A Survey of Fungal Community Composition along a Gradient of Recovery on the Mine Sites in the Carolinas

Ruolin Miao

Under the supervision of Dr. Rytas Vilgalys, Department of Biology, Duke University And Dr. Alejandro Rojas, Department of Entomology and Plant Pathology, University of Arkansas

April 22, 2019

______Research Supervisor

______Faculty Reader

______Director of Undergraduate Studies

Honors thesis submitted in partial fulfillment of the requirements for graduation with Distinction in Biology in Trinity College of Duke University

Abstract In the era of Anthropocene, an increasing part of the terrestrial environments is losing their ecosystem services and function, negatively affecting both human economics and the ecological system. Phytoremediation, the use of plants to reverse degradation and to restore ecological function, has been a promising approach. However, the symbiotic soil microbiota that influence the effectiveness of this method is not fully understood. I sampled the soil and roots of Pinus spp. (pines) at four sites along a gradient of vegetation recovery on the Superfund Site Brewer Gold Mine (SC), the Henry Knob Mine (SC), and Russell Gold Mine. The acidity, nutrient profile and heavy-metal contamination of collected soil is determined. DNA is extracted from the soil and root samples with PowerSoil DNA Isolation Kit, followed by preparation of multiplex PCR samples of the ITS region. Sequence reads generated through Illumina Miseq is processed through QIIME pipeline and taxonomy assigned through UNITE database. The results show a pattern of succession in fungal communities along a recovery gradient. While the mycorrhizal fungi on the least recovered site are dominated by Rhizopogon sp. and Pisolithus sp., sites with more recovered vegetation reveal a more diverse array of symbiotic fungi, including Amanita sp. and Russula sp.. These diverse fungi, although came later in the succession pattern, likely brings more diverse benefits to help their hosts cope with the stressful environment. This “bioprospecting” method could be applied to extract and amplify symbiotic fungi to facilitate revegetation efforts.

Introduction A major feature of the Earth’s most current geological epoch, the Anthropocene, is the unmistakeable impact of human activity on ocean and terrestrial environments. One prime example of major terrestrial impacts are mine sites, whose activities result in loss of topsoil, reduced soil fertility and biodiversity, and high concentrations of pollutants such as toxic metals (Wang, 2017). The use of plants has been proposed as a major mechanism to reverse degradation processes and restore ecological function at mine sites through phytoremediation (Singh, Raghubanshi, & Singh, 2002), which is assisted by mutualistic plant-associated microbes that play an important role in revegetation and succession (Dickie et al., 2013). Mutualistic symbiotic fungi, including mycorrhizal fungi and endophytic fungi, confer a range of fitness benefits that help plants adapt to stress, including improved nutrient acquisition, and tolerance to drought, heat, metals, herbivory, and organic contaminants (Marrs, 2016). [Write about mycorrhizal fungi in particular] The study of such fungi-plant symbiosis on mining-impacted sites could provide valuable insights into the utilization of beneficial fungi to facilitate ecological restoration. A number of studies have demonstrated the potential of symbiotic fungi to assist plant survival at heavy-metal contaminated sites. Mycorrhizal fungi in particular are known to improve the metal tolerance of their host plants. For example, AM fungi have been reported to occur on the roots of Viola calaminaria in Zn/Pb-rich soil and Berkheyla coddii in Ni-rich soil, accumulating and immobilizing heavy metals in their structures (Gaur & Adholeya, 2004). In experiment with zinc-rich soil, ectomycorrhiza Suillus bovinus have been shown to improve the nutrient uptake and growth of Pinus sylvestris (Adriaensen, Vangronsveld, & Colpaert, 2006). Similar benefits to hosts have been observed in endophytic fungi: Shi et al. show that Fusarium sp. and Penicillium sp. associated with Brassica napus improved the rape biomass and metal extraction efficiency of their host in Cd and Pb contaminated soil (Shi et al., 2017). Many of these in-vitro studies have focused on a limited array of fungi. The most heavily studied fungal genera include Suillus (Adriaensen, van der Lelie, Van Laere, Vangronsveld, & Colpaert, 2003; Colpaert, Vandenkoornhuyse, Adriaensen, & Vangronsveld, 2000), Thelephora (Van Tichelen, Colpaert, & Vangronsveld, 2001), and Glomus (Amna et al., 2015). Indigenous fungal communities likely hold other promising fungal strains for phytoremediation that may adapt better to the local environment, establish a more mutualistic interaction with their host and pose less interruption to the surrounding soil microbiota (Faye et al., 2013; Mummey, Antunes, & Rillig, 2009). Study of these fungal communities could provide new insights into the co-existence of multiple strains of symbiotic fungi under various ecological stresses (different heavy-metal contamination, acidity, drought). Early-stage, or early successional fungi are known to inhabit mineral soils of a low nutrient content or with extreme pH values (Münzenberger, Golldack, Ullrich, Schmincke, & Hüttl, 2004). Environmental filtering acts on the fungal communities on mine sites: ECM communities are influenced strongly by the soil features of the sites, such as the low levels of organic matter, soil structure, pH values, soil temperature and moisture (Huang et al., 2012; Staudenrausch, Kaldorf, Renker, Luis, & Buscot, 2005). ECM fungal species that are found on the roots of Pinus massoniana growing on Pb-Zn mine sites includes Cenococcum geophilum, Rhizopogon buenoi, Tomentella ellisii, Inocybe curvipes and Suillus granulatus on Pinus massoniana (Huang et al., 2012). Pisolithus tinctorius is a typical, widespread early-stage ECM fungus, reported to be well- adapted to the low pH and high temperatures of anthracite and hard coal spoils (Schramm, 1966; Marx, 1975). Fungal community might shift through the succession. The fungal communities in pioneer pine forests on metal mines in Belgium revealed that it is initially dominated by dark Ascomycota, but metal tolerance basidiomycetes such as Suillus luteus become more frequent within two years (Op De Beeck et al., 2015). A review of studies on ectomycorrhizal fungal genera across soil chronosequences suggests putative trait groupings of different genera: early successional genera likely include ones with low host-specificity, forming small, inconspicuous sporocarps that are long-term persistent (Cenococcum, Tomentella / Thelephora, Inocybe, Cortinarius and Laccaria); early-mid successional genera may include ones with high host- specificity and large sporocarp (Rhizopogon and Suillus); and the genera most common in mature ecosystems include Amanita, Russula, Lactarius and Boletus (Dickie et al., 2013). In reforestation of the mine sites, inoculation of selected ECM fungi potentially confers key advantages for tree seedlings to adapt to environmental stress and adverse substrates during the first few years after outplanting (Kałucka & Jagodziński, 2016). Such inoculation is most effective when fungal inocula are adapted to the environmental conditions of transplantation sites (Rincón, de Felipe, & Fernández-Pascual, 2017). A better understanding of the ECM fungal community indigenous to the mine sites and surrounding forest areas is therefore crucial to facilitate better restoration of sites impacted by anthropogenic activities. In this study, the fungal communities associated with pines (Pinus spp.) were studied on three abandoned mineral and metal mine sites in the Carolinas—the Brewer Gold Mine, Henry’s Knob Mine (kyanite), and Russell Gold Mine. The goals of this study were to characterize the fungal community, in particular the ECM fungi, on pines growing on the mine sites and to evaluate the effects of soil characteristics on these communities.

