Survey of macrofungal diversity and analysis of edaphic factors influencing the fungal community of church forests in Dry Afromontane areas of Northern Ethiopia

Forest Ecology and Management 496 (2021) 119391 Available online 2 June 2021 0378-1127/© 2021 The Author(s). Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/). Survey of macrofungal diversity and analysis of edaphic factors influencing the fungal community of church forests in Dry Afromontane areas of Northern Ethiopia Demelash Alem a , b , Tatek Dejene a , b , Juan Andr´ es Oria-de-Rueda a , Pablo Martín-Pinto a , * a Sustainable Forest Management Research Institute, University of Valladolid, Avda. Madrid 44, 34071 Palencia, Spain b Ethiopian Environment and Forest Research Institute (EEFRI), P. O. Box 30708 Code 1000, Addis Ababa, Ethiopia ARTICLE INFO Keywords: Conservation Edaphic variables Fragmented forests Macrofungi Church forests Sporocarps ABSTRACT The Dry Afromontane forests in Northern Ethiopia have been cleared for agriculture and reduced to small and isolated fragments. Most of these forests are located around church territories and are they called church forests. The church forests are known to be biodiversity islands and provide key ecosystem services to local communities. However, to date, the fungal resources of these forests have not been assessed and, therefore, the contribution of fungi to their conservation value is unknown. In 2019, we investigated the fungal diversity of three Dry Afromontane church forests. In each forest, we established nine permanent plots (2 m ×50 m), which were surveyed weekly during the rainy season to quantify the fungal diversity and sporocarp production levels. Explanatory variables were also analyzed to determine their relationship with macrofungal species composition. We collected 13,736 sporocarps corresponding to 188 taxa. Of these, 81% were saprotrophic and 14% were ectomycorrhizal. Sixty-eight species were edible, including economically valuable species such as Tricholoma and Termitomyces. This suggests that these fragmented forest systems could be managed to provide valuable non-timber forest products such as mushrooms and socioeconomic benefits for local communities. Although many species were present in all three forests, some were only found in one forest, highlighting the importance of conserving individual forests. The correlation of the Shannon diversity indices of the two communities showed a positive trend in spite of the lack of correlation between their richness. Macrofungal communities as a whole were influenced by edaphic, spatial and climate variables. This study indicates that church forests support a wide diversity of fungi, including potentially novel fungal species, and highlights the need for forest managers to consider the importance of fungi in forest ecosystem management and to provide habitats that will maintain fungal diversity and sporocarp production when planning conservation strategies. 1. Introduction The Ethiopian highlands constitute large parts of the Afromontane regions in Africa (Aynekulu et al., 2016; Nyssen et al., 2014). These highlands are dominated by Dry Afromontane forests and are mainly found in the Northern part of Ethiopia (Eshete et al., 2011; Friis et al., 2010a,b). Dry Afromontane forests are rich in biodiversity and are dominated by pioneers, shrubs, and high-quality trees (Abiyu et al., 2018; Lemenih et al., 2011) that are able to grow at high altitudes (Friis et al., 2010a,b). The main tree species that constitute the Dry Afromontane forests include Juniperus procera, Podocarpus falcatus, Hagenia abyssinica and Olea africana (Kassa et al., 2009). These trees serve as a vital source of timber to the country (Kassa et al., 2009) and thus an indication of a need for the sustainable management of these forests. The Dry Afromontane forests also produce several Non-Timber Forest Products (NTFPs) such as wild edible fruits and medicinal plants that are vital for the socioeconomics of the local communities (Shumi, 2009). Furthermore, edible mushrooms from the forest systems have been utilized as important sources of food and medicine by rural communities for their livelihoods in few specific regions in Ethiopia (Abate, 2008). However, high levels of historical human landscape alteration and landuse pressure have resulted in widespread deforestation and the degradation of forest land (Aerts et al., 2016; Aynekulu et al., 2016; Darbyshire et al., 2003; Nyssen et al., 2014). The ever-increasing demand for * Corresponding author. E-mail addresses: [email protected] (J.A. Oria-de-Rueda), [email protected] (P. Martín-Pinto). Contents lists available at ScienceDirect Forest Ecology and Management journal homepage: www.elsevier.com/locate/foreco https://doi.org/10.1016/j.foreco.2021.119391 Received 14 April 2021; Received in revised form 19 May 2021; Accepted 22 May 2021 Forest Ecology and Management 496 (2021) 119391 2 wood products as well as crop and grazing land expansion, stimulated by rapid population and livestock growth are also factors aggravating the degradation of the Dry Afromontane forests in the country (Bekele and Lemenih, 2008). As a result of the many physical and biological changes to the Dry Afromontane forests in Northern Ethiopia, natural forests have been reduced to small and isolated fragments, most of which belong to the church or are located around church forest territories (Aerts et al., 2016; Aynekulu et al., 2016; Wassie et al., 2009). These forest fragments have survived because of the cultural or religious values held by local communities, which have contributed to the conservation of their biodiversity. Owing to the small size of these forest fragments, there are variations in their biodiversity between forest fragments (Lemenih et al., 2011; Wassie et al., 2005) and probably of their