp. 62–69, In B. Onken and R. Reardon (Eds.). Proceedings of the Fourth Symposium on Hemlock Woolly Adelgid in the eastern United States. FHTET-2008-01, USDA Forest Service, Forest Health Technology Enterprise TEAM. Morgantown, WV. Kaiser, C., M. Koranda, B. Kitzler, L. Fuchslueger, J. Schnecker, P.Chweiger, F. Rasche, S. Zechmeister-Boltenstern, A. Sessitsch, and A. Richter. 2010. Belowground carbon allocation by trees drives seasonal patterns of extracellular enzyme activities by altering microbial community composition in a beech forest soil. New Phytologist 187:843–858. Southeastern Naturalist R. Baird, et al. 2014 210 Vol. 13, Special Issue 6 Kernaghan, G., S.M. Berch, and R. Carter. 1995. Effect of urea fertilization on ectomycorrhizae of 20-year-old Tsuga heterophylla. Canadian Journal of Forest Research 25(6):891–901. Kornerup, A., and J.H. Wanscher. 1978. Methuen Handbook of Colour. Eyre Methuen, London, UK. 252 pp. Kranabetter, S.M., D.M. Durall, and D.H. Mackenzie. 2009. Diversity and species distribution of ectomycorrhizal fungi along productivity gradients of a southern boreal forest. Mycorrhiza 19:99–111. Krebs, C.J. 1989. Ecological Methodology. Harper and Ross Publications, New York, NY. 654 pp. Largent, D.L., D. Johnson, and R. Watling. 1977. How to Identify Mushrooms to Genus III: Microscopic Features. Mad River Press, Eureka, CA. 148 pp. Lewis, J.D., and B.R. Strain. 1994. Effects of elevated CO2 on mycorrhizal colonization of Loblolly Pine (Pinus taeda L.) seedlings. Plant Soil 165:81–88. Lewis, J.D., J. Licitra, A.R. Tuininga, A. Sirulnik, G.D. Turner, and J. Johnson. 2008. Oak seedling growth and ectomycorrhizal colonization are less in Eastern Hemlock stands infested with Hemlock Woolly Adelgid than in adjacent oak stands. Tree Physiology 28:629–636. Lilleskov, E.A., and T.D. Bruns. 2005. Spore dispersal of a resupinate ectomycorrhizal fungus, Tomentella sublilacina, via soil food webs. Mycologia 97:762–769. Luoma, D.L., J.L. Eberhart, and M.P. Amaranthus. 1997. Biodiversity of ectomycorrhizal types from southwest Oregon. Pp. 249–253, In T.N. Kaye, A. Liston, D.L. Luoma, R.J. Meinke, and M.V. Wilson (Eds.). Management of Native Plants and Fungi. Native Plant Society of Oregon, Corvallis, OR. Lynch, M.D.J., and R.G. Thorn. 2006. Diversity of basidiomycetes in Michigan agricultural soils. Applied Environmental Microbiology 72:7050–7056. Magurran, A.E. 2004. Measuring Biological Diversity. Blackwell Publishing, Oxford, UK. Martin, M.M. 1987. Invertebrate-microbial Interactions: Ingested Fungal Enzymes in Arthropod Biology. Comstock Publishing Association, Ithaca, NY. 148 pp. Massicotte, H.B., R. Molina, L.E. Tackaberry, J.E. Smith, and M.P. Amaranthus. 1999. Diversity and host specificity of ectomycorrhizal fungi retrieved from three adjacent forest sites by five host species. Canadian Journal of Botany 77:1053–1 076. McLenon-Porter, T.M. 2008. Above- and below-ground fungal diversity in a hemlockdominated forest plot in southern Ontario and the phylogenetic placement of a new Ascomycota subphylum. Ph.D. Dissertation. University of Toronto, ON, Canada. 235 pp. Moore, J.C., D.E. Walter, and H.W. Hunt. 1988. Arthropod regulation of micro-and mesobiota in below-ground detrital food webs. Annual Review of Entomology 33:419–439. Mullin, C.A., M.C. Saunders, T.W. Leslie, D.J. Biddinger, and S.J. Fleischer. 