A contribution to the knowledge of Cunninghamella in Brazil: a new species isolated from soil with an updated key to the genusAbstract
A new species of Cunninghamella was isolated during an expedition searching for mucoralean fungi from soil in an upland tropical forest area in the state of Pernambuco, Brazil. The new species was described based on morphophysiological and phylogenetic data (ITS and LSU rDNA sequences). It typically forms unbranched sporophores, though there is some monopodial and sympodial branching. Verticillate branches arising in whorls of up to three from the main sporophore are rarely observed. Vesicles varied-shaped with one to several broken pedicels or warts on their surface and commonly with immature pedicellate sporangiola are formed. Chlamydospores are globose. Better growth was observed at temperatures of 25 and 30 °C on MEA. The new species was positioned closer to a clade containing C. clavata, C. subclavata, and C. verrucosa with high support values. In this study, a new species of Cunninghamella is proposed and an updated identification key for Cunninghamella is provided.
Keywords:
ITS and LSU rDNA; Mucorales; Mucoromycota; Mucoromycetes; Taxonomy; upland tropical forest
Introduction
The genus Cunninghamella, established by Matruchot (1903), belongs to the family Cunninghamellaceae, order Mucorales and phylum Mucoromycota (Wijayawardene et al., 2022). Species of this genus are morphologically characterized by the formation of sporophores of different sizes and branching patterns, bearing pedicellate and unispored sporangiola on the surface of a vesicle (Zheng & Chen, 2001).
For many years, the taxa of this genus were delimited according to the maximum growth temperature, color and texture of the colonies, branching pattern of the sporophores, shape and size of the vesicles and sporangiola and the presence or absence of spines on the sporangiola. Zycha (1935), Alcorn & Yeager (1938), Naumov (1939), Cutter (1946), Milko & Beljakova (1967) and Samson (1969) have reported 43 species names for Cunninghamella, however, only three species were reported in common in these works. Most of the cited taxa were species taxonomically similar to others already published, which were later synonymized or invalidated (Zheng & Chen, 2001).
After a taxonomic study using almost 200 specimens of Cunninghamella, Zheng & Chen (2001) delimited 12 species for the genus, which were later confirmed based on phylogenetic analysis of the ITS rDNA region (Liu et al., 2001). Currently, 26 species of Cunninghamella are accepted (Wijayawardene et al., 2022; Zhao et al., 2023) from which only seven species and three varieties have been reported for Brazil, namely C. bertholletiae Stadel, C. blakesleeana Lendn., C. elegans Lendn., C. clavata R.Y. Zheng & G.Q. Chen, C. echinulata var. echinulata (Thaxt.) Thaxt. ex Blakeslee, C. echinulata var. antarctica (Caretta & Piont.) R.Y. Zheng & G.Q. Chen, C. echinulata var. verticillata (F.S. Paine) R.Y. Zheng & G.Q. Chen, C. gigacellularis A.L. Santiago, C.L. Lima & C.A.F. de Souza, and C. phaeospora Boedijn (Assis et al., 2010; Hyde et al., 2016; Alves et al., 2017; Lima et al., 2018; Wang et al., 2022; Flora e Funga do Brasil, 2024; Index Fungorum, 2024).
Cunninghamella species are saprobic, generally isolated from soil, mammalian excrement, fruits and decaying organic matter (Baijal & Mehrotra, 1980; Yu et al., 2014; Nguyen et al., 2017). However, some species of the genus have been reported to cause severe infections in humans (e.g. C. bertholletiae, C. echinulata, C. elegans and C. blakesleeana). Reports are usually in immunosuppressed individuals, such as patients with uncontrolled diabetes or with hematological deficiency (Yu et al., 2014). However, C. arunalokei V. Hallur, S. Rudramurthy & H. Prakash has been described in India as causing disease in an immunocompetent individual (Hallur et al., 2021). Cunninghamella bertholletiae has also been isolated after causing fatal pneumonia in a captive killer whale (Orcinus orca Linnaeus) and causing meningoencephalomyelitis in a free-ranging bottlenose dolphin (Tursiops truncatus Montagu) (Abdo et al., 2012; Isidoro-Ayza et al., 2014; Bragulat et al., 2017).