Methods Field sites Brewer Gold Mine Brewer Gold Mine (BGM) is a Superfund site near Jefferson, South Carolina. BGM has mined significant ore from several open pits (over 12 million tons of ore and waste rock from 1987 to 1995). When EPA took over the site in 1995, serious concerns were raised over multiple contamination sources, including pollution from the cyanide solution used for extracting gold, acid mine drainage (AMD) and heavy-metal contamination. BGM thus serves as a prime field site to study plant-fungal symbiosis under multiple stress factors. Upon consultation with our mine site contact, we decided on four experimental sites (Figure 1) with one control site in the nearby forest. From each site, we collected three soil samples under Pinus taeda (loblolly pine) and the corresponding root samples.

Henry’s Knob Mine The Henry’s Knob Mine (HKM) was utilized for the extraction of kyanite, an aluminum silicon oxide used in the manufacture of high-alumina brick and other high temperature, refractory materials. Mining activities ceased in the 1970’s and with common duty of care at that time, mine tailings were present at several large areas of fine grained mineral sediment located down slope of former mineral processing facilities. The site is approximately 185 acres in size, mainly a combination of wooded tree growth and revegetated tailings impoundments. We collected in four separate areas of concern (AOC) – site HKM_1, HKM_2, HKM_3, and HKM_4 (Figure 2). A control sample were also collected from a nearby forest (HKM_F).

Russel Gold Mine The Russell Gold Mine (RGM) operated a series of pits and shafts around 1894. The geology of the site is composed of white, iron-stained, siliceous material bordered by phyllite, with pyrite scattered through the zone (Carpenter, 1976). After consulting with the US Forest Service staff, we decide to collect on the old pit (RGM_Pit) and processing site (RGM_Process) (Figure 3).

Silver Hill Silver Hill mine (SHM) is located in south-central Davidson County, and was under active mining from 1838-1882. Exposed mining shafts are still present on SH (Hudson Institute of Mineralogy, 2019). Samples were collected from four sites, including bare mine soil (SHS) with no plants growing above the ground, b) soil colonized by copper moss (SHM), c) isolated colonizing pines (SHP), d) adjacent Pine forest (F) (Figure 4).

Collection on the field BGM, HKM, and RGM samples were collected during 2017 and 2018. At BGM and HKM, we sampled four sites of varying level of disturbance from the mine site (BGM_1~4, HKM_1~4), and one control site from the nearby forest (BGM_F and HKM_F). For each site, three pine trees, preferably P. taeda, were randomly chosen. For each tree, a composite soil sample of 4 cores was sampled from A-horizon and fine root sample was taken. At RGM, we collected soil and fine root samples from P. taeda growing on two sites—an old open pit and the metal processing site (RGM_Pit and RGM_Process). SH samples were collected during a pilot study in 2015. A-horizon soil samples were collected from four sites: (1) the exposed mining shaft (SH_S), (2) patches of copper moss growing alongside the shaft (SH_M), (3) pine trees growing at the edge of the clearing (SH_P), and (4) the forest area surrounding the site (SH_F).

Soil analysis and DNA analysis The soil from each site is sent to the Soil, Plant and Water Laboratory at University of Georgia to test: acidity and other soil characteristics (pH, lime buffering capacity, Cation Exchange Capacity), total C and N, and plant available elements (Al, As, Ca, Cd, Cr, Cu, Fe, K, Mg, Mn, Mo, Na, Ni, P, Pb, Zn) with the Mehlich 1 method. The composition of the fungal community was assessed using similar approached used by the Vilgalys Lab to survey pine and cottonwood microbiomes (Bonito, Reynolds et al. 2014, Talbot, Bruns et al. 2014, Glassman, Peay et al. 2015). DNA was from the soil sample (PowerSoil DNA Isolation Kit from Mobio) and from the root sample (Phenol:Chloroform extraction), followed by preparation of a pooled library of ITS PCR products. The samples were sequenced in three sequencing runs: the SH samples were sequenced with Illumina MiSeq, BGM sequences were sampled with Illumina NanoSeq, and BGM, HKM and RGM samples were sequenced with Illumina MiSeq. All the sequencing data were combined for downstream analysis.

Data analysis and visualization Sequenced reads were de-multiplexed, quality filtered, and processed using the QIIME pipeline (Caporaso, Kuczynski et al. 2010) and clustered into operational taxonomic units (OTUs) employing both de novo and reference based methods (against the QIIME 12_11 alpha release of curated ITS sequences from the UNITE database). Sequences were clustered into OTUs with USEARCH using a 97% sequence similarity threshold and a minimal cluster size of two. Taxonomy was assigned by searching representative sequences from each OTU against the UNITE database (Abarenkov, Henrik Nilsson et al. 2010). The OTUs were annotated by FUNGuild to ecological guilds (Nguyen et al., 2016). To visualize the most abundant OTUs from the experimental sites (excluding the forest sites), top 25 OTUs from each mine were selected out. The relative abundance of each OUT on each site (including both experimental and control sites) were mapped onto a table. The R package “metacoder” was used to visualize the most common taxa of the class Agaricomyetes on the mine sites compared to the forest sites (Rice, Longden, & Bleasby, 2000). The top 20 OTUs across all mine sites were mapped to a heat map representing their relative abundance on each site through the R package “ampvis” (Andersen, Kirkegaard, Karst, & Albertsen, 2018). The R package “phyloseq” was used to perform Principal Coordinate Analysis (PCoA) (McMurdie & Holmes, 2013). Because Henry’s Knob Mine forest sample have larger than expected distance from the other samples, they are taken out during the associated environmental variables analysis.

Results Soil Analysis On both BGM and HKM, we do not observe a consistent high concentration of heavy metals compared to the nearby forest site (Table 2). However, the forest sites near both mines have higher Cation Exchange Capacity (CEC), a characteristic of the soil that retains nutrients in plant-available form (Table 1). This could explain why the forest sites have higher concentration of elements, some of which are macronutrients (C, N, P, K, Ca, Mg) and micronutrients (Fe, Mn, Mo, Cu, Zn, Ni).

Metagenomic Analysis 2794103 reads were processed. Overall, 1473 OTUs were identified in 177 samples with 97% threshold. A variety of fungal taxa are present in the soil and pine roots on these mine sites. The classes Agaricomycetes, which contains many mycorrhizal members, and Dothidiomycetes were abundant across the sites that we sampled (Figure 5). Many mycorrhizal genera such as Boletus, Pisolithus, Rhizopogon, and Scleroderma found on these sites (Figure 6, Table 3). The FUNguild analysis confirms the abundance of mycorrhizal fungi on these sites (Figure 7). The heat trees suggest that there is a consistent trend when disturbed mine sites were compared with nearby forest sites: species more common on mine sites includes Pisolithus, Rhizopogon, and species of the Thelephoraceae family; species more common on forest sites include Amanita, Russula, Clavulina, Lactarius, Piloderma, Trechispora, and Inocybe (Figure 8, Figure 9). PCoA suggests that there are distinct local community assemblages on each site (Figure 10, Figure 11). In addition, there is a clear shift in community composition from mine sites to forest sites. The environmental factors analysis confirms with findings from the soil analysis that the forest soils tend to have higher CEC, a range of elements, and higher pH value (Figure 12, Figure 13); these environmental variables plays a role in driving the shifts in community composition, although the PCoA suggests that there is not a clear trend.