fungal communities. Fungal communities are an important component of forest ecosystems and have a broad range of functions (Song et al., 2019; Tedersoo et al., 2014a; Wagg et al., 2014). As decomposers, they are important for the degradation of organic matter and play a vital role in nutrient cycling (Chen et al., 2019; Ferris et al., 2000). Mycorrhizal fungi form symbiotic associations with higher plants, facilitating plant uptake of water and nutrients (Egli, 2011; Hall et al., 2003; Tedersoo et al., 2020). Fungi can also be used as bioindicators to assess the quality of forests (Egli, 2011; Van Bruggen and Semenov, 2000). In addition to their ecological roles, fungi have been used by humans for thousands of years in different ways (Boa, 2004; Oria-De-Rueda et al., 2008) and are sold in markets worldwide, providing an important source of rural income (Boa, 2004; Pettenella et al., 2007). Indeed, in some cases, forest fungi provide significant complimentary benefits to forest managers (Bonet et al., 2014; Martín-Pinto et al., 2006). Fungi also provide food and habitats for other organisms and, therefore, interactions between fungi and other organisms in forest systems cannot be overlooked (Jonsell and Nordlander, 2000). Pathogenic fungi also impact ecosystems, mainly by acting as natural population regulators, thereby influencing productivity, species diversity, and composition (Ruiz-Almenara et al., 2019). For all these reasons, fungi are considered a strategic component in the conservation and management of forest systems (Bonet et al., 2014). Today, biodiverse remnants of the Dry Afromontane forests survive in the agricultural landscape and church areas as forest islands (Aynekulu et al., 2016). Several studies have evaluated the conservation value of these fragmented forests in the Northern landscapes of Ethiopia (Aerts et al., 2016; Aynekulu et al., 2016; Wassie et al., 2010, 2005; Nyssen et al., 2014). However, the ecology and conservation status of the macrofungi in these isolated and fragmented forest systems is unknown. Consequently, the fungal taxonomy and ecology are very poorly described and, hence, fungi are neglected when decisions need to be made regarding forest management and conservation actions. Recently, there has been an interest in surveying fungi in particular habitats (Alem et al., 2020), to describe and predict the extent of their diversity on a larger scale (Danielsen et al., 2005; Peay, 2014). This information is important to enable the integration of fragmented forests into global biodiversity conservation strategies (Hundera et al., 2013; Aerts et al., 2016; Aynekulu et al., 2016) and to understand what actions are required to conserve fragmented forests and their biological components, including fungi, which are known for their exceptionally high diversity levels (Burgess et al., 2006). The practice of using plant communities as surrogates to predict fungal diversity has been reported by previous studies (McMullan-Fisher et al., 2010; Rudolf et al., 2013). Fungal communities have also been associated with essential ecosystem parameters such as edaphic variables (Chen et al., 2018), as well as other relevant drivers of fungal richness at a global level such as climate (Tedersoo et al., 2014b). Furthermore, fungal diversity is considered to reflect niche diversity given that reducing niche similarity drives species assemblage (Silvertown, 2004). The Dry Afromontane church forests in Ethiopia are reported to have high levels of plant species diversity (Mokria et al., 2015; Tsegaye et al., 2010). However, there is no evidence that the high level of plant diversity in the church Dry Afromontane forests system anticipates correspondingly high macrofungal diversity. In addition, as yet, the environmental variables that govern fungal communities in these fragmented forest systems have not been identified given that these forests vary in their status (size, density, species composition etc.), topography and altitude (Bongers and Tenngkeit, 2010). Thus, investigating the fungal community composition and how this community changes across sites in fragmented forests should help us to understand different aspects of fungal interaction within these systems and their function in the ecosystem (Genevieve et al., 2019). This information may also be a means to understand how to improve natural fungal richness and sporocarp production and also help us to facilitate the conservation of economically and ecologically important macrofungal species in these fragmented forest systems. This study is the first attempt to provide baseline information about macrofungi assemblage, diversity, and sporocarp production in Dry Afromontane church forests with priority status in Northern Ethiopia. Priority forests are those that have been designated as reserves to give them additional protection. The information generated should help to guide management and conservation strategies for these priority forests and supplement our knowledge of macrofungal species in Ethiopia. Furthermore, determining whether edible mushrooms are produced in these forests could provide an opportunity for harvesting edible mushrooms for either subsistence or commercial use. Despite fragmentation, the forests in the study areas are suggested to be relatively rich in plant species (Aerts et al., 2016; Wassie et al., 2010). On average, there are 25 vascular tree species per forest patch (Aerts et al., 2016). The tree species composition of these forests also varied with their status, topography and altitude (Bongers and Tenngkeit, 2010), with a wide distribution over the landscapes (Aerts et al., 2016). Given that fungal diversity is related positively to plant richness (Tedersoo et al., 2014b), we hypothesized that the fungal diversity of the study plots in the church forests would be high in