Species of Cunninghamella are used as reliable models for drug metabolism studies (Asha & Vidyavathi, 2009). Cunninghamella elegans is capable of neutralizing some pollutants and polycyclic aromatic hydrocarbons (PAH) (Lisowska & Dlugonski, 2003), in addition to industrial dyes (Cha et al., 2001) and degrades the polycyclic aromatic nitrated hydrocarbons considered mutagenic agents and carcinogens (Pothuluri, 1996). Cunninghamella blakesleeana is also able to produce polyunsaturated fatty acids such as omega-6 (Sukrutha et al., 2014). A strain of C. elegans (UCP 0542) has been reported to be a good biosurfactant producer using food industry waste, highlighting its biotechnological potential to reduce bioprocessing costs (Medeiros et al., 2022).
During a study on the diversity of fungi in an upland forest area in Pernambuco, northeastern Brazil, specimens of Cunninghamella were isolated from soil samples. Their identity was confirmed by morphological and molecular (ITS and LSU regions of rDNA) data. In this work, we describe and illustrate a new species, and provide an updated identification key for the genus Cunninghamella.
Materials and methods
Sampling site and soil collection
Soil samples were collected in September and October 2021 in the district of Jenipapo, municipality of Sanharó (8°17'08.6"S 36°30'53.9"W and 8°17'13.6"S 36°30'52.7"W), located in the state of Pernambuco, Brazil. The local vegetation comprises subdeciduous and deciduous forests. The climate is tropical rainy, with dry summers. The rainy season starts in January/February, ends in September, and can extend until October. The average annual temperature is 31 °C, with an average annual rainfall of 496 mm (Beltrão et al., 2005). Using sterilized spatulas, soil samples were collected at a depth of 5 cm, packed in sterile plastic bags and stored in styrofoam boxes with ice during transport to the Laboratory of Zygosporic Fungi at the Universidade Federal de Pernambuco (UFPE).
Isolation and purification of the strains
Five milligrams of soil were added to wheat germ agar culture medium (Benny, 2008), with the addition of chloramphenicol (80 mg L-1), contained in Petri dishes. Colony growth was monitored for 72 hours at room temperature (26 ± 2 °C). Fragments of mycelium were removed directly from the Petri dishes under a Leica EZ4 stereomicroscope and transferred to malt extract agar (MEA) plates (Benny, 2008). Mycelium fragments from the specimens were transferred to slides with 2 % KOH or lactophenol blue and observed under light microscopy (Leica DM500).
Experiments
Pure/axenic cultures from specimens were cultured in triplicate, in MEA and potato dextrose agar (PDA), and incubated at 10, 15, 20, 25, 30, 35, and 40 °C, for 7 days in the dark. Fragments of mycelia were removed from cultures, placed on microscope slides with KOH (3 %) and observed under a light microscope (Leica DM500). For species description forty measurements were made for each fungal structure from Petri dishes incubated at 25 °C for 7 days on MEA in the dark. The color designation of colonies was established according to Kornerup & Wanscher (1978). Mating experiments were carried out on three MEA and PDA plates at 25, 30 and 35 °C, on which one 5 mm colony disk of each isolate was placed on opposite sides of the plates. The holotype was deposited in a metabolically inactive state by lyophilization in the URM Culture Collection of the Federal University of Pernambuco. Living cultures were deposited in test tubes containing slanting PDA in the same Culture Collection.