2005. Toxic and behavioral effects to Carabidae of seed treatments used on Cry3Bb1- and Cry1Ab/cprotected corn. Environmental Entomology 34:1626–1636. Newell, K. 1984a. Interaction between two decomposer basidiomycetes and a collembolan under Sitka Spruce: Distribution, abundance, and selective grazing (Mycena galopus, Marasmius androsaceus, Picea sitchensis). Soil Biology and Biochemistry 16:227–233. Newell, K. 1984b. Interaction between two decomposer basidiomycetes and a collembolan under Sitka Spruce: Grazing and its potential effects on fungal distribution and litter decomposition (Mycena galopus, Marasmius androsaceus, Onychiurus latus). Soil Biology and Biochemistry 16:235–239. Parkinson, D., S. Visser, and J.B. Whittaker. 1979. Effects of collembolan grazing on fungal colonization of leaf litter. Soil Biology and Biochemistry 11:529–535. Southeastern Naturalist 211 R. Baird, et al. 2014 Vol. 13, Special Issue 6 Pena, R., C. Offermann, J. Simon, P.S. Naumann, A. Gessler, J. Holst, M. Dannenmann, H. Mayer, I. Kogel-Knabner, H. Rennenberg, and A. Polle. 2010. Girdling affects ectomycorrhizal fungal (EMF) diversity and reveals functional differences in EMF community composition in a beech forest. Applied Environmental Microbiology 76:1831–1841. Porter, T.M., C.W. Schadt, L. Rizui, A.P. Martin, S.K. Schmidt, L. Scott-Denton, R. Vilgalys, and J.M. Moncalvo. 2008. Widespread occurrence and phylogenetic placement of a soil clone group adds a prominent new branch to the fungal tree of life. Molecular Phylogenetic Evolution 46:635–644. Price, P.W. 1997. Insect Ecology, 3rd Edition. John Wiley and Sons, New York, NY. 874 pp. Rastin, N., G. Schlechte, A. Hüttermann, and K. Rosenplänter. 1990. Seasonal fluctuation of some biological and biochemical soil factors and their dependence on certain soil factors on the upper and lower slope of a spruce forest. Soil Biology and Biochemistry 22:1049–1061. Read, D.J. 1991. Mycorrhizas in ecosystems. Experientia 47:376–39. Reynolds, W.N. 2008. Imidacloprid insecticide treatments for Hemlock Woolly Adelgid, Adelges tsugae Annand (Hemiptera: Adelgidae), affect a non-target soil arthropod community surrounding Eastern Hemlock, Tsuga canadensis (L.) Carriere. M.Sc. Thesis. The University of Tennessee, Knoxville, TN. 125 pp. Rossow, L.J., J.P. Bryant, and K. Kielland. 1997. Effects of above ground browsing by mammals on mycorrhizal infection in an early successional tiaga ecosystem. Oecologia 110:94–98. Rygiewicz, P.T., K.J. Martin, and A.R. Tuininga. 2000. Morphotype community structure of ectomycorrhizas on Douglas-fir (Pseudotsuga menziesii Mirb. Franco) seedlings grown under elevated atmospheric CO2 and temperature. Oecologia 124:299–308. Saikkonen, K., U. Ahonen-Jonnarth, A.M. Markkola, M. Helander, J. Tuomi, M. Roitto, and H. Ranta. 1999. Defoliation and mycorrhizal symbiosis: A functional balance between carbon sources and below-ground sinks. Ecological Letters 2:19–26. SAS Institute. 1999. SAS/STAT software changes and enhancements through release V8 SAS Institute, Cary, NC. Schomaker, M.E., S.J. Zarnoch, W.A. Bechtold, D.J. Latelle, W.G. Burkman, and S.M. Cox. 2007. Crown-condition classification: A guide to data collection and analysis. General Technical Report SRS-102. US Department of Agriculture, Forest Service, Southern Research Station, Asheville, NC. 78 pp. Shaw, P. 1992. Fungi, fungivores, and fungal food webs. Pp. 295–310, In G.C. Carroll and D.T. Wicklow (Eds.). The Fungal Community: Its Organization and Role in the Ecosystem. Marcel Dekker, New York, NY. Simard, S.W. 1997. Intensive management of young mixed forests: Effects on forest health. Pp. 48–54, In R. Sturrock (Ed.). Proceedings of the 45th Western International Forest Disease Work Conference, 15–19 September 1997. Prince George, BC, Canada. Singh, J., and D.K. Singh. 