Discussion The infertility of most soils from Brewer Gold Mine and Henry’s Knob Mine is best attributed to nutrient leaching and acid mine drainage, instead of heavy metal contamination. Future mine site restoration should pay specific attention to combined effects of acidity, leaching and potential metal contamination. Making sure that the plants are resilient to acidity and able to obtain enough nutrients through deliberate choice of plant species, adding amendments, or increasing the benefits from symbiotic fungi are potential strategies to make restoration projects more effective. The recovering mine soils harbor a diverse community of symbiotic fungi that likely play an important role in the restoration of mine sites. We observed a high abundance of Agaricomycetes, many members of which are mycorrhizal, on mine sites. Two fungal genera (Pisolithus and Rhizopogon) and one fungal family (Thelephoraceae) stand out as abundant members of the fungal communities in the less recovered site. These species are likely to be essential for the early stage of succession. Pisolithus has been reported to be common to a variety of habitats across the United States and around the world (Grand, 1976). In particular, it is consistently found in areas of poor conditions and are sparsely vegetated (Medve & Gill, 1982). Both Pisolithus and Rhizopogon have been frequently applied to reforestation (PHC Reclamation, Inc., n.d.; Steinfield, Amaranthus, & Cazares, 2003). In the context of phytomrediation, these strains could be augmented and inoculated to young pine seedlings during initial phase of revegetation to facilitate establishment. Ectomycorrhizal fungi that come later in the recovery process include Russula spp. and Amanita spp.. Our findings contribute to the understanding of the indigenous fungal community present on the mine sites and on nearby undisturbed sites, and points to a successional trend of fungal community on Carolinian mine sites; such understanding could be harnessed to improve reforestation practices by intentionally facilitating the mutualistic mycorrhizal relationship. However, one question remains unsolved: what contributes to the successional pattern? In other words, are the early succession species such as Pisolithus and Rhizopogon better symbionts in poor condition, or are they poor competitors with other ectomycorrhizal fungi (Lampky & Peterson, 1963)? Future studies should look into the effectiveness of different strategies of enhancing soil mycorrhizal community to improve plant growth, including augmenting early-succession species and late-comers. Whether or how each environmental variable impact the composition of mycorrhizal community remains unclear. We recommend more studies to be done on the impact of soil characteristics on the fungal community composition, with data from more mine sites around the world.

Conclusion In sum, our study contributes to a growing literature on the succession of fungal community on disturbed mine sites. In the era of Anthropocene, we are faced with the challenge to improve and harness our understandings of the “coherent, dynamic unit” of the tree- rhizhosphere interrelationship to ultimately restore the “stability and resilience” of the ecosystem (Perry, Molina, & Amaranthus, 1987). Our findings point to a promising array of ectomycorrhizal fungi on the mine sites, with a successional trend where genera such as Pisolithus and Rhizopogon establish in the early successional stage, and Amanita and Russula became more common in the later stage. We recommend more studies on the application of ectomycorrhizal fungi from different succession stages in aiding phytoremediation, and more understanding in the impact of environmental variables on the succession of fungal community.

Figure 1. Four experimental sites on BGM, along a pollution gradient.

Figure 2. Google Earth image of four experimental sites on Henry’s Knob Mine.

Figure 3. Google Earth image of two experimental sites on Russel Gold Mine.

(A) (B) Figure 4. A) Goggle Earth image of Silver Hill Mine. B) Silver Hill mine drainage field showing sites where soil samples were collected.

Table 1. Properties of soils from different sites.

LBC Equivalent CEC Base Saturation (%) C (%) N (%) (ppm CaCO3/ pH) Water pH (meq/100g) BGM 1 382 4.12 4.58 6.69 0.54 0.02

BGM 2 322 4.01 5.07 5.88 0.86 0.02

BGM 3 526 3.85 2.27 9.83 0.14 <0.02

BGM 4 511 4.22 3.69 8.56 1.12 0.05

BGM Forest 1960 3.96 2.62 35.49 6.85 0.20

HKM 1 173 4.06 15.89 3.12 1.24 0.04

HKM 2 437 4.17 6.40 7.66 1.81 0.05

HKM 3 386 4.43 27.38 7.92 2.29 0.07

HKM 4 237 4.34 9.53 4.01 0.46 <0.02

HKM Forest 2998 3.43 5.67 65.81 23.54 0.67

RGM Process 475 4.67 11.14 7.22 4.25 0.25

RGM Pit 569 4.95 45.87 12.50 4.40 0.29

Silver Hill Forest 264 5.25 56.27 6.13 1.91 0.06

Silver Hill Pine 120 6.57 90.69 2.33 0.27 <0.02

Silver Hill Moss 354 5.59 41.38 4.94 0.17 <0.02 Table 2. Plant-available element concentration in soils from different sites. All units are in mg/kg (ppm).

Al As Ca Cd Cr Cu Fe K Mg Mn Mo Na Ni P Pb Zn

BGM 1 127.8 <0.15 35.19 <0.05 <0.05 2.49 41.9 13.60 6.86 2.95 <0.05 8.85 <0.05 1.35 0.17 0.40

BGM 2 131.7 <0.14 28.97 <0.04 <0.04 1.05 34.8 10.18 5.69 0.54 <0.04 18.40 <0.04 1.36 0.04 0.23

BGM 3 185.5 <0.13 28.10 <0.04 0.07 1.93 37.1 3.72 5.56 <0.21 <0.04 6.28 <0.04 0.36 <0.02 <0.21

BGM 4 248.8 <0.13 29.63 <0.04 0.06 10.20 51.4 20.07 5.02 <0.21 <0.04 17.09 0.08 2.74 0.49 0.69

BGM Forest 686.3 0.33 98.15 0.11 <0.06 1.23 224.2 45.02 27.99 2.35 <0.06 20.65 0.13 3.85 2.70 0.97

HKM 1 47.2 <0.18 57 <0.06 <0.06 0.31 107.1 20.8 16.5 9.0 <0.06 4.3 <0.06 1.8 1.4 0.9

HKM 2 160.3 <0.19 48 <0.06 <0.06 0.50 67.8 41.7 15.0 13.6 <0.06 4.3 <0.06 2.1 0.8 0.6

HKM 3 157.3 <0.17 338 <0.06 <0.06 1.57 82.9 38.1 42.7 7.3 <0.06 6.1 0.19 4.9 0.5 1.8

HKM 4 35.8 <0.17 40 <0.06 <0.06 0.33 38.8 19.1 13.4 <0.28 <0.06 4.6 <0.06 2.9 5.5 0.4

HKM Forest 742.7 <0.46 446 <0.15 <0.15 <0.76 173.1 112.1 129.5 30.3 <0.15 30.8 0.89 15.5 6.0 6.7

RGM Process 34.7 <0.24 115 <0.08 <0.08 <0.39 39.5 16.2 20.2 9.2 <0.08 4.6 0.13 3.0 0.5 0.6

RGM Pit 176.2 <0.23 894 <0.07 <0.07 <0.37 27.3 68.9 125.8 145.1 <0.07 9.1 0.90 10.7 0.6 5.4

Silver Hill Forest 175.7 <0.18 388 0.13 <0.06 9.33 50.7 80.0 98.1 41.8 <0.06 111.1 0.15 12.5 80.1 18.9

Silver Hill Pine 99.5 0.25 250 1.24 <0.04 14.37 43.9 19.4 82.0 22.3 <0.04 29.7 0.10 28.2 120.6 416.8

Silver Hill Moss 47.7 0.35 319 0.62 <0.04 40.76 62.7 28.0 27.5 28.4 <0.04 33.5 0.07 33.0 98.1 283.6

Figure 5. Fungal OTUs on each site assigned to class level.