terms of total fungal species. In addition, the edaphic conditions, vascular plant richness and diversity would follow given the differences in climate and topography among the different fragmented forests, we also hypothesized that the composition of macrofungal communities would differ among the studied forests, resulting in an overall higher richness value for the study sites because fungi will be driven mainly by vegetation condition and site conditions such as soil fertility conditions (Casta˜ no et al., 2018; Vaˇ sutov´ a et al., 2017). Thus, our specific aims were to study three church forests in Dry Afromontane areas of Northern Ethiopia: (1) to describe fungal species richness, diversity, and sporocarp production; (2) to correlate the macrofungal and plant diversity of the three church forests; and (3) to determine whether the macrofungal community composition was governed by the soil fertility status of the three forest sites. 2. Materials and methods 2.1. Description of the study areas The study was conducted in three church forests located in three different districts of the Amhara region, namely the Taragedam forest located in Libokemkem Woreda, the Alemsaga forest located in Farta Woreda, and the Banja forest located in Banja Woreda (Fig. 1). These forests are fragments of the remnant Dry Afromontane forests in Northern Ethiopia (Gebeyehu et al., 2019; Masresha et al., 2015; Zegeye et al., 2011). The Taragedam and Banja forests were designated as reserves in 1979 (Zegeye et al., 2011) and 1994 (Abere et al., 2017), respectively, to prevent any kind of encroachments. The Alemsaga forest was designated as a priority forest in 1978 to serve as a seed source, to conserve the remnant natural forest, and to rehabilitate the degraded area in Northern part of the country (Masresha et al., 2015). Comprehensive descriptions of the forests are provided in Table 1. Within each of the forests, plots were established systematically about 500 m apart in 2019. The plots were laid out randomly in the forests to avoid D. Alem et al. Forest Ecology and Management 496 (2021) 119391 3 confounding spatial effects inherent to such a plot-based design (Hiiesalu et al., 2017; Rudolph et al., 2018). The plots were analyzed as independent samples as suggested by Ruiz-Almenara et al.(2019). However, the present study provides a starting place in broadening management objectives for NTFPs in the Dry Afromontane church forests. Thus, the result should be considered as a case study and as a preliminary indication and conclusions regarding other similar studies need to be taken with caution. 2.2. Sporocarp sampling In total, 27 sample plots were established, nine in each of the three church forests, as described in Gassibe et al. (2011) and Hern´ andez- Rodríguez et al. (2013). Each plot was rectangular in shape (2 m ×50 m) and covered an area of 100 m 2 . Within each of the selected church forests, we studied three different blocks including three plots per block. The plots were established about a minimum distance of 500 m apart. All fungal fruit bodies found were harvested weekly during the major rainy season in July and August of 2019. Fresh weight measurements were taken in situ using a digital sensitive balance (SF-400) to determine fruit body production in kilograms per hectare per year. The number of sporocarps of each species in each plot was also recorded. Specimens were photographed in the field and their morphological features and ecological characteristics were noted to facilitate taxonomic identification processes in the laboratory (Adeniyi et al., 2018). Specimens of each macrofungus were taken to the laboratory and dried to preserve as herbaria specimens, and then used for morphological species identification. 2.3. Species identification In the laboratory, the morphological features of the fruit bodies were examined using appropriate monographs, including Antonin (2007), Hama et al. (2010), Heinemann (1956), Hjortstam and Ryvarden (1996), Morris (1990), Pegler (1968, 1969, 1977), Rammeloo and Walleyn (1993), and Singer (1965), to determine the genus and species of the macrofungal specimens. Up-to-date fungal species names and authors’ names were obtained from the Mycobank database (http://myc obank.org). Ecological functions at the genus level were identified using a FUNGuild (www.funguild.org) search and provided in Table 2. In addition, the edibility of the fruiting bodies collected from the study sites was assessed following the criteria used by Bonet et al. (2004). Species described in the literature as both non-edible and edible in the literature were classified as non-edible. Species described in the literature as having doubtful edibility were classified as non-edible. Only species classified as edible by a large majority of the literature consulted were classified as edible fungi (E). 2.4. Soil sampling and analysis To relate macrofungal composition to edaphic variables, soil samples were collected from each of the sample plots established in the three forests. Composite samples were collected by grouping each plot into relatively homogeneous subsamples. After clearing and removing plant matter and debris from the soil surface, five soil cores were extracted from the center and the four corners of each plot using an auger (2 cm radius, 20 cm deep and 250 cm 3 ). Subsamples collected from each plot were mixed thoroughly and a composite sample of approximately 500 g was placed in a plastic bag for analysis. Soil samples were dried under a shed until a constant weight was obtained and ground to <2 mm sieved soil is used in the analysis. The soil pH and electrical conductivity were determined by analyzing a soil:water (1:2.5) suspension and the supernatant from the same suspension with the aid of a pH meter and an electrical conductivity meter, respectively (Reeuwijk, 