DNA extraction, amplification, purification, and sequencing
Fungal biomass was obtained from MEA slant cultures incubated at 28 °C for up to five days and was transferred to 2 mL microtubes with screw caps. To each tube, 0.5 g of acid-washed glass beads of two different diameters (150-212 μm and 425-600 μm, 1:1) (Glass beads, Sigma-Aldrich, Darmstadt, Germany) were added and the fungal biomass was crushed by stirring at high speed in a FastPrep homogenizer (FastPrep-24, MP Biomedicals, California, USA). The genomic DNA extraction procedure was conducted as described by Oliveira et al. (2016), where the mycelium was homogenized in CTAB lysis buffer [2 % cetyltrimethylammonium bromide, 20 mM EDTA, 0.1 M Tris-HCl (pH 8.0), 1.4 M NaCl; Doyle & Doyle, 1987, 1990], and washed with chloroform: isoamyl alcohol (24:1). The DNA-containing supernatant was then separated from the hyphal residues. The supernatant was mixed with an equal volume of isopropanol followed by DNA precipitation after incubation at -20 °C for 30 min. After centrifugation at 13.000 rpm for 15 min the resulting DNA pellet was washed with 70 % ethanol and resuspended in 50 μL ultrapure water. For the amplification of the ITS and the LSU regions of rDNA, the primer pairs ITS1/ITS4 and LR1/LSU2 (White et al., 1990; Van Tuinen et al., 1998; Santiago et al., 2014) were used, respectively. The final amplicons were purified with an enzymatic mix (NucleoSAP, Molecular Biotecnologia, Belo Horizonte, Brazil) and used for sequencing at Plataforma Multiusuários de Sequenciamento, Centro of Biociências, Universidade Federal de Pernambuco - UFPE (Pernambuco, Brazil). Direct sequencing of the ITS region from PCR products of strains URM 8557 and URM 8842 failed. PCR products were cloned using the pGEM-T Easy Vector (Promega, Madison, WI, USA), following the manufacturer’s instructions. These clones were sequenced using the primers M13F forward (5’-GTAAAACGACGGCCAGT-3’) and M13R reverse (5’-GCGGATAACAATTTCACACAGG-3’).
Sequence alignment and phylogenetic analysis
The sequences of the strains URM 8557 and URM 8842 were compared with those available in the National Center for Biotechnology Information GenBank database using the tool BLASTn to find the genetically closest sequences (Table 1). Sequences were aligned using MAFFT v.7 (https://mafft.cbrc.jp/alignment/server) (Katoh & Standley, 2013) for each of the molecular markers. Sequences were edited and alignments of the ITS and LSU (rDNA) regions were manually concatenated in MEGA v.7 prior to phylogenetic analyzes (Kumar et al. 2016). Bayesian inference (BI) and maximum likelihood (ML) analyses were performed with MrBayes v.3.2.2 (Ronquist et al., 2012) on XSEDE and RAxML-HPC BlackBox v.8.2.8 (Stamatakis et al., 2008; Stamatakis, 2014), respectively, using the CIPRES Science Gateway (http://www.phylo.org/) (Miller et al., 2010). The ML analysis was performed using the GTR+I+G standard nucleotide substitution model, and the BI analysis was performed using the best nucleotide model selected by AIC in MrModeltest 2.3. (Nylander, 2004). Bayesian Inference analysis was conducted with 1 × 106 generations with a burn-in value of 25 %. Phylogenetic trees were viewed and arranged using Interactive Tree of Life (iTOL) v4 (https://itol.embl.de/) (Letunic & Bork, 2019). Values less than 0.90 BI posterior probability and 70 % ML bootstrap were not depicted on the tree. The newly obtained sequences were deposited in the GenBank database. GenBank accession numbers are listed in Table 1.
Results
The alignment of ITS and LSU consisted of 55 sequences and 2102 characters with 1391 and 711 characters used in the ITS and LSU analyses, respectively. The phylogenetic analysis characterized the isolated sequences within Cunninghamella with high support values. The topology of the tree, as well as the ML bootstrap values and Bayesian inference posterior probabilities (>70 % and >0.95, respectively), are shown in Figure 1. URM 8557 and URM 8842 were placed closer to a clade containing C. clavata, C. subclavata H. Zhao, Y.C. Dai, Yuan Yuan & X.Y. Liu, and C. verrucosa H. Zhao & X.Y. Liu with high support values.