Figure 6. Fungal OTUs on each site assigned to genus level. Table 3. Top 25 OUTs from disturbed sites on each mine and their relative abundance (percentage) on each site. R denotes for root samples, S denotes for soil samples. Taxa OTU BGM_1 BGM_2 BGM_3 BGM_4 BGM_F HKM_1 HKM_2 HKM_3 HKM_4 HKM_F

R S R S R S R S R S R S R S R S R S R S c__Agaricomycetes;f__Amanitaceae;g 15.0 20.6 17.7 OTU_35 1.3 0 0.07 0.39 5.62 0.01 0 __Amanita 4 6 2 c__Agaricomycetes;f__Boletaceae;g__ 13.1 OTU_128 0 0 0 0.03 0 0.05 2.29 0 0 Boletus 8 c__Agaricomycetes;f__Clavulinaceae; OTU_65 0 0 0.65 8.05 0.01 0 0 0 0 0 g__Clavulina c__Agaricomycetes;f__Cortinariaceae; 16.5 OTU_24 0 0 0.01 2.56 0.02 0 0.01 0 0 g__Cortinarius 9 c__Agaricomycetes;f__Exidiaceae;g__ OTU_394 unidentified c__Agaricomycetes;f__Hydnodontacea OTU_202 e;g__Trechispora c__Agaricomycetes;f__Hygrophoracea OTU_105 e;g__Hygrocybe c__Agaricomycetes;f__Inocybaceae;g_ OTU_304 _Inocybe c__Agaricomycetes;f__Pisolithaceae;g 31.0 12.8 OTU_137 0 0.01 0.03 0.16 5.37 0.33 0 0 __Pisolithus 5 8 c__Agaricomycetes;f__Rhizopogonace OTU_261 8.74 2.5 0 0.89 0.71 0.01 3.42 0.98 0 0 ae;g__Rhizopogon c__Agaricomycetes;f__Russulaceae;g_ OTU_106 0 0 3.42 4.24 0 0.48 0 0.05 0 0 _Russula c__Agaricomycetes;f__Russulaceae;g_ OTU_409 _Russula c__Agaricomycetes;f__Russulaceae;g_ OTU_100 _Russula c__Agaricomycetes;f__Russulaceae;g_ OTU_155 _unidentified c__Agaricomycetes;f__Sclerodermatac OTU_108 eae;g__Scleroderma c__Agaricomycetes;f__Thelephoraceae OTU_185 3.9 3.58 4.83 2.17 0 0 0 0.01 0 0 ;g__unidentified c__Agaricomycetes;f__unidentified;g_ 13.3 OTU_39 4.13 0 0 0 0 0 0 0 0 0 0 0.01 4.61 0 0 0 0.01 0 0 _unidentified 9 c__Agaricomycetes;f__unidentified;g_ 10.2 OTU_63 9.58 0 0.01 0 0 4.65 0.04 0 0 _unidentified 1 c__Agaricomycetes;f__unidentified;g_ OTU_46 0 0 6.85 3.87 0.01 0 0 0 0 0 _unidentified c__Agaricomycetes;f__unidentified;g_ OTU_473 _unidentified c__Archaeorhizomycetes;f__Archaeor hizomycetaceae;g__Archaeorhizomyce OTU_36 0 1.67 0.01 3.7 0.01 0.36 0 0.95 0 0 s c__Dothideomycetes;f__Didymellacea OTU_182 e;g__Phoma c__Dothideomycetes;f__Gloniaceae;g_ 31.4 OTU_26 0.21 0.83 0.03 0.57 2.93 1.08 1.36 0.32 0.13 _Cenococcum 1 c__Dothideomycetes;f__Leptosphaeria 44.1 OTU_1417 0.05 0.1 0.02 0.01 1.09 0.14 0.04 0 0.1 ceae;g__Leptosphaeria 8 c__Dothideomycetes;f__Leptosphaeria 27.3 OTU_1361 0.01 0.01 0 0.02 0 0.06 0.01 0 0.01 ceae;g__Leptosphaeria 8 c__Dothideomycetes;f__Leptosphaeria OTU_1416 5.05 2.74 2.69 1.88 3.68 2.66 3.09 2.16 0.28 0.41 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1450 0.01 0.17 0.01 0.01 0.01 1.21 2.02 9.85 0.02 0.01 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1435 0.05 0.58 0.01 0.2 0 0.05 0.15 10.2 0 0.01 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria 10.8 OTU_1371 0.04 0.01 0.01 0.01 0 0.09 0.01 0 0.01 ceae;g__Leptosphaeria 9 c__Dothideomycetes;f__Leptosphaeria OTU_1427 0.1 0.08 0.02 0.02 0.04 10.1 0.03 0.05 0.12 0.05 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1213 0.76 1 0.55 6.58 0.34 0.55 0.25 0.32 0.04 0.11 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1426 0.23 0.04 0.03 7.83 0.01 0.12 0.05 0.13 0 0.37 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1382 0.26 0.03 0.01 0.03 0.11 0.04 0.77 6.79 0.59 0.03 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1405 0.01 0.12 0.05 6.37 0.01 0.11 0.03 0.05 0 0.04 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1358 0.19 0.02 0.85 0.05 0.04 0.05 0 4.79 0.1 0.02 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1488 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1487 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1525 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1495 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1483 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1285 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1269 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1368 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1527 ceae;g__Leptosphaeria c__Dothideomycetes;f__Leptosphaeria OTU_1455 ceae;g__Leptosphaeria c__Dothideomycetes;f__Teratosphaeri OTU_52 aceae;g__Devriesia c__Dothideomycetes;f__unidentified;g 16.2 OTU_1404 0.06 0.16 0.01 0.06 0.2 0.02 0.02 0.02 0.07 __unidentified 5 c__Dothideomycetes;f__unidentified;g 13.8 OTU_1428 0.72 0.01 0.05 0.05 0.02 0.06 0.04 0.05 0.01 __unidentified 2 c__Dothideomycetes;f__unidentified;g 13.7 OTU_1442 0.02 0.07 0.49 0.05 0.01 0.01 0.14 0.07 0.11 __unidentified 9 c__Dothideomycetes;f__unidentified;g 11.3 OTU_1406 1.36 0.01 0 0.01 0 0.28 0.01 0 0.04 __unidentified 9 c__Dothideomycetes;f__unidentified;g OTU_1448 0.03 10.5 0.01 0.02 0.05 0.01 0.46 0.03 0.08 0.04 __unidentified c__Dothideomycetes;f__unidentified;g 10.0 OTU_1429 0.01 0.04 0.1 0.02 0 0.04 0 0.03 0.01 __unidentified 5 c__Dothideomycetes;f__unidentified;g OTU_1459 0 9.12 0.01 0.04 0 0.02 0 0 0 0.02 __unidentified c__Dothideomycetes;f__unidentified;g OTU_1390 0.03 0.02 0.09 7.2 0.09 0.13 0.05 0.03 0.12 0.08 __unidentified c__Dothideomycetes;f__unidentified;g OTU_1463 0.04 6.7 0.01 0.02 0.2 0.01 0.06 0.01 0.55 0.01 __unidentified c__Dothideomycetes;f__unidentified;g OTU_1443 0.24 0.02 0.03 0 0.66 5.36 0 0.01 0.61 0 __unidentified c__Dothideomycetes;f__unidentified;g OTU_1444 0 0 0 0 0 0.01 0.03 6.3 0.2 0 __unidentified c__Dothideomycetes;f__unidentified;g OTU_1511 0.01 0 0 0 0.04 0 0 4.99 0 0.01 __unidentified c__Dothideomycetes;f__unidentified;g OTU_1397 __unidentified c__Dothideomycetes;f__unidentified;g OTU_1454 __unidentified c__Dothideomycetes;f__unidentified;g OTU_1505 __unidentified c__Eurotiomycetes;f__Aspergillaceae; OTU_435 g__Aspergillus c__Eurotiomycetes;f__unidentified;g__ OTU_34 0.1 0.47 7.4 3.28 1.44 2.78 3.54 1.18 1.72 0.9 unidentified c__Eurotiomycetes;f__unidentified;g__ OTU_372 unidentified c__Lecanoromycetes;f__Parmeliaceae; OTU_776 g__Protoparmelia c__Leotiomycetes;f__Helotiaceae;g__ 18.9 OTU_28 4.42 5.4 1.58 9.13 3.16 5.51 2.08 5.89 1.4 Meliniomyces 1 c__Leotiomycetes;f__Helotiales_fam_I OTU_300 ncertae_sedis;g__Leptodontidium c__Leotiomycetes;f__Hyaloscyphaceae 16.8 10.2 OTU_76 2.2 1.24 0.24 1.89 1.31 0.2 5.52 2.45 ;g__unidentified 2 7 c__Leotiomycetes;f__Hyaloscyphaceae OTU_64 0.41 0.05 9.55 2.13 6.18 2.06 3.11 0.71 0 0.16 ;g__unidentified c__Leotiomycetes;f__Hyaloscyphaceae OTU_94 ;g__unidentified c__Leotiomycetes;f__Myxotrichaceae; OTU_89 0.18 0.47 4.4 1.33 2 1.71 8.99 1.65 0.29 0.2 g__Oidiodendron c__Leotiomycetes;f__Myxotrichaceae; OTU_91 0.03 0.44 0.53 2.32 0.43 1.82 2.22 1.37 0.02 0.21 g__Oidiodendron c__Leotiomycetes;f__Myxotrichaceae; OTU_284 g__Oidiodendron c__Leotiomycetes;f__unidentified;g__ OTU_183 unidentified c__Leotiomycetes;f__Vibrisseaceae;g_ OTU_687 _Phialocephala c__Mortierellomycetes;f__Mortierellac OTU_60 0.14 4.7 0.03 6.18 0.01 0.02 0 0 0.08 1.52 eae;g__Mortierella c__Mortierellomycetes;f__Mortierellac OTU_61 0 1.06 0.05 6.65 0.01 0 0.4 0.13 0 0 eae;g__Mortierella c__Mortierellomycetes;f__Mortierellac OTU_27 0.16 3.46 0.06 1.27 0.01 2.42 0.09 0.01 0.05 0.25 eae;g__Mortierella c__Mortierellomycetes;f__Mortierellac OTU_1122 eae;g__Mortierella c__Rhizophydiomycetes;f__unidentifie OTU_38 d;g__unidentified c__Rozellomycotina_cls_Incertae_sedi OTU_133 s;f__unidentified;g__unidentified c__Umbelopsidomycetes;f__Umbelops OTU_29 0 0 0.17 5.8 0 0.3 0.09 0.34 0.01 1.21 idaceae;g__Umbelopsis c__unidentified;f__unidentified;g__uni OTU_59 1.47 0.28 5.36 2.61 0.03 0 1.71 3.05 0 0 dentified c__unidentified;f__unidentified;g__uni 24.5 OTU_58 1.3 0 0 0 0 4.2 0 0 0 dentified 6 c__unidentified;f__unidentified;g__uni OTU_21 dentified c__unidentified;f__unidentified;g__uni OTU_33 dentified c__unidentified;f__unidentified;g__uni OTU_25 dentified c__unidentified;f__unidentified;g__uni OTU_75 dentified c__unidentified;f__unidentified;g__uni OTU_23 dentified c__unidentified;f__unidentified;g__uni OTU_30 dentified c__unidentified;f__unidentified;g__uni OTU_528 dentified c__unidentified;f__unidentified;g__uni OTU_116 dentified c__unidentified;f__unidentified;g__uni OTU_422 dentified c__unidentified;f__unidentified;g__uni OTU_37 dentified