2002). The organic carbon content of the soil was determined using wet digestion (Walkley and Black, 1934). The Kjeldahl procedure was used to determine the total N content in soils (Kim et al., 2005). Sodium bicarbonate (0.5 M NaHCO3) was used as an extraction solution to determine the available phosphorus (P) (Kim, 1996). Available sodium (Na), potassium (K), calcium (Ca) and magnesium (Mg) were also determined. To assess soil particle size we used a hydrometer (Bouyoucos, 1951) and sodium hexametaphosphate (Calgon solution) was used as a dispersing agent. After calculating the proportions of sand, silt, and clay, the soil was assigned a textural class name using ASTM free software, Version 4, Available: http://www.astm.org. The results of the soil analysis are provided in Table 1. The soil analysis was conducted by the Amhara Design and Supervision Works Enterprise at Bahir Dar, Ethiopia. Fig. 1. Map of the Amhara region in Northern Ethiopia showing the location of the three church forests in which the study plots were located. D. Alem et al. Forest Ecology and Management 496 (2021) 119391 4 2.5. Vegetation sampling To relate the vegetation characteristics to the macrofungal richness and diversity, vegetation inventories were conducted in the plots established for macrofungal species sampling as described above. Vascular plants identified in each plots were recorded using their vernacular names. For those species difficult to identify their scientific name in the field, specimens were collected and their taxonomic identification was conducted using published volume of the flora of Ethiopia and Eritrea (Hedberg and Sue, 1989). Large trees growing outside the plots were included in the survey if their crowns overhung the plots because tree crown projection areas can affect macrofungal occurrence (Collins et al., 2018). Furthermore, large trees create their own microhabitat and develop a large root system, providing more space for fungal associations (Sch¨ on et al., 2018). Then, the vascular plant species richness and diversity parameters were determined (Table 3). Plant parameters and their correlations were also used for further interpretation of macrofungal pattern from each study areas. The mycorrhizal status of the vascular tree species found in each of the studied plots were checked using freely accessible databases (Soudzilovskaia et al., 2020) and the data is provided (Table S1). 2.6. Statistical analysis Data were transformed when needed to achieve the parametric criteria of normality and homoscedasticity. The macrofungi data were normalized by rarefying the abundance data to the smallest number of macrofungi per plot. Also, the data from soil variables were scaled using base R and used for subsequent statistical analyses. Shannon’s H′diversity index, H′=–Σpi (lnpi) (Shannon and Weaver, 1949), was estimated for each forest, where pi indicates the relative abundance of the species (Kent and Coker, 1993). Simpson’s diversity, D =1 −Σ(pi2), where pi is the importance probability in element i; and the evenness, J =H′/H′max, where H′is the number derived from the Shannon diversity index and the H′max is the maximum possible value of H′were also calculated (Magurran, 1988). In addition to species richness values, macrofungi biomass production levels in each forest were estimated and converted in to Kg bases. All diversity measures for macrofungi and vascular plants were analyzed using the Biodiversity R package (Kindt and Coe, 2005) in R version 4.0.3 (R Core Team, 2020). The difference in the soil, vegetation and sporocarps variables across forests were assessed by Linear Mixed Effects models (LME, Pinheiro et al., 2016), where block (a set of plots in a same site in each forest) was defined as random and forest was defined as fixed factor. The LME used to prevent the false positive associations due relatedness structure in the sampling. Tukey Test was later used to check significant differences (p ≤0.05) between forests when needed. Species accumulation curves were constructed to compare the rate at which new fungal species were found in the three forests and to provide an estimate of macrofungal species richness. Curves were generated using a sample-based estimator of EstimateS Version 9 (Colwell, 2013). The number of fungal species collected during each weekly visit to a plot within a forest constituted the sample. Curves were generated based on the total of the weekly sampling datasets. A R´ enyi diversity profile (T´ othm´ er´ esz, 1995) was also used to depict the diversity curves of the three church forests. When parameter alpha =0, this function gives the total species number and when alpha =1, this gives an index proportional to the Shannon index. The relationship of macrofungal composition with the edaphic, climate and location parameters was visualized using non-metric multidimensional scaling (NMDS), based on absence and presence species data matrix and environmental scaled data. A permutation-based nonparametric MANOVA (PerMANOVA) (Anderson, 2001) using Bray–Curtis distance was conducted to analyze differences in macrofungal communities across forests. The isolines of the elevation also plotted on the NMDS ordinations using the ordisurf function. The correlation of NMDS axes scores with explanatory variables was assessed using envfit function in R. To test the influence of categories of the edaphic, climate and location variables on the fungal community, we used Mantel Test (Bray-Curtis distance) on total species matrix and scaled environmental parameters. Also, an analysis of similarity percentages (SIMPER; Clarke, 1993) was performed to identify macrofungal species that were most responsible for the observed patterns and was also used to determine the percentage contribution of macrofungal species to significant dissimilarities between the three forests (Parravicini et al., 2010). The SIMPER analysis was performed using the sim function of the Vegan package in R (R Core Team, 2020). 