2005. Bacterial, azotobacter, actinomycetes, and fungal population in soil after diazinon, imidacloprid, and lindane treatments in Groundnut (Arachis hypogaea L.) fields. Journal of Environmental Science and Health. Part. B: Pesticides, Food Contaminants, and Agricultural Wastes 40:785–800. Sirulnik, A., J. Lewis, A. Tuininga, J. Johnson, and C. Louis. 2005. Soil conditions, host community, and infestations of the Hemlock Woolly Adelgid (Adelges tsugae) affects ectomycorrhizal diversity in eastern temperate forests. Ecological Society of America (ESA) 90th Annual Meeting, 4–12 August 2005. Montreal, QC, Canada. Smith, S.E., and D.J. Read. 1997. Mycorrhizal Symbiosis. Academic Press, San Diego, CA. 605 pp. Southeastern Naturalist R. Baird, et al. 2014 212 Vol. 13, Special Issue 6 Smith, S.E., and D.J. Read. 2008. Mycorrhizal Symbiosis, 3rd Edition, Academic Press, London, UK. 800 pp. Snyder, C., J. Young, D. Smith, D. Lemarie, R. Ross, and R. Bennett. 2004. Stream ecology linked to Eastern Hemlock decline in Delaware Water Gap National Recreation Area. US Geological Survey, Kearneysville, WV. Available online at http://www.lsc.usgs.gov/ aeb/2048-03/dewa.asp. Accessed 2013. Soloway, S.B., A.C. Henry, W.D. Kollmeyer, W.M. Padgett, J.E. Powell, S.A. Roman, C.H. Tieman, R.A. Corey, and C.A. Horne. 1978. Nitromethylene Insecticides. Pp. 206–217, In H. Geissbuhler, G.T. Brooks, and C. Kearney (Eds.). Advances in Pesticide Science, Part 2. Pergamon Press, Oxford, UK. Stephenson, S.L. 1989. Distribution and ecology of myxomycetes in temperate forests II. Patterns of occurrence on bark surface of living trees, leaf litter, and dung. Mycologia 81:608–621. Stephenson, S.L., M. Schnittler, and C. Lado. 2004. Ecological characterization of a tropical myxomycete assemblage—Maquipucuna Cloud Forest Reserve, Ecuador. Mycologia 96:488–497. Steward, V.B., and T.A. Horner. 1994. Control of Hemlock Woolly Adelgid using soil injections of systemic insecticides. Journal of Arboriculture 20:287–288. Swaty, R.L., C.A. Gehring, M. Van Ert, T.C. Theimer, P. Keim, and T.G. Whitham. 1998. Temporal variation in temperature and rainfall differentials affects ectomycorrhizal colonization at two contrasting sites. New Phytologist 139:733–739. Tedersoo, L., T. Jairus, B.M. Horton, K. Abarenkov, T. Suvi, I. Saar, and U. Kõljalg. 2008. Strong host preference of ectomycorrhizal fungi in Tasmanian wet sclerophyll forest revealed by DNA barcoding and taxon-specific primers. 180:479–490. Trappe, J.M. 1977. Selection of fungi for ectomycorrhizal inoculation in nurseries. Annual Review of Phytopathology 15:203–222. Tubbs, C.H. 1977. Manager’s handbook for northern hardwoods in the North Central States. USDA Forest Service, General Technical Report NC-39. North Central Forest Experiment Station, St. Paul, MN. 29 pp. Tuininga, A.R., and J. Dighton. 2004. Changes in ectomycorrhizal communities and nutrient availability following prescribed burning in two upland pine-oak forests in the New Jersey Pine Barrens. Canadian Journal of Forest Research 34:1755–1765. Van der Drift, J., and E. Jansen. 