Taxa OTU SH_F_ SH_F_ SH_P_ SH_P_ RGM_Proces RGM_Proce RGM_Pit Root Soil Root Soil s_Root ss_Soil _Soil c__Agaricomycetes;f__Amanitaceae;g__Amani ta OTU_35 c__Agaricomycetes;f__Boletaceae;g__Boletus OTU_128 c__Agaricomycetes;f__Clavulinaceae;g__Clavu lina OTU_65 c__Agaricomycetes;f__Cortinariaceae;g__Corti narius OTU_24 c__Agaricomycetes;f__Exidiaceae;g__unidentif ied OTU_394 3.13 6.08 0.05 0 c__Agaricomycetes;f__Hydnodontaceae;g__Tre chispora OTU_202 0 5.29 0.03 0.45 c__Agaricomycetes;f__Hygrophoraceae;g__Hy grocybe OTU_105 0 0 10.78 1.5 c__Agaricomycetes;f__Inocybaceae;g__Inocyb e OTU_304 0 0 4.67 0 c__Agaricomycetes;f__Pisolithaceae;g__Pisolit hus OTU_137 c__Agaricomycetes;f__Rhizopogonaceae;g__R hizopogon OTU_261 c__Agaricomycetes;f__Russulaceae;g__Russula OTU_106 c__Agaricomycetes;f__Russulaceae;g__Russula OTU_409 0.1 0 10.95 10.28 c__Agaricomycetes;f__Russulaceae;g__Russula OTU_100 0 0 0 12.34 c__Agaricomycetes;f__Russulaceae;g__unident ified OTU_155 0.01 0 0 3.4 c__Agaricomycetes;f__Sclerodermataceae;g__S cleroderma OTU_108 0 0 47.19 c__Agaricomycetes;f__Thelephoraceae;g__unid entified OTU_185 c__Agaricomycetes;f__unidentified;g__unidenti fied OTU_39 c__Agaricomycetes;f__unidentified;g__unidenti fied OTU_63 c__Agaricomycetes;f__unidentified;g__unidenti fied OTU_46 c__Agaricomycetes;f__unidentified;g__unidenti fied OTU_473 0 0 0 3.86 c__Archaeorhizomycetes;f__Archaeorhizomyce taceae;g__Archaeorhizomyces OTU_36 1.54 0 0 3.16 c__Dothideomycetes;f__Didymellaceae;g__Pho ma OTU_182 0 2.12 0.01 c__Dothideomycetes;f__Gloniaceae;g__Cenoco ccum OTU_26 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1417 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1361 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1416 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1450 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1435 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1371 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1427 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1213 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1426 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1382 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1405 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1358 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1488 0 2.71 0 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1487 0 2.4 0 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1525 0 2.4 0 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1495 0 2.15 0 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1483 0 2.1 0 24.38 0 0 0.02 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1285 0 1.81 0 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1269 0 1.75 0 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1368 0.02 26.89 0.01 0.04 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1527 0 0 4.23 0.06 c__Dothideomycetes;f__Leptosphaeriaceae;g__ Leptosphaeria OTU_1455 0 0.13 0.01 3.24 c__Dothideomycetes;f__Teratosphaeriaceae;g_ _Devriesia OTU_52 0 6.77 0.2 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1404 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1428 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1442 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1406 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1448 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1429 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1459 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1390 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1463 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1443 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1444 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1511 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1397 0 1.55 0 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1454 16 0 0 0.01 c__Dothideomycetes;f__unidentified;g__uniden tified OTU_1505 0.01 0.18 0.18 13.39 c__Eurotiomycetes;f__Aspergillaceae;g__Asper gillus OTU_435 0 2.11 0 c__Eurotiomycetes;f__unidentified;g__unidenti fied OTU_34 c__Eurotiomycetes;f__unidentified;g__unidenti fied OTU_372 0 1.84 0.01 c__Lecanoromycetes;f__Parmeliaceae;g__Proto parmelia OTU_776 0 2.05 0.01 c__Leotiomycetes;f__Helotiaceae;g__Meliniom yces OTU_28 c__Leotiomycetes;f__Helotiales_fam_Incertae_ sedis;g__Leptodontidium OTU_300 5.85 2.69 0.03 0.82 c__Leotiomycetes;f__Hyaloscyphaceae;g__unid entified OTU_76 c__Leotiomycetes;f__Hyaloscyphaceae;g__unid entified OTU_64 c__Leotiomycetes;f__Hyaloscyphaceae;g__unid entified OTU_94 10.34 0.01 0 0 c__Leotiomycetes;f__Myxotrichaceae;g__Oidio dendron OTU_89 c__Leotiomycetes;f__Myxotrichaceae;g__Oidio dendron OTU_91 c__Leotiomycetes;f__Myxotrichaceae;g__Oidio dendron OTU_284 3.51 2.81 0.03 0.08 c__Leotiomycetes;f__unidentified;g__unidentifi ed OTU_183 5.13 2.08 0 0.09 c__Leotiomycetes;f__Vibrisseaceae;g__Phialoc ephala OTU_687 10.2 8.67 0.04 0.45 c__Mortierellomycetes;f__Mortierellaceae;g__ Mortierella OTU_60 c__Mortierellomycetes;f__Mortierellaceae;g__ Mortierella OTU_61 c__Mortierellomycetes;f__Mortierellaceae;g__ Mortierella OTU_27 0.66 0.18 15.47 6.39 c__Mortierellomycetes;f__Mortierellaceae;g__ Mortierella OTU_1122 0 0 3.58 0.48 c__Rhizophydiomycetes;f__unidentified;g__uni dentified OTU_38 0 7.42 0.36 c__Rozellomycotina_cls_Incertae_sedis;f__unid entified;g__unidentified OTU_133 0 9.65 0.22 0 c__Umbelopsidomycetes;f__Umbelopsidaceae; g__Umbelopsis OTU_29 0 0.01 1.58 1.55 c__unidentified;f__unidentified;g__unidentified OTU_59 0 2.14 0 c__unidentified;f__unidentified;g__unidentified OTU_58 c__unidentified;f__unidentified;g__unidentified OTU_21 0 8.79 0.4 c__unidentified;f__unidentified;g__unidentified OTU_33 0 0 7.27 c__unidentified;f__unidentified;g__unidentified OTU_25 0 5.65 0 c__unidentified;f__unidentified;g__unidentified OTU_75 0 5.34 0.09 c__unidentified;f__unidentified;g__unidentified OTU_23 0 5.37 0 c__unidentified;f__unidentified;g__unidentified OTU_30 0 3.88 0 c__unidentified;f__unidentified;g__unidentified OTU_528 0 1.99 0 c__unidentified;f__unidentified;g__unidentified OTU_116 0 1.73 0.03 c__unidentified;f__unidentified;g__unidentified OTU_422 0 1.7 0.01 c__unidentified;f__unidentified;g__unidentified OTU_37 0.01 0.01 10.01 0.28