Phylogenetic tree of Cunninghamella neoverrucosa (URM 8557 and URM 8842) and related species based on the combined ITS and LSU rDNA sequences. Support values from maximum likelihood analyses and Bayesian inference. Bootstrap values lower than 70% and 0.95 or absent are marked with “-”. New taxa in blue and bold. Absidia aguabelensis URM 8213 and A. saloaensis URM 8209 were used as outgroups. “T” and “NT” represent ex-type and ex-neotype, respectively.
Taxonomy
Cunninghamella neoverrucosa F.R.S. Santos, H.B. Lee & A.L. Santiago sp. nov. - Figure 2
Mycobank: MB849789
Colony floccose, white (3-1A), reverse yellow (3-8A), with rapid growth, reaching the entire Petri dish (9 cm diam. and 1.5 cm in height) in 3 days on MEA at 25 °C. Rhizoids hyaline, long and short, root- or finger-shaped. Stolons hyaline, up to 12 μm wide. Sporophores hyaline, erect, rarely recurved, 7-480 (-600) × 2-12 μm, arising from stolons and from aerial hyphae, commonly unbranched, occasionally simply and monopodially branched, smooth-walled. Long and short branches may occasionally arise from the same sporophore, and lateral branches, some re-branching up to 4 times, may arise next to the apical vesicle. Verticillate branches are rare with up to 3 branches in a whorl. The main sporophore may be forked. Sometimes two branches arise from the same point on the sporophores. Some sporophores may arise directly from the vesicles. Axial vesicles, hyaline, mainly globose 15-35 (-40) μm diam., some subglobose 25-30 (-35) × 17-30 μm; lateral vesicles mostly clavate slightly angulated on the surface, some subglobose, 12-35 (-50) × 9.5-25 (-30) μm. Pedicels 2.5-5 μm long. Vesicles with one to several broken pedicels or warts on their surface and commonly with immature pedicellate sporangiola. Sporangiola hyaline, with greenish internal content, mostly globose, (5-) 7-21.5 (-25) μm diam., some subglobose 12-14.5 × 14.5-20 μm, unispored, echinulate. Chlamydospores globose 12-14.5 (-20) μm diam. Zygospores not formed.
Cunninghamella neoverrucosa (Holotype URM 8557h). A. Colony surface (left) and reverse (right) on malt extract agar (MEA) at 25 °C. B. Unbranched sporophore with sporangiola. C. Branched sporophore with vesicles and warts (arrows) on their surface. D. Repeatedly branched sporophore with vesicles. E. Branched sporophore with vesicles and immature pedicellate sporangiola. F. Forked sporophore with vesicles and sporangiola. G. Branched sporophore with two sporophores arising directly from the vesicle (arrow). H. Rhizoids. I. Sporangiola.
Etymology: referring to the formation of sporophores with vesicles with one to several broken pedicels or warts on its surface as observed in Cunninghamella verrucosa.
Material examined: Brazil, Pernambuco, the district of Jenipapo, municipality of Sanharó (8°17'08.6"S 36°30'53.9"W), in soil, 14 September 2021, collected by F.R.S. Santos (Holotype URM 8557h; ex-type URM 8557). GenBank accession numbers: OR502876 (ITS) and OR502877 (LSU).
Other material examined: Brazil, Pernambuco, the district of Jenipapo, municipality of Sanharó (8°17'13.6"S 36°30'52.7"W), in soil, 16 October 2021, collected by F.R.S. Santos (URM 8842). GenBank accession numbers: OR502875 (ITS) and OR885680 (LSU).
Habitat: Soil.
Distribution: Pernambuco state (Brazil).
Media and temperature test. - On MEA, at 10 °C - no growth, at 15 °C - slow growth (7.5 cm diam. after 168 h); at 20 °C - good growth (9 cm diam. after 120 h), at 25 and 30 °C - excellent growth (9 cm diam. after 72 h); at 35 °C - slow growth (7 cm diam. after 192 h), at 40 °C - no growth. Cunninghamella neoverrucosa exhibited similar growth and development of reproductive structures in MEA and PDA culture media at all temperatures tested.
Identification key to species of the genus Cunninghamella [adapted from Zheng & Chen (2001 )].