3. Results 3.1. Macrofungal richness and diversity In total, 13,736 sporocarps were collected from the three church Table 1 Characteristics of the study sites and selected edaphic properties. Descriptions Forests Taragedam Alemsaga Banja Geographical location 12◦06′–12◦07′N 37◦46′– 37◦47′E 11◦54′–11◦56′N 37◦55′–37◦57′E 10◦57′–11◦03′N 36◦39′– 36◦48′E Altitude range (m asl) 2142–2484 2180–2470 1870–2570 Mean annual precipitation (mm) 1098 1926 1884.3 Mean annual temperature (◦C) 19.5 15.8 18.7 Forest area (ha) 875 814 897 Density of trees ha −1 48.11 17.19 43.13 Sand (%) 58.89 ±2.93b 51.78 ±2.99b 68.67 ±2.21a Silt (%) 28.44 ±2.38a 32.44 ±2.13a 20.00 ±1.76b Clay (%) 12.67 ±1.37a 15.78 ±1.93a 11.33 ±1.33a pH H2O 1:2.5 7.04 ±7.03a 5.85 ±6.59b 5.60 ±6.24c EC (dS/m) 0.43 ±0.05b 0.28 ±0.03b 0.81 ±0.14a Ex.Ca (cmol (+)/kg) 13.95 ±0.60a 9.19 ±0.52b 13.55 ±0.87a Ex.Mg (cmol (+)/kg) 6.16 ±0.10a 4.58 ±0.15c 5.54 ±0.20b Ex.Na (cmol (+)/kg) 1.95 ±0.05a 2.05 ±0.10a 1.82 ±0.12a Ex.K (cmol (+)/kg) 0.73 ±0.06a 0.61 ±0.04a 0.77 ±0.06a CEC (cmol (+)/kg) 47.21 ±1.36a 34.89 ±0.92b 44.51 ±1.96a Organic matter (%) 4.46(0.60)a 3.35(1.34)b 4.87(0.10)a Nitrogen (%) 0.23 ±0.01a 0.17 ±0.02b 0.26 ±0.01a P (ppm) 17.18 ±5.72a 7.8 ±0.73b 17.64 ±6.05a Dominant species in each plots Maytenus obscura, Carissa edulis, Olea sp. Acacia abyssinica, Buddleja polystachya, Acacia nilotica Albizia gummifera, Prunus africana, Brucea antidysenterica References Gedefaw and Soromessa (2014), Zegeye et al. (2011), Zerihun et al. (2013) Birhane et al. (2017), Masresha et al. (2015), Wubet et al. (2004) Abere et al. (2017) Note: Values shown are means; standard errors of the means are indicated in parentheses. Values with different lowercase letters are significantly different (p <0.05). The mean annual precipitation and mean annual temperature are given based on nearby stations data of each study area by the year 2019. Abbreviations: EC, electrical conductivity; CEC, cation exchange capacity; m, meter; mm, millimeter; asl, above sea level. The references listed are related to the climatic and geographical descriptions of the study areas. D. Alem et al. Forest Ecology and Management 496 (2021) 119391 5 Table 2 Fungal sporocarps collected in July and August in three church forests in Northern Ethiopia. Taxa Order Family T A B E LM Agaricus augustus Fr. Agaricales Agaricaceae x E SS Agaricus bitorquis (Qu´ el.) Sacc. Agaricales Agaricaceae x E SS Agaricus campestris L. Agaricales Agaricaceae x x x E SS Agaricus cupreobrunneus (Jul.Sch¨ affer & Steer ex F.H.Møller) Pil´ at Agaricales Agaricaceae x x E SS Agaricus megalosporus J. Chen, R.L. Zhao, Karun. & K.D. Hyde Agaricales Agaricaceae x x x E SS Agaricus moelleri Wasser Agaricales Agaricaceae x x E SS Agaricus murinaceus Bull. Agaricales Agaricaceae x E SS Amanita vaginata (Bull.) Lam. Agaricales Amanitaceae x x EM Amanita sp. Pers. Agaricales Amanitaceae x x E EM Amanita verna (Bull.) Lam. Agaricales Amanitaceae x x x E EM Ampulloclitocybe clavipes (Pers.) Redhead, Lutzoni, Moncalvo & Vilgalys Agaricales Tricholomataceae x x E LS Artomyces pyxidatus (Pers.) Jülich Russulales Amylostereaceae x x x WS Auricularia auricula-judae (Bull.) Qu´ el. Auriculariales Auriculariaceae x E WS Bisporella citrina (Batsch) Korf & S.E.Carp. Helotiales Helotiaceae x WS Bjerkandera adusta (Willd.) P.Karst. Polyporales Meruliaceae x x WS Bolbitius sp. Fr. Agaricales Bolbitiaceae x x DS Bovista aestivalis (Bonord.) Demoulin Agaricales Agaricaceae x x SS Bovista plumbea Pers. Agaricales Agaricaceae x SS Calvatia cyathiformis (Bosc) Morgan. Agaricales Agaricaceae x x E SS Calvatia gigantea (Batsch) Lloyd Agaricales Agaricaceae x E SS Calvatia sp. Fr. Agaricales Agaricaceae x E SS Cantharellula umbonata (J.F.Gmel.) Singer Agaricales Tricholomataceae x x LS Cantharellus cinnabarinus (Schwein.) Schwein. Cantharellales Hydnaceae x E EM Chlorophyllum molybdites (G. Mey.) Massee Agaricales Agaricaceae x x x E LS Chlorophyllum rhacodes (Vittad.) Vellinga Agaricales Agaricaceae x x x E LS Clavaria falcata Pers. Agaricales Clavariaceae x SS Climacodon septentrionalis (Fr.) P. Karst. Polyporales Phanerochaetaceae x WS Clitocybe carolinensis H.E. Bigelow & Hesler Agaricales Tricholomataceae x x E LS Clitocybe cistophila Bon & Contu Agaricales Tricholomataceae x E LS Clitocybe foetens Melot. Agaricales Tricholomataceae x x x E LS Clitocybe fragrans (With.) P.Kumm. Agaricales Tricholomataceae x x x E LS Clitocybe geotropa (Bull.ex DC.) Qu´ el Agaricales Tricholomataceae x E LS Clitopilus hobsonii (Berk. & Broome) P.D. Orton Agaricales Entolomataceae x x x LS Conocybe apala (Fr.) Arnolds Agaricales Bolbitiaceae x SS Conocybe aurea (Jul.Sch¨ aff.) Hongo Agaricales Bolbitiaceae x x SS Conocybe dumetorum (Velen.) Svrcek Agaricales Bolbitiaceae x x SS Conocybe tenera (Schaeff.) Fayod Agaricales Bolbitiaceae x x x SS Conocybe velutipes (Velen.) Hauskn. & Svrcek Agaricales Bolbitiaceae x x x SS Coprinellus disseminatus (Pers.) J.E.Lange Agaricales Psathyrellaceae x x SS Coprinellus micaceus (Bull.) Vilgalys, Hopple & Jacq. Johnson Agaricales Psathyrellaceae x x x SS Coprinopsis sp. P. Karst. Agaricales Coprinaceae x x SS Coprinus comatus (O.F.Müll.) Pers. Agaricales Coprinaceae x x x E DS Coprinus lagopus (Fr.) Fr. Agaricales Coprinaceae x x DS Coprinus micaceus (Bull.) Fr. Agaricales Coprinaceae x E DS Coprinus niveus (Pers.) Fr. Agaricales Coprinaceae x x x E DS Cortinarius rubellus Cooke Agaricales Cortinariaceae x x EM Craterellus ignicolor (R.H. Petersen) Dahlman, Danell & Spatafora Cantharellales Hydnaceae x E EM Crepidotus applanatus (Pers.) P. Kumm. Agaricales Inocybaceae x x x E WS Crepidotus mollis (Schaeff.) Staude Agaricales Inocybaceae x x x E WS Crucibulum