Figure 7. FUNguild analysis showing the relative abundance of OTUs of each ecological guild.

BGM Mine and Forest Site Differences HKM Mine and Forest Site Differences

s__unidentified s__Russula_vinacea s__Tricholoma_albobrunneum s__Russula_emetica s__unidentified s__unidentified s__Piloderma_sphaerosporum g__Tricholoma g__Amphinema g__Russula s__Russula_cascadensis

g__Inocybe g__Piloderma s__unidentified s__Laccaria_angustilamella f__Tricholomataceae f__Atheliaceae s__unidentified s__Clavulina_cinerea g__Lactarius g__unidentified g__Laccaria f__Inocybaceae f__Russulaceae s__unidentified s__Lactarius_areolatus s__unidentified g__Clavulina f__Hydnangiaceae s__Amanita_novinupta o__Atheliales g__Russula o__Agaricales s__unidentified f__Russulaceae f__Clavulinaceae g__Trechispora g__Amanita s__Scleroderma_citrinum f__Amanitaceae o__Russulales o__Russulales g__Scleroderma f__Hydnodontaceae s__Amanita_flavoconia f__Sclerodermataceae o__Cantharellales o__Trechisporales c__Agaricomycetes s__unidentified g__unidentified o__Boletales s__unidentified o__unidentified f__unidentified g__unidentified g__Rhizopogon f__Rhizopogonaceae f__unidentified s__Rhizopogon_sp c__Agaricomycetes s__unidentified o__Thelephorales o__Sebacinales g__Tomentella o__unidentified

f__unidentified s__Thelephora_corticioides f__Boletaceae o__Thelephorales g__unidentified g__Thelephora g__Boletus f__Thelephoraceae s__unidentified f__Thelephoraceae o__Boletales s__unidentified g__unidentified f__unidentified Nodes g__Tomentella s__unidentified f__Pisolithaceae f__Inocybaceae Nodes s__Amanita_pachycolea g__Thelephora −1.000 1.0 o__Agaricales g__Inocybe s__Tomentella_sublilacina −1.000 1.00 g__unidentified g__Pisolithus f__Amanitaceae f__Rhizopogonaceae −0.667 1.5 s__Inocybe_tubarioides g__Amanita −0.667 1.78 s__Pisolithus_arhizus f__Tricholomataceae f__Hygrophoraceae s__Amanita_novinupta s__Thelephora_corticioides s__unidentified −0.333 3.0 f__Cortinariaceae −0.333 4.11 g__Tricholoma s__Amanita_flavoconia 0.000 5.5 0.000 8.00 g__Rhizopogon g__Hygrophorus s__unidentified 9.0 s__Tricholoma_aestuans 0.333 g__Cortinarius 0.333 13.40 s__Rhizopogon_evadens 0.667 13.5 s__unidentified s__Cortinarius_mucosus 0.667 20.40 Number of OTUs s__Rhizopogon_sp Number of OTUs s__unidentified 1.000 19.0 1.000 29.00 Log2 ratio median proportionsLog2 ratio Log2 ratio median proportionsLog2 ratio SH Mine and Forest Site Differences

s__Amanita_brunnescens s__unidentified

s__Mycena_epipterygia

g__Amanita g__Sebacina s__unidentified

g__Mycena s__unidentified

f__Amanitaceae f__Sebacinaceae g__Sistotrema f__Tricholomataceae

s__Tricholoma_equestreg__Tricholoma f__Cantharellales_fam_Incertae_sedis o__Agaricaleso__Sebacinales s__Basidiodendron_caesiocinereum

o__Cantharellales g__Basidiodendron

f__Exidiaceae o__Auriculariales

g__Luellia c__Agaricomycetes s__Luellia_recondita f__Hydnodontaceae

o__Trechisporales o__unidentified f__unidentified g__unidentifieds__unidentified o__Russulales

f__Trechisporales_fam_Incertae_sedis Nodes o__Boletales −1.000 1.00 g__Sistotremastrum f__Russulaceae −0.667 1.44 f__Boletaceae f__Sclerodermataceae s__Sistotremastrum_suecicum g__Tylopilus −0.333 2.78 s__Tylopilus_rubrobrunneus g__Lactarius 0.000 5.00 g__Scleroderma g__unidentified s__Lactarius_areolatus 0.333 8.11

0.667 12.10 s__Lactarius_imperceptus

s__Scleroderma_polyrhizum s__unidentified Number of OTUs 1.000 17.00

Log2 ratio median proportionsLog2 ratio Figure 8. Metacoder heat tree comparison of the abundant species in the class Agaricomycetes on mine sites (green) v.s. nearby forest (brown) for BGM (top left), HKM (top right), SH (bottom).