1977. Grazing of springtails on hyphal mats and its influence on fungal growth and respiration. Pp. 203–209, In U. Lohm and T. Persson (Eds). Soil Organisms as Components of Ecosystems. Ecological Bulletin 25. van der Heijden, M.G.A., J.N. Klironomos, M. Ursic, P. Moutoglis, R. Streitwolf-Engel, T. Boiler, A. Wiemken, and I.R. Sanders. 1998. Mycorrhizal fungal diversity determines plant diversity, ecosystem variability, and productivity. Nature 396:69–72. Visser, S. 1995. Ectomycorrhizal fungal succession in Jack Pine stands following wildfire. New Phytologist 129:389–401. Vogt, K.A., and R.L. Edmonds. 1980. Patterns of nutrient concentration in basidiocarps in western Washington. Canadian Journal of Botany 58:694–698. Walker, J.F., O.K. Miller, Jr., and J.L. Horton. 2005. Hyperdiversity of ectomycorrhizal fungus assemblages on oak seedlings in mixed forests in the southern Appalachian Mountains. Molecular Ecology 14:829–838. Walker, J.F., O.K. Miller, Jr., and J.L. Horton. 2008. Seasonal dynamics of ectomycorrhizal fungus assemblages on oak seedlings in the southeastern Appalachian Mountains. Mycorrhiza 18:123–132. Southeastern Naturalist 213 R. Baird, et al. 2014 Vol. 13, Special Issue 6 Ward, J.S., M.E. Montgomery, C. Cheah, B.P. Onken, and R.S. Cowles. 2004. Eastern Hemlock forests: Guidelines to minimize the impacts of Hemlock Woolly Adelgid. USDA Forest Service, NA-TP-03-04, Northeastern Area State and Private Forestry, Morgantown, WV. 27pp. Webb, R.E., R.J. Frank, and M.J. Raupp. 2003. Eastern Hemlock recovery from Hemlock Wooly Adelgid damage following imidacloprid therapy. Journal of Aboriculture 29:298–302. Wright, S.H.A., S.M. Berch, and M.L. Berbee. 2009. The effect of fertilization on the below- ground diversity and community composition of ectomycorrhizal fungi associated with Western Hemlock (Tsuga heterophylla). Mycorrhiza 19:267–276. Southeastern Naturalist R. Baird, et al. 2014 214 Vol. 13, Special Issue 6 Appendix A. Frequency and relative abundance of macrofungi associated with Eastern Hemlock, Great Smoky Mountains National Park, September 2006–2008. Abundance values are based on mean percent relative abundance of macrofungi taxa from two sites: CC = Copeland Creek (487 m elevation), GM = Gabes Mountain (1158 m elevation) and 3 imidacloprid application rates/site: full = 11.8 ml/cm dbh, half = 5.9 ml/cm dbh, and control = no imidacloprid application. n = 20 subplots per location (40 total). Frequency indicates mean percent occurrence of macrofungi species per tree, n = 20 subplots per location. Overall abundance values are based on mean percent relative abundance of total macrofungi taxa collected within Eastern Hemlock study sites, n = 20 subplots per location (6 control, 7 Full, 7 half). * indicates species previously reported in the park from University of Tennesse Herbarium listings online (http://tenn.bio.utk.edu/fungus/database/ fungus-browse-results.asp?; GSMNP=GRSM); ** indicates species previously reported in the park from a report of collections by Andrew Miller of the University of Illinois (http://usmo4.discoverlife.org/mp/20p?res=640&see = I_ANM78). EM = ectomycorrhizal; W = saprobic or wood decay; and P = parasitic species. Abundance by Site Rate Overall Fungal taxon Authority Frequency CC GM Full Half Control abundance Ascomycota Galiella rufa* DW, H-111 (Schwein.) Nannf. & Korf 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Hypoxylon howeanum** W, H-121 Peck 0.0 0.0 2.0 0.0 0.0 0.0 <1.0 Leotia lubrica* W, H-122 (Scop.) Pers. 