laeve (Huds.) Kambly Agaricales Agaricaceae x LS Cystodermella granulosa (Batsch) Harmaja Agaricales Agaricaceae x x LS Dacrymyces palmatus (Schwein.) Burt Dacrymycetales Dacrymycetaceae x WS Daedaleopsis confragosa (Bolton) J.Schr¨ ot. Polyporales Polyporaceae x x WS Daldinia concentrica (Bolton) Ces. & De Not. Xylariales Hypoxylaceae x WS Deconica montana (Pers.) P.D. Orton Agaricales Strophariaceae x x x LS Entoloma asprellum (Fr.) Fayod. Agaricales Entolomataceae x x x SS Entoloma olivaceohebes Noordel. & Hauskn. Agaricales Entolomataceae x x SS Entoloma poliopus (Romagn.) Noordel. Agaricales Entolomataceae x x SS Entoloma sp. Fr. ex P. Kumm. Agaricales Entolomataceae x x SS Entoloma undatum (Gillet) M.M. Moser Agaricales Entolomataceae x x SS Favolaschia calocera R. Heim Agaricales Marasmiaceae x WS Galerina badipes (Pers.) Kühner. Agaricales Strophariaceae x x x WS Geastrum triplex Jungh. Geastrales Geastraceae x x x LS Geoglossum sp. Pers. Geoglossales Geoglossaceae x x SS Gymnopilus sp1. P.Karst. Agaricales Omphalotaceae x x WS Gymnopilus sp2. P.Karst. Agaricales Omphalotaceae x x WS Gymnopilus sp3. P.Karst. Agaricales Omphalotaceae x WS Gymnopus dryophilus (Bull.) Murrill Agaricales Omphalotaceae x x x LS Gymnopus luxurians (Peck) Murrill Agaricales Omphalotaceae x LS Gymnopus putillus (Fr.) Antonín, Halling & Noordel. Agaricales Omphalotaceae x x LS Hebeloma eburneum Malençon Agaricales Strophariaceae x EM Hemimycena delectabilis (Peck) Singer. Agaricales Tricholomataceae x x x LS Hexagonia tenuis (Hook.) Fr. Polyporales Polyporaceae x x x WS Hohenbuehelia petalodes (Bull.) Schulzer. Agaricales Pleurotaceae x WS (continued on next page) D. Alem et al. Forest Ecology and Management 496 (2021) 119391 6 Table 2 (continued) Taxa Order Family T A B E LM Hygrocybe chlorophana (Fr.) Wünsche Agaricales Hygrophoraceae x x x E SS Hygrocybe chlorophana var. aurantiaca Bon. Agaricales Hygrophoraceae x E SS Hygrophoropsis aurantiaca (Wulfen) Maire Boletales Hygrophoropsidaceae x x x LS Hygrophorus hypothejus Fr. (Fr.) Agaricales Hygrophoraceae x x x E EM Hymenagaricus sp1. Heinem. Agaricales Agaricaceae x E SS Hymenagaricus sp2. Heinem. Agaricales Agaricaceae x SS Inocybe viridiumbonata Pegler Agaricales Inocybaceae x EM Laccaria glabripes McNabb. Agaricales Hydnangiaceae x EM Laccaria laccata (Scop.) Cooke Agaricales Hydnangiaceae x EM Laetiporus sulphureus (Bull.) Murrill Polyporales Fomitopsidaceae x x x E PP Lentinellus cochleatus (Pers.) P. Karst. Russulales Auriscalpiaceae x x E WS Lepiota cristata (Bolton) P.Kumm. Agaricales Agaricaceae x LS Lepiota ermine (Fr.) P.Kumm. Agaricales Agaricaceae x x LS Lepiota himalayensis Khalid & Razaq Agaricales Agaricaceae x x LS Lepiota sp1. (Pers.) Gray Agaricales Agaricaceae x LS Lepiota sp2. (Pers.) Gray Agaricales Agaricaceae x LS Lepiota sp3. (Pers.) Gray Agaricales Agaricaceae x x LS Leptonia lampropus (Fr.) Qu´ el. Agaricales Entolomataceae x x x SS Leucoagaricus americanus (Peck) Vellinga. Agaricales Agaricaceae x x x E SS Leucoagaricus purpureolilacinus Huijsman Agaricales Agaricaceae x x x E SS Leucoagaricus sp1. Locq. ex Singer Agaricales Agaricaceae x x E SS Leucoagaricus sp2. Locq. ex Singer Agaricales Agaricaceae x SS Leucocoprinus cepaestipes (Sowerby) Pat. Agaricales Agaricaceae x x SS Leucocoprinus fragilissimus (Berk. &M.A.Curtis) Pat. Agaricales Agaricaceae x SS Lyophyllum infumatum (Bres.) Kühner Agaricales Lyophyllaceae x EM Macrolepiota procera (Scop.) Singer Agaricales Agaricaceae x E LS Macrolepiota sp. Singer Agaricales Agaricaceae x E LS Marasimus sp1. Fr. Agaricales Marasmiaceae x E LS Marasmiellus chamaecyparidis (Hongo) Hongo Agaricales Omphalotaceae x LS Marasmius arborescens (Henn.) Beeli Agaricales Marasmiaceae x x LS Marasmius candidus Fr. Agaricales Marasmiaceae x E LS Marasmius guyanensis Mont. Agaricales Marasmiaceae x x x E LS Marasmius oreades (Bolton) Fr. Agaricales Marasmiaceae x x E LS Marasmius purpureostriatus Hongo Agaricales Marasmiaceae x x x E LS Marasmius scorodonius (Fr.) Fr. Agaricales Marasmiaceae x E LS Marasmius siccus Schwein. ex Fr. Agaricales Marasmiaceae x x E LS Marasmius sp2. Fr. Agaricales Marasmiaceae x x x E LS Marasmius sp3. Fr. Agaricales Marasmiaceae x x E LS Marasmius undatus (Berk.) Fr. Agaricales Marasmiaceae x x x E LS Micropsalliota sp. H¨ ohn. Agaricales Agaricaceae x SS Mycena griseoviridis A.H. Sm. Agaricales Mycenaceae x x LS Mycena interrupta (Berk.) Sacc. Agaricales Mycenaceae x LS Mycena rhenana Maas Geest. & Winterh. Agaricales Mycenaceae x x x LS Mycena rosea Gramberg Agaricales Mycenaceae x LS Mycena sp1. (Pers.) Roussel Agaricales Mycenaceae x LS Mycena sp2. (Pers.) Roussel Agaricales Mycenaceae x LS Mycena stipata Maas Geest. & Schw¨ obel Agaricales Mycenaceae x x x LS Mycena tenerrima (Berk.) Qu´ el. Agaricales Mycenaceae x LS Neopaxillus plumbeus Singer & Lodge. Boletales Serpulaceae x x SS Onnia tomentosa (Fr.) P.Karst. Hymenochaetales Hymenochaetaceae x x WS Panaeolina foenisecii (Pers.) Maire Agaricales Psathyrellaceae x x x SS Panaeolus fimicola (Fr.) Qu´ el. Agaricales Psathyrellaceae x DS Panaeolus papilionaceus (Bull.) Qu´ el Agaricales Psathyrellaceae x x DS Panellus mitis (Pers.) Singer Agaricales Mycenaceae x x WS Phaeolus schweinitzii (Fr.) Pat. Polyporales Fomitopsidaceae x x WS Phellinus noxius (Corner) G. Cunn. Hymenochaetales Hymenochaetaceae x x PP Phellinus populicola Niemel¨ a Hymenochaetales Hymenochaetaceae x PP Pholiota aurivella (Batsch) P. Kumm. Agaricales Strophariaceae x x E WS Pleurotus luteoalbus Beeli Agaricales Pleurotaceae x x x E WS Pleurotus populinus O.Hilber &O.K.Mill. Agaricales Pleurotaceae x x E WS Pleurotus pulmonarius (Fr.) Qu´ el. Agaricales Pleurotaceae x x E WS Pluteus longistriatus (Peck) Peck Agaricales Pluteaceae x LS Pluteus mammillatus (Longyear) Minnis, Sundb. & Methven. Agaricales Pluteaceae x LS