BGM Mine BGM Forest RGM Mine HKM Mine HKM Forest SH Mine

SH Forest

HKM Forest

s__unidentified s__Russula_cascadensis BGM Forest s__unidentified s__unidentified g__unidentified g__Russula

g__unidentified s__Russula_vinacea s__unidentified f__Russulaceae

f__unidentified g__Amphinema s__Clavulina_cinerea g__Clavulina f__Atheliaceae o__Russulales s__Cortinarius_mucosus o__unidentified f__Clavulinaceae

g__Cortinarius o__Atheliales o__Cantharellales s__unidentified g__unidentified f__Cortinariaceae f__unidentified s__Tricholoma_albobrunneum g__Tricholoma c__Agaricomyceteso__Sebacinales f__Tricholomataceae o__Agaricales g__Boletus f__Boletaceae s__unidentified

f__Hydnangiaceae o__Thelephorales g__Laccaria Nodes s__Laccaria_angustilamella o__Boletales f__Amanitaceae tions f__Pisolithaceae r −5.00 1.00 g__Pisolithus s__Pisolithus_arhizus −3.33 1.56

f__Sclerodermataceae TUs g__Amanita f__Thelephoraceae f__Rhizopogonaceae s__Amanita_novinupta −1.67 3.22 O 0.00 6.00 g__unidentified g__Rhizopogon s__Amanita_flavoconia g__Thelephora g__Scleroderma s__Rhizopogon_sp 1.67 9.89 3.33 14.90 s__unidentified s__Thelephora_corticioides s__unidentified Number of s__Scleroderma_polyrhizum atio median propo r 5.00 21.00 Log2

Figure 9. Metacoder matrix heat tree for pairwise comparison between sites. PCoA Mines

Root Soil 0.6 Site BGM_1 BGM_2 BGM_3 0.4 BGM_4 BGM_F HKM_1 HKM_2 HKM_3 HKM_4 0.2 HKM_F RGM_Pit RGM_Process SH_F Axis.2 [5.4%] Axis.2 SH_M 0.0 SH_P

Mine BGM HKM −0.2 RGM SH

−0.2 0.0 0.2 0.4 −0.2 0.0 0.2 0.4 Axis.1 [8.2%]

Figure 10. Principal Coordinate Analysis of the fungal communities on each mine. PCoA Mines

Root Soil

Site BGM_1 0.2 BGM_2 BGM_3 BGM_4 BGM_F HKM_1 HKM_2 HKM_3 0.0 HKM_4 RGM_Pit RGM_Process

Axis.2 [4.6%] Axis.2 SH_F SH_M SH_P

−0.2 Mine Forest Mine

−0.2 0.0 0.2 0.4 −0.2 0.0 0.2 0.4 Axis.1 [8.5%]

Figure 11. Principal Coordinate Analysis of the fungal communities on each mine. With Henry’s Knob Mine forest samples taken out.

0.25

0.00 Root Na Cu Zn Pb Site −0.25 Cd Equiv_water_pH BGM_1 P Base_saturation Mo BGM_2 Ni Cr BGM_3 BGM_4 Mn Ca Fe −0.50 Al BGM_F Mg LBC HKM_1 As HKM_2 CEC K C HKM_3 N HKM_4 RGM_Pit RGM_Process 0.25 SH_F PCoA 2 [4.6%] SH_M SH_P

0.00 Treatment Forest

Na Soil Mine Cu Zn Pb −0.25 Cd Equiv_water_pH P Base_saturation

Ni Cr Mo Mn Ca Fe −0.50 Al Mg LBC As CEC K N C −0.25 0.00 0.25 0.50 PCoA 1 [8.5%]

Figure 12. Principal Coordinate Analysis of the fungal communities on each mine, with soil property variables. Henry’s Knob Mine is taken out of the analysis as it has larger than expected distance from the other samples. Supplementary Information – Root Fungi Isolation and Greenhouse Bioassay Background With the fine root samples from BGM, we isolated a variety of root fungi isolates. In particular, three fungal species turns out to be very common isolates from roots collected on the disturbed sites: Acephala macrosclerotiorum, Rhizopogon fuscorubens, and Pisolithus tinctorius (Figure 13). The analysis of next-gen data confirms that the abundance of genera Acephala, Rhizopogon and Pisolithus in the root and soil samples from BGM.

Methods We set up a greenhouse bioassay to test 1) if the indigenous fungal community in the soil exerts influences the growth of pine hosts; 2) if artificially inoculating the pine seedlings growing on soil from disturbed sites will improve the growth of the seedlings; 3) if inoculation of a fungal mix will bring different effect from inoculation of a single species.

Soil from BGM_1 (more recovered site) and BGM_3 (more disturbed site) were used for the bioassay. Five replicates of Pinus taeda seeds were planted for each of combinations shown in Table 4. Three types of control treatments were applied: 1) sterile sand (no nutrient; no disturbance; no original spore bank); 2) sterile mine sites soil (nutrient and disturbance; no original spore bank); 3) mine site soil without inoculation (nutrient and disturbance; original spore bank).

Inocula were cultivated in perlite with pure strain culture of each species isolated from BGM mine sites. Four months after the initial seeding of P. taeda and three months after inoculating perlite substrate with fungal culture, a thin, plastic pistol was used to apply the perlite into the soil of each bioassay cone (Figure 14). For single-species inocula, each seeding receives about 12 ml of the inocula, applied in three separate times, in three different directions from the seeding. For mixed-species inocula, each seedling receives about 4ml of each type of inocula. The experiment will be harvested in summer 2019.

(A) (B) (C) Figure 13. Mycorrhizal fungi isolates from BGM: A) Acephala macrosclerotiorum; B) Rhizopogon fuscorubens; C) Pisolithus tinctorius.

(A) (B) Figure 14. A) Greenhouse bioassay to test the effect of indigenous fungal community; B) artificial inoculation on Pinus taeda growth.