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Spathularia velutipes* W, H 123, 126, 151 Cooke & Farl. 4.9 2.0 0.0 0.0 2.6 0.0 1.6 Xylaria sp.* W, H-133 Hill ex Schrank 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Basidiomycota Amanita bisporigera* EM, H-3 G.F. Atk. 4.9 1.5 0.0 0.0 1.3 2.8 1.2 Amanita brunnescens* EM, H-37, 61, 62 G.F. Atk. 4.9 <1.0 0.0 0.0 2.6 0.0 <1.0 Amanita cinereopannosa EM H-101b Bas 2.4 2.5 0.0 6.8 0.0 0.0 2.0 Amanita citrina* EM, H-28 (Schaeff.) Pers. 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Amanita citrina var. lavendula* EM, H-5 (Coker) Sartory & Maire 17.1 6.9 0.0 9.5 5.3 5.6 5.5 Amanita flavoconia* EM, H-65 G.F. Atk. 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Amanita frostiana* EM, H-7 (Peck) Sacc. 4.9 <1.0 0.0 1.4 1.3 0.0 <1.0 Amanita gemmata* EM, H-67, 101 (Fr.) Bertill. 1.0 0.0 1.0 <1.0 1.0 0.0 <1.0 Amanita onusta* EM, H-82 (Howe) Sacc. 2.4 0.0 2.0 0.0 1.3 0.0 <1.0 Southeastern Naturalist 215 R. Baird, et al. 2014 Vol. 13, Special Issue 6 Abundance by Site Rate Overall Fungal taxon Authority Frequency CC GM Full Half Control abundance Amanita rubescens* EM, H- Pers. 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Amanita russuloides* EM, H-64b (Peck) Sacc. 9.8 2.0 0.0 4.1 0.0 2.8 1.6 Amanita virosa* EM H-94 (Fr.) Bertill. 2.0 <1.0 0.0 0.0 1.3 1.2 <1.0 Ampulloclitocybe clavipes* W, H-162 (Pers.) Redhead, Lutzoni, 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Moncalvo & Vilgalys Armillaria mellea* P, H-124, 142 (Vahl) P. Kumm. 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Austroboletus gracilis* EM, H-57 (Peck) Wolfe 14.6 5.4 0.0 8.1 5.3 2.8 4.3 Boletellus russellii* EM, H-102 (Frost) E.J. Gilbert 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Boletus spadiceus var. gracilis* EM, H-38 A.H. Sm. & Thiers 2.4 <1.0 0.0 0.0 0.0 2.8 <1.0 Cantharellus minor* EM, H-103 Peck 2.4 <1.0 0.0 0.0 0.0 2.8 <1.0 Clavulina cristata* EM, H-60 (Holmsk.) J. Schröt. 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Clitocybe sp.* W, H-125 2.4 <1.0 2.0 0.0 1.3 0.0 <1.0 Clitocybe hygrophoroides W, H-189 H.E. Bigelow 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Clitocybe trullaeformis* W, H-116 (Fr.) P. Karst. 2.4 <1.0 0.0 0.0 0.0 2.8 <1.0 Coltricia montagnei* W, H-185 (Fr.) Murrill 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Cortinarius croceofolius EM, H-29 Peck 4.9 1.5 0.0 0.0 2.6 2.8 1.2 Cortinarius sp.* EM, H-39, 69 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Craterellus fallax* EM, H-197 A.H. Sm. 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Crepidotus albatus* W, H-132 Hesler & A.H. Sm. 0.0 0.0 3.9 0.0 0.0 0.0 <1.0 Crepidotus appalachiensis* W, H-192 Hesler & A.H. Sm. 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Crepidotus crocophyllus* W, H-148 (Berk.) Sacc. 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Crepidotus malachius* W, H-120 Sacc. 0.0 <1.0 2.0 0.0 0.0 0.0 <1.0 Crepidotus sp.* W, H-168 2.4 0.0 2.0 0.0 0.0 2.8 <1.0 Dacrymyces sp.* W, H-190 2.4 <1.0 0.0 0.0 0.0 2.8 <1.0 Entoloma incanum* W, H-152 (Fr.) Hesler 4.9 <1.0 3.9 1.4 1.3 0.0 1.2 Fomitopsis cajanderi* W, H-157, 164 (P. Karst.) Kotl. and Pouzar 7.3 2.0 0.0 0.0 2.6 2.8 1.6 Ganoderma applanatum* W, P, H-196 (Pers.) Pat. 2.4 0.0 2.0 1.4 0.0 0.0 <1.0 Ganoderma tsugae* W, P, H-197 Murrill 0.0 0.0 2.0 0.0 0.0 0.0 <1.0 Southeastern Naturalist R. Baird, et al. 2014 216 Vol. 13, Special Issue 6 Abundance by Site Rate Overall Fungal taxon Authority Frequency CC GM Full Half Control abundance Gomphus clavatus* EM, 194 (Pers.) Gray 2.4 <1.0 0.0 0.0 0.0 2.8 <1.0 Gymnopus dichrous W, H-180 (Berk. & M.A. Curtis) Halling 0.0 0.0 2.0 0.0 0.0 0.0 <1.0 Gymnopus dryophilus* W, H-138, 141, 155 (Bull.) Murrill 9.8 2.5 3.9 1.4 3.9 0.0 2.7 Hemistropharia albocrenulata* W, H-181 (Peck) Jacobsson & E. Larss. 