1. Homothallic, sporophores unbranched or simply branched, zygospores tuberculate..................................................................................................C. homothallica
1. Heterothallic, sporophores often more branched, zygospores absent...........................2
2. Pinkish-cinnamon brown colonies, sporophores branched in a zigzag pattern, with many septa...........................................................................................................C. septata
2. Colonies not pinkish-cinnamon brown, sporophores not zigzag branching, with no septa or only a few septa....................................................................................................3
3. Persistently white colonies.............................................................................................4
3. Colonies first white, then pigmented..............................................................................6
4. Vesicles typically inflated in the lower part (4/5) and narrowing in the upper part (1/5)................................................................................................................C. vesiculosa
4. Vesicles never as above.................................................................................................5
5. Giant cells not formed; vesicles commonly with one to several broken pedicels or warts on their surface……...................................................................................C. neoverrucosa
5. Giant cells formed; vesicles without broken pedicels or warts on their surface.......................................................................................................C. gigacellularis
6. Sporangiola exclusively globose...................................................................................7
6. Sporangiola not exclusively globose...........................................................................14
7. Rhizoids infrequent, root-like, when present, always branched……………...C. arrhiza
7. Rhizoids abundant, root-like or finger-like, branched or unbranched………………….8
8. Vesicles clavate and/or subclavate formed……………………………………………9
8. Vesicles clavate and/or subclavate not formed……………………………………….11
9. Pedicels never exceeding 4 μm in length………………………………... C. subclavata
9. Pedicels commonly longer ……………………….……………………….…………10
10. Sporophores branches 1-4, single, in pairs, or 3-4 arranged pseudoverticillately; vesicles without broken pedicels on their surface; sporangiola 9.5-20 (-27) μm diam..... .............................................................................................................................C. clavata
10. Sporophores usually unbranched, occasionally simply, monopodially or verticilatelly branched; vesicles commonly with one to several broken pedicels on their surface; sporangiola up to 15.5 μm diam.......................................................................C. verrucosa
11. Vesicles globose to subglobose, commonly angular.................................C. intermedia
11. Vesicles globose, subglobose and/or ovoid, never angular........................................12
12. Vesicles exclusively globose……………………………………….…...C. arunalokei
12. Vesicles globose, subglobose and/or ovoid………………………………………………13
13. Colonies brownish-grey; vesicles globose and subglobose; sporangiola 9-15 μm diam...............................................................................................................C. globospora
13. Colonies light grey, vesicles globose, ovoid; sporangiola 6.5-10 μm diam………………………………………………………………………….C. regularis
14. Colonies cream-colored to brownish-cream or yellowish-white................................15
14. Colonies not cream-colored to brownish-cream or yellowish-white..........................20
15. Giant dark sporangiola present in old cultures............................................................16
15. Giant dark sporangiola absent in old cultures.............................................................19
16. Sporophores typically nodular; sporangiola globose and broadly ovoid; dark giant sporangiola present after 16 days; vesicles somewhat angular; pedicels 4.5-7.5 (-10) μm long......................................................................................................................C. nodosa
16. Sporophores not nodular; sporangiola globose, broadly ovoid and lacrimoid; dark giant sporangiola present after 5 days; vesicles not angular; pedicels 2-4 (-7.5) μm long..................................................................................................................................17
17. Sporophores with or without a terminal vesicle on the main axis; branches typically long and short in the same sporophore and repeatedly branched; dark and giant sporangiola relatively rare to few............................................C. echinulata var. antarctica
17. Sporophores usually with a terminal vesicle on the main axis; branches mainly similar in length but sometimes rarely long and short in some sporophores and usually not repeatedly branched; dark giant sporangiola usually abundant........................................18
18. No verticillate branches found in the sporophores, primary branches of sporophores 0-4 (-10) in number, not short and compact; terminal vesicles on the main axes globose to subglobose, 13-30 (-40) μm diam......................................C. echinulata var. echinulata