Table 4. Treatments applied in the greenhouse bioassay. Treatment Replicates Inoculation Sand 5 no inoculation Sterile BGM_1 Soil 5 NA Sterile BGM_3 Soil 5 NA BGM_1 Soil 5 NA BGM_3 Soil 5 NA BGM_1 Soil 5 Acephala macrosclerotiorum BGM_1 Soil 5 Rhizopogon fuscorubens BGM_1 Soil 5 Pisolithus tinctorius BGM_1 Soil 5 Acephala x Rhizopogon x Pisolithus BGM_3 Soil 5 Acephala macrosclerotiorum BGM_3 Soil 5 Rhizopogon fuscorubens BGM_3 Soil 5 Pisolithus tinctorius BGM_3 Soil 5 Acephala x Rhizopogon x Pisolithus

Acknowledgements We would like to heartily thank Dr. Khalid Hameed for his help in the field and instructions for isolating the fungal strains. We would also like to thank Dr. Brian Looney and Yi-Hong Ke for their help in the field. Many thanks to Jim McLain for his accompany in the field and knowledge about Brewer Gold Mine, to Loften Carr from EPA for putting us in touch with the mine, to the managing staff of Henry’s Knob Mine for permission and accompany in the field, to Rodney Smith and Simon Donald from the US Forest Service for permission and directions for collection on the Russell Gold Mine. We are indebted to Abyy Maciejewski and Hui-Ling Liao for their contribution to the pilot study on Silver Hill Mine. This research is supported by the Duke University Dean’s Summer Research Fellowship, the Mycological Society of America Undergraduate Student Award, and North Carolina Independent Universities and Colleges (NCICU) Undergraduate Research Program.

Bibliography Adriaensen, K., Vangronsveld, J., & Colpaert, J. V. (2006). Zinc-tolerant Suillus bovinus improves growth of Zn-exposed Pinus sylvestris seedlings. Mycorrhiza, 16, 553–558. Amna, Ali, N., Sajid, M., Mukhtar, T., Kamran, M. A., Rafique, M., … Chaudhary, H. J. (2015). Differential effects of cadmium and chromium on growth, photosynthetic activity, and metal uptake of Linum usitatissimum in association with Glomus intraradices. Environ Monit Assess (2015), 187, 311. Andersen, K., Kirkegaard, R., Karst, S., & Albertsen, M. (2018). ampvis2: an R package to analyse and visualise 16S rRNA amplicon data. BioRxiv. Retrieved from https://www.biorxiv.org/content/10.1101/299537v1.abstract Carpenter, P. A. (1976). Metallic Mineral Deposits of the Carolina Slate Belt, North Carolina. North Carolina Department of Natural Resources and Community Development, Division of Land Resources, Geological Survey Section. Dickie, I. A., Martínez-García, L., Koele, N., Grelet, G.-A., Tylianakis, J. M., Peltzer, D. A., & Richardson, S. J. (2013). Mycorrhizas and mycorrhizal fungal communities throughout ecosystem development. Plant and Soil, 367(1–2), 11–39. Faye, A., Dalpé, Y., Ndung’u-Magiroi, K., Jefwa, J., Ndoye, I., Diouf, M., & Lesueur, D. (2013). Evaluation of commercial arbuscular mycorrhizal inoculants. Canadian Journal of Plant Sciences, 93, 1201–1208. Gaur, A., & Adholeya, A. (2004). Prospects of arbuscular mycorrhizal fungi in phytoremediation of heavy metal contaminated soils. Current Science, 86(4), 528–534. Grand, L. F. (1976). Distribution, Plant Associates and Variation in Basidiocarps of Pisolithus tinctorius in the United States. Mycologia, 68(3), 672–678. Huang, J., Nara, K., Lian, C., Zong, K., Peng, K., Xue, S., & Shen, Z. (2012). Ectomycorrhizal fungal communities associated with Masson pine (Pinus massoniana Lamb.) in Pb–Zn mine sites of central south China. Mycorrhiza, 22, 589–602. Hudson Institute of Mineralogy. (2019, April 22). Silver Hill Mine, Silver Hill, Cid District, Carolina Slate Belt, Davidson Co., North Carolina, USA. Retrieved April 22, 2019, from https://www.mindat.org/loc-7918.html Kałucka, I., & Jagodziński, A. (2016). Successional traits of ectomycorrhizal fungi in forest reclamation after surface mining and agricultural disturbances: A review. Dendrobiology, 76, 91–104. Lampky, J. R., & Peterson, J. E. (1963). Pisolithus tinctorius associated with pines in Missouri. Mycologia, 55(5), 675–678. Marrs, R. H. (2016). Ecological restoration: Soil microbes call the shots. Ecological Restoration, 2, 16117. McMurdie, P. J., & Holmes, S. (2013). An R Package for Reproducible Interactive Analysis and Graphics of Microbiome Census Data. PLoS ONE, 8(4), e61217. Medve, R. J., & Gill, S. M. (1982). Distribution and Ecology of Pisolithus tinctorius on Bituminous Stripmine Spoils in Western Pennsylvania. Bulletin of the Torrey Botanical Club, 109(1), 35–38. Mummey, D. L., Antunes, P. M., & Rillig, M. (2009). Arbuscular mycorrhizal fungi pre- inoculant identity determines community composition in roots. Soil Biology & Biochemistry, 41, 1173–1179. Münzenberger, B., Golldack, J., Ullrich, A., Schmincke, B., & Hüttl, R. F. (2004). Abundance, diversity, and vitality of mycorrhizae of Scots pine (Pinus sylvestris L.) in lignite recultivation sites. Mycorrhiza, 14, 193–202. Nguyen, N. H., Song, Z., Bates, S. T., Branco, S., Tedersoo, L., Menke, J., … Kennedy, P. G. (2016). FUNGuild: an open annotation tool for parsing fungal community datasets by ecological guild. Fungal Ecology, 20, 241–248. Op De Beeck, M., Ruytinx, J., Smits, M. M., Vangronsveld, J., Colpaert, J. V., & Rineau, F. (2015). Belowground fungal communities in pioneer Scots pine stands growing on heavy metal polluted and non-polluted soils. Soil Biology & Biochemistry, 86, 58–66. Perry, D. A., Molina, R., & Amaranthus, M. P. (1987). Mycorrhizae, mycorrhizospheres, and reforestation: current knowledge and research needs. Canadian Journal of Forest Research, 17, 929–940. PHC Reclamation, Inc. (n.d.). Rice, P., Longden, I., & Bleasby, A. (2000). EMBOSS: The European Molecular Biology Open Software Suite. Rincón, A., de Felipe, M. R., & Fernández-Pascual, M. (2017). Inoculation of Pinus halepensis Mill. with selected ectomycorrhizal fungi improves seedling establishment 2 years after planting in a degraded gypsum soil. Mycorrhiza, 18. Shi, Y., Xie, H., Cao, L., Zhang, R., Xu, Z., Wang, Z., & Deng, Z. (2017). Effects of Cd- and Pb- resistant endophytic fungi on growth and phytoextraction of Brassica napus in metal- contaminated soils. Environ Sci Pollut Res, 24, 417–426. Singh, A. N., Raghubanshi, A. S., & Singh, J. S. (2002). Plantations as a tool for mine spoil restoration. Current Science, 82(12), 1436–1441. Staudenrausch, S., Kaldorf, M., Renker, C., Luis, P., & Buscot, F. (2005). Diversity of the ectomycorrhiza community at a uranium mining heap. Biology and Fertility of Soils, 41, 439–446. Steinfield, D., Amaranthus, M., & Cazares, E. (2003). Survival of Ponderosa Pine (Pinus Ponderosa Dougl. ex laws.) Seedlings Outplanted with Rhizopogon Mycorrhizae Inoculated with Spores at the Nursery. Journal of Arboriculture, 29(4), 197–207. Van Tichelen, K. K., Colpaert, J. V., & Vangronsveld, J. (2001). Ectomycorrhizal protection of Pinus sylvestris against copper toxicity. New Phytologist, 150. Wang, F. (2017). Occurrence of arbuscular mycorrhizal fungi in mining-impacted sites and their contribution to ecological restoration: Mechanisms and applications. Critical Reviews in Environmental Science and Technology, 0(0), 1–57.