2.4 0.0 2.0 0.0 1.3 0.0 <1.0 Hohenbuehelia mastrucata* W, H-183 (Fr.) Singer 0.0 0.0 2.0 0.0 0.0 0.0 <1.0 Hydnellum spongiosipes* EM, H-95 (Peck) Pouzar 4.9 1.5 0.0 2.7 1.3 0.0 1.2 Hydnum albidum* EM, H-27b Peck 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Hygrocybe appalachiensis* EM, H-156 (Hesler & A.H. Sm.) Kronaw. 4.9 0.0 3.9 1.4 0.0 2.8 <1.0 Hygrocybe miniata* EM, H-129 (Fr.) P. Kumm. 2.4 0.0 3.9 2.7 0.0 0.0 <1.0 Hygrocybe psittacina* EM, H=131 (Schaeff.) P. Kumm. 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Hygrophorus pudorinus* EM, H-83 (Fr.) Fr. 2.4 0.0 2.0 1.4 0.0 0.0 <1.0 Laccaria laccata* EM, H-9 (Scop.) Cooke 9.8 3.4 0.0 1.4 0.0 13.9 2.7 Laccaria ochropurpurea* EM, H-96 (Berk.) Peck 2.4 <1.0 0.0 0.0 0.0 2.8 <1.0 Lactarius aquifluus EM, H-11 Peck 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Lactarius deceptivus* EM, H-88 Peck 4.9 <1.0 0.0 0.0 2.6 0.0 <1.0 Lactarius glaucescens* EM, H-40 Crossl. 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Lactarius griseus* EM, H-46 Peck 7.3 <1.0 3.9 2.7 1.3 0.0 1.6 Lactarius sp. 1* EM, H-12 (griseus-like) 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Lactarius mucidus* EM, H-13 Burl. 4.9 <1.0 0.0 0.0 2.6 0.0 <1.0 Lactarius oculatus* EM, H-18 (Peck) Burl. 7.3 <1.0 2.0 1.4 1.3 2.8 <1.0 Lactarius piperatus* EM, H-41 (L.) Pers. 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Lactarius sp. 2 EM, H-107 2.4 <1.0 0.0 0.0 0.0 2.8 <1.0 Lactarius sp. 3 EM, H-106 Pers. 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Lactarius speciosus* EM, H-15 Burl. 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Lactarius subpurpureus* EM, H-16, 59 Peck 4.9 <1.0 0.0 1.4 0.0 2.8 <1.0 Lactarius theiogalus* EM, H-70 (Bull.) Gray 4.9 <1.0 2.0 0.0 2.6 0.0 <1.0 Lactarius vellereus* EM, H-56 (Fr.) Fr. 2.4 <1.0 0.0 0.0 0.0 2.8 <1.0 Southeastern Naturalist 217 R. Baird, et al. 2014 Vol. 13, Special Issue 6 Abundance by Site Rate Overall Fungal taxon Authority Frequency CC GM Full Half Control abundance Laetiporus sulphureus* EM, H-136 (Bull.) Murrill 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Leccinum scabrum* EM, H-60 89 (Bull.) Gray 2.4 <1.0 0.0 0.0 0.0 5.6 <1.0 Lepiota cristata* EM, H-140 Barla 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Lepiota sp.* EM, H-118 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Leptonia incana*EM, H-119 (Fr.) Gillet 2.4 0.0 2.0 0.0 1.3 0.0 <1.0 Leucopholiota decorosa* EM, H-161 (Peck) O.K. Mill., T.J. Volk, 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 & Bessette Lycoperdon perlatum* EM, 153 Pers. 2.4 0.0 2.0 1.4 0.0 0.0 <1.0 Marasmius fulvoferrugineus* EM, H-193 Gilliam 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Marasmius siccus* W, H-112, 191 (Schwein.) Fr. 2.4 1.5 2.0 0.0 1.3 0.0 1.6 Megacollybia platyphylla* W, H-184 (Pers.) Kotl. & Pouzar 4.9 <1.0 5.9 0.0 1.3 2.8 1.6 Meripilus giganteus* W, H-171 (Pers.) P. Karst. 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Mycena sp.* W, H-139 0.0 0.0 2.0 0.0 0.0 0.0 <1.0 Mycorrhaphium adustum* W, H-182 (Schwein.) Maas Geest. 