18. Sporophores usually with many whorls of verticillate or pseudoverticillate branches, primary branches of sporophore (0-)4-15(-25) in number, short and compact; terminal vesicles on the main axis somewhat depressed-globose, 22-50(-70) μm diam.......................................................................................C. echinulata var. verticillata
19. Maximum growth temperature at 40 °C; chlamydospores formed........C. saisamornae
19. Maximum growth temperature at 36-38 °C; chlamydospores not formed........................................................................................................C. blakesleeana
20. Giant dark sporangiola present……………………………………………...C. varians
20. Giant dark sporangiola absent……………………………………………………....21
21. Main axes of sporophores typically with long and short branches on the same sporophore and/or repeatedly branched...........................................................................22
21. Main axes of sporophores with branches subequal in length on the same sporophore and not repeatedly branched, some branches may show secondary branches...................23
22. Sporophore branching typically binary; sterile branches sometimes present; axial vesicles present and sometimes with irregular shapes; lacrimoid sporangiola reaching 32 μm long….........................................................................................................C. binareae
22. Sporophore branching not binary; absence of sterile branches; axial vesicles present or absent, usually not irregularly shaped when present; lacrimoid sporangiola up to 16 μm long………...........................................................................................C. bertholletiae
23. Colonies with an irregular margin or reverse like a laminated cloud..........................24
23. Colonies not as above………………………….……………………………………25
24. Colonies with an irregular margin; sporophores unbranched, or simply branched, in pairs, sympodial, but never verticillate; vesicles ovoid, subglobose, globose and rarely clavate, even irregular ………………………………………………………C. irregularis
24. Colonies with reverse like a laminated cloud; sporophores unbranched or monopodially, verticillatelly, usually laterally branched; vesicles globose to subglobose……………………...………………………………………...C. guizhouensis
25. Presence of lipid droplets in sporangiola……………………………...........C. guttata
25. Without lipid droplets in sporangiola ………………………………………....….26
26. Colonies dark grey; sporophores relatively scanty as compared with other taxa, predominantly simple or monopodially branched; branching solitary or in pairs, very rarely whorled or pseudo-whorled...............................................................C. phaeospora
26. Colonies pale grey; sporophores not scanty, monopodially and commonly verticillately branched.....................................................................................................27
27. Maximum growth temperature at 42 °C; many vesicles irregular in shape and slightly angular...................................................................................................C. multiverticillata
27. Maximum growth temperature at 34-36 °C; vesicles not as above………...C. elegans
Discussion
In this study, Cunninghamella neoverrucosa is proposed as a new species based on morphological characteristics and phylogenetic analyses. Our concatenated ITS and LSU rDNA tree demonstrated that the new species is distinct from all other species, as it forms a highly supported clade sister to a clade with C. subclavata, C. verrucosa and C. clavata (Fig. 1).
The new species differs from C. subclavata by the formation of chlamydospores, and by its larger sporangiola that can reach 21.5 (-25) μm in diam., while those of C. subclavata are up to 17 μm in diam. (Zhao et al., 2023). In addition, only C. neoverrucosa forms vesicles with one to several broken pedicels or warts on their surface and commonly with immature pedicellate sporangiola. Cunninghamella neoverrucosa also differs from C. subclavata by forming sporophores that arise directly from vesicles. The color of the vesicles of C. neoverrucosa has a grayish tone, whereas the ones of C. verrucosa are hyaline or sub-hyaline.
Cunninghamella neoverrucosa seems morphologically similar to C. verrucosa, as both species form sporophores with vesicles with one to several broken pedicels or warts on their surface, which is a prominent feature of both species. However, after analyzing the two species in more detail, some differences became quite evident. Firstly, the colony of C. neoverrucosa is persistently white, differing from C. verrucosa which turns grey with age. Secondly, sporophores of the latter are unbranched or simply and monopodially branched, differing from the ones of the new species that are simply, monopodially but also sympodially (up to 4 times), branched. Thirdly, vesicles of C. neoverrucosa are globose (majority) or subglobose, whereas C. verrucosa forms vesicles which are usually clavate and oval, sometimes subglobose. Finally, C. verrucosa forms sporangiola up to 17.6 μm diam. (Wang et al., 2022), smaller than the ones of the new species.
Cunninghamella neoverrucosa differs from