Pluteus umbrosus (Pers.) P. Kumm. Agaricales Pluteaceae x x x LS Polyporus brumalis (Pers) Fr. Polyporales Polyporaceae x x x WS Polyporus tenuiculus (P. Beauv.) Fr. Polyporales Polyporaceae x x WS Polyporus varius (Pers.) Fr. Polyporales Polyporaceae x x x WS Psathyrella candolleana (Fr.) Maire Agaricales Psathyrellaceae x x x WS Psathyrella corrugis (Pers.) Konrad & Maubl. Agaricales Psathyrellaceae x x WS Psathyrella multipedata (Peck) A.H. Sm. Agaricales Psathyrellaceae x x x WS Psathyrella gracilis (Fr.) Qu´ el. Agaricales Psathyrellaceae x x x WS Psathyrella ammophila (Durieu &L´ ev.) P.D. Orton Agaricales Psathyrellaceae x x x WS Psathyrella piluliformis (Bull.) P.D.Orton Agaricales Psathyrellaceae x x x WS Psathyrella sp1. Fr. ex Qu´ el. Agaricales Psathyrellaceae x WS Psathyrella sp2. Fr. ex Qu´ el. Agaricales Psathyrellaceae x WS Psathyrella sp3. Fr. ex Qu´ el. Agaricales Psathyrellaceae x x WS (continued on next page) D. Alem et al. Forest Ecology and Management 496 (2021) 119391 7 forests and classified into 258 fungal taxa (Table 2). Although identification of sporocarps down to species level was not possible, out of the total taxa collected, 155 (60%) were identified to species level, 33 (13%) to genus level and further 69 (27%) were completely unidentified. The unidentified sporocarps were excluded from further analysis. The Basidiomycota was the dominant phylum and was represented by 10 orders, 62 families, 90 genera, and 180 species. Ascomycota was represented by three orders, seven families, seven genera, and eight species (Table 2). Among the taxa identified, the Agaricaceae was the most diverse family with 58 different taxa, followed by Psathyrellaceae (26), Tricholomataceae (22), and Mycenaceae (20), which together accounted for 33.6% of the total collected taxa (Table 2). The most abundant genera were Termitomyces, Psathyrella, Leucoagaricus, Marasmius, and Mycena. The proportions of macrofungal taxa at the genus level are provided (Fig. 2A). The Agaricales was the most prevalent order in the three forests (77.66%). Since many Agaricales are conspicuous macrofungi, it is not surprising to find a higher abundance during sampling. The family to genus and genus to species ratios were 0.70 and 0.50, respectively. Total numbers of fungal taxa per family encountered in the three studied forests are provided (Fig. 2B). In terms of the trophic groups, the majority of species were saprophytic (81%) followed by ectomycorrhizal (14%) and parasitic taxa (4%). The accumulation curves (Fig. 3A) generated for the taxa identified in the three forests show that the saturation of macrofungal richness was not reached during the survey given that the curves showed a steady increase with additional samplings. Although there was no significant difference in species richness between the three forests (p >0.05), the taxa accumulation curve for Banja forest showed a relatively steeper rising slope and yielded higher macrofungal richness values than the other forests. The highest macrofungal diversity values were obtained for Taragedam forest; however, diversity was not significantly different to that of the other two forests (Fig. 3B). The occurrence of macrofungi was more uneven in Banja forest than in the other forests (Table 3), with no fungal species found at all sampling events and certain macrofungal Table 2 (continued) Taxa Order Family T A B E LM Psathyrella sp4. Fr. ex Qu´ el. Agaricales Psathyrellaceae x WS Psathyrella sp5. Fr. ex Qu´ el. Agaricales Psathyrellaceae x x x WS Psathyrella sp6. Fr. ex Qu´ el. Agaricales Psathyrellaceae x WS Pseudoclitocybe cyathiformis (Bull.) Singer Agaricales Tricholomataceae x LS Pseudohydnum gelatinosum (Scop.) P.Karst. Auriculariales Exidiaceae x x x WS Pseudoomphalina pachyphylla (Fr.) Knudsen. Agaricales Tricholomataceae x LS Psilocybe ovoideocystidiata Guzm´ an & Gaines Agaricales Strophariaceae x x LS Psilocybe samuiensis Guzm´ an, Bandala & J.W.Allen Agaricales Strophariaceae x LS Ramaria stricta (Pers.) Qu´ el. Gomphales Gomphaceae x x x E EM Rhizopogon luteolus Krombh. Boletales Rhizopogonaceae x x x E EM Rhizopogon pseudoroseolus A.H. Sm. Boletales Rhizopogonaceae x E EM Russula gracillima Jul. Sch¨ aff. Russulales Russulaceae x EM Russula ochroleuca Pers. Russulales Russulaceae x x x EM Sarcoscypha occidentalis (Schwein.) Sacc. Pezizales Sarcoscyphaceae x x x WS Scleroderma areolatum Ehrenb. Boletales Sclerodermataceae x EM Scleroderma aurantium (L.) Pers. Boletales Sclerodermataceae x EM Sebacina concrescens (Schwein.) P. Roberts Auriculariales Exidiaceae x EM Skeletocutis carneogrisea A.David Polyporales Polyporaceae x x WS Suillus luteus (L.) Roussel Boletales Suillaceae x E EM Suillus sp. Gray Boletales Suillaceae x EM Terfezia leonis (Tul. & C.Tul.) Tul. Pezizales Terfeziaceae x x E EM Termitomyces clypeatus R.Heim Agaricales Lyophyllaceae x x x E LS Termitomyces microcarpus (Berk. & Broome) R. Heim Agaricales Lyophyllaceae x x E LS Termitomyces robustus (Beeli) R. Heim Agaricales Lyophyllaceae x x E LS Termitomyces sp. R. Heim Agaricales Lyophyllaceae x x x E LS Termitomyces schimperi (Pat.) R.Heim Agaricales Lyophyllaceae x x x E LS Trichaptum biforme (Fr.) Ryvarden Polyporales Polyporaceae x WS Tricholoma portentosum (Fr.) Qu´ el. Agaricales Tricholomataceae x E EM Tricholoma saponaceum (Fr.) P.Kumm. Agaricales Tricholomataceae x E EM Tricholoma sp. (Fr.) Staude Agaricales Tricholomataceae x E EM Tricholomopsis rutilans (Schaeff.: Fr.) Sing. Agaricales Tricholomataceae x x x WS Volvariella speciosa (Fr.) P.Kumm. Agaricales Pluteaceae x x LS Wilcoxina mikolae (Chin S. Yang & H.E. Wilcox) Chin S. Yang & Korf Pezizales Pyronemataceae x x E EM Xeromphalina caulicinalis (Bull.) Kühner & Maire Agaricales Mycenaceae x x x WS Xeromphalina tenuipes (Schwein.) A.H.Sm. Agaricales Mycenaceae x x x WS Xerula radicata (Relhan) D¨ orfelt Agaricales Physalacriaceae x x PP Xylaria hypoxylon (L.) Grev. Xylariales Xylariaceae x WS Xylaria scruposa (Fr.) Fr. Xylariales Xylariaceae x x WS Note: Abbreviations: T =the Taragedam forest group; A =the Alemsaga forest group; B =the Banja forest group; x =sporocarp production; E =edible; LM =mode of life; PP =Plant pathogen; EM =ectomycorrhizal, SS =Soil saprotroph, WS =Wood saprotroph, LS =Litter saprotroph, DS =Dung saprotroph. Table 3 Macrofungal and vascular plant richness and diversity indices in three church forests in Northern Ethiopia. Forest status Banja forest Taragedam forest Alemsaga forest All macrofungi Richness 22.56 ±3.02a 18.44 ±2.34a 22.67 ±1.84a Shannon 2.03 ±0.23a 2.57 ±0.13a 2.06 ±0.20a Simpson 0.73 ±0.05b 0.88 ±0.02a 0.77 ±0.05ab Evenness 0.38 ±0.03c 0.60 ±0.03a 0.47 ±0.02b Vascular plants Richness 5.78 ±0.55c 16.89 ±1.25a 12.67 ±1.04b Shannon 1.38 ±0.12b 2.18 ±0.08a 2.04 ±0.07a Simpson 0.67 ±0.05b 0.83 ±0.02a 0.82 ±0.02a Evenness 0.73 ±0.04a 0.55 ±0.03b 0.63 ±0.03ab Ectomycorrhizal fungi Richness 3.88 ±0.64a 2.57 ±0.3a 2.67 ±0.21a Shannon 1.09 ±0.1a 0.80 ±0.06a 0.90 ±0.07a Simpson 0.61 ±0.03a 0.53 ±0.02a 0.57 ±0.02a Evenness 0.85 ±0.05a 0.91 ±0.05a 0.94 ±0.03a Note: Values shown are means ±the SE of the mean. Different lowercase letters indicate a significant difference (p <0.05) in richness or diversity between forests. D. Alem et al. Forest Ecology and Management 496 (2021) 119391 8 species were more dominant in Banja forest than in the other two forests. The Shannon index and richness for vascular plants were significantly correlated with Shannon and Simpson diversity indices for the fungal communities (Fig. 4). Interestingly, for all these variables, the highest values were found in Taragedam forests and the lowest values were observed in Banja forests (Table 3). Although the three forests were not significantly different (p >0.05; Table 3) in terms of measure of diversity of their ectomycorrhizal fungal species and richness, more ectomycorrhizal species were collected from Banja forest (20) than from Taragedam (15) or Alemsaga (7) forests (Table 2). 3.2. Sporocarp production Taragedam forest produced the greatest quantity of sporocarps (25.4 kg ha −1 ), although production levels were not significantly different (p =0.63) to those of Alemsaga forest (21.6 kg ha −1 ; Fig. 5). However, both of these forests produced significantly greater quantities of sporocarps than Banja forest (p <0.05). Sixty eight (36%) of the total macrofungi collected were deemed to be edible (Table 2). Banja forest produced the greatest quantity of edible fungi (mean fresh weight, 1.8 kg ha −1 ) and Alemsaga forest produced the least (0.4 kg ha −1 ); however, the production of edible species did not differ significantly among the three forests (Fig. 5; p =0.01). Fig. 2. (A) The proportions of macrofungal taxa at the genus level (name of genus; the number of species; percentage); and (B) total numbers of fungal taxa per family encountered in the three studied forests. Fig. 3. Taxa accumulation curves generated for the fungal community found in the three studied forests using a rarefaction sample-based estimator (A) and R´ enyi diversity profiles (B). D. Alem et al. Forest Ecology and Management 496 (2021) 119391 9 3.3. Macrofungal communities and edaphic variables The perMANOVA analyses indicated the three church forests differed significantly in their macrofungal composition (F =2.05, R 2 =0.14, p = 0.001; Fig. 6). With respect to the explanatory variables, categorized edaphic, climate and location parameters were correlated to the macrofungal community composition (p <0.05; Table 4). Of these, Mantel test confirmed that location variables aggregately had a strongly significant effect on macrofungal community structure (p =0.000) than that of the climate (p =0.009) and the edaphic variables (p =0.112). The significance of each explanatory variable and their aggregated contribution to the difference of the macrofungal community compositions is provided (Table 4). The SIMPER analysis also identified macrofungal species that distinguished between the three forests (Table 5). The overall betweengroup dissimilarity (Sørensen) was 88.73% for Taragedam and Alemsaga forests, 94.44% for Alemsaga and Banja forests, and 93.76% for Taragedam and Banja forests. In this regard, the Coprinellus species are found the most important in distinguishing all forest locations along with the others (Table 5). The cumulative contribution of the most influential macrofungal species for the dissimilarity between these forests is shown in Table 5. 4. Discussion Although fragmentation poses major threats to forest ecosystems, the Dry Afromontane forests in the highland region of Ethiopia, including forest fragments owned by the church or located around church forest territories, are considered to be major reservoirs of biodiversity (Aerts et al., 2016; Aynekulu et al., 2016; Darbyshire et al., 2003; Nyssen et al., 2014). This study provides a comprehensive analysis of macrofungal communities and showed the differences in fungal community compositions of the fragmented forest systems in Northern Ethiopia. The difference in macrofungal species among the three forests might be due to the difference in vegetation composition or the variation in ecological factors such as soils, which are among the most important factors that could affect macrofungal species (Oria-de-Rueda et al., 2010). The availability of suitable substrates due to the difference in plant inputs on Fig. 4. Scatter plot matrices showing correlation coefficients between the entire tree and macrofungal variables and their significance levels. Abbreviations: T = tree, F =fungi. On the bottom of the diagonal, bi-variate scatter plots with a fitted line are displayed. 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