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Paxillus involutus* W, H-186 (Batsch) Fr. 4.9 0.0 3.9 1.4 1.3 0.0 <1.0 Pholiota flavida* W, 159 (Schaeff.) Singer 0.0 0.0 2.0 0.0 0.0 0.0 <1.0 Pleurocybella porrigens* W, H-144 (Pers.) Singer 4.9 0.0 3.9 1.4 0.0 2.8 <1.0 Pleurotus dryinus* W, H-134 (Pers.) P. Kumm. 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Pluteus cervinus* W, H-147, 158 (Schaeff.) P. Kumm. 2.4 <1.0 2.0 0.0 0.0 2.8 <1.0 Polyporus varius* W, H-137, 176 (Pers.) Fr. 4.9 2.5 0.0 4.1 0.0 0.0 2.0 Postia caesius* W, H-129 (Schrad.) P. Karst., 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Psathyrella delineata* W, H-146 (Peck) A. H. Sm. 2.4 <1.0 2.0 1.4 0.0 0.0 1.2 Pseudohydnum gelatinosum* W, H-157, 177 (Scop.) P. Karst. 4.9 0.0 3.9 1.4 1.3 0.0 <1.0 Ramaria formosa* EM, H-42 (Pers.) Quél. 7.3 2.5 0.0 0.0 5.3 2.8 2.0 Ramaria stricta* EM, H-72 (Pers.) Quél. 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Retiboletus ornatipes* EM, H-4 (Peck) Manfr. Binder & 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Bresinsky Russula densifolia* EM, H-34, 73 Secr. ex Gillet 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Southeastern Naturalist R. Baird, et al. 2014 218 Vol. 13, Special Issue 6 Abundance by Site Rate Overall Fungal taxon Authority Frequency CC GM Full Half Control abundance Russula fragrantissima* EM, H-24, 42, 43, Romagn. 22.0 5.4 3.9 9.5 3.9 8.3 5.1 51, 90, 91 Russula krombholzii EM, H-46, 75, 91 Shaffer 7.3 3.9 0.0 1.4 7.9 0.0 3.1 Russula sp. 1 (red) EM, H-76 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Russula sp. 2 (cream, tacky) EM, H-98 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Russula sp. 3 (cream) EM, H-34 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Russula sp. 4 (red) EM, H-108 2.4 <1.0 0.0 0.0 0.0 2.8 <1.0 Russula sp. 5 (red) EM, H-100 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Sarcodon scabrosus* EM, H-21 (Fr.) P. Karst. 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Scleroderma citrinum* EM, H-52, 85 Pers. 4.9 0.0 3.9 1.4 1.3 0.0 <1.0 Strobilomyces confuses* EM, H-77 Singer 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Strobilurus conigenoides* W, H-175 (Ellis) Singer 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Tapinella atrotomentosa* W, H-124, 172 (Batsch) Šutara. 7.3 2.5 0.0 1.4 1.3 2.8 2.0 Trametes versicolor* W, H-150 (L.) Lloyd 2.4 0.0 3.9 0.0 1.3 0.0 <1.0 Trichaptum biforme* W, H-132 (Fr.) Ryvarden 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Tricholoma flavovirens* EM, H-93 S. Lundell 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Tricholoma portentosum* EM, H-93b (Fr.) Quél. 7.3 1.5 0.0 1.4 2.6 0.0 1.2 Tricholoma sp. 1 (striate), EM, H-33 2.4 <1.0 0.0 0.0 1.3 0.0 <1.0 Tricholoma sp. 2 (white), EM, H-33b 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Tricholoma sp. 3 (grey) EM, H-33c 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Tricholomopsis decora* EM, H-167 (Fr.) Singer 0.0 0.0 2.0 0.0 0.0 0.0 <1.0 Tricholomopsis flavissima EM (A.H. Sm.) Singer 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Tricholomopsis formosa* EM, H-149 (Murrill) Singer 0.0 <1.0 0.0 0.0 0.0 0.0 <1.0 Tylopilus violatinctus* EM, H-78 T.J. Baroni & Both 7.3 2.0 0.0 1.4 1.3 2.8 1.6 Tyromyces chioneus* W, H-143, 188 (Fr.) P. Karst. 0.0 1.5 3.9 0.0 0.0 0.0 2.0 Xanthoconium stramineum EM, H-81 (Murrill) Singer 2.4 <1.0 0.0 1.4 0.0 0.0 <1.0 Xeromphalina tenuipes* W, H-135, 187 (Schwein.) A.H. Sm. 2.4 <1.0 2.0 1.4 0.0 0.0 <1.0 Xerula furfuracea* W, H-113, 114 (Peck) Redhead, Ginns, 4.9 1.5 0.0 1.4 1.3 0.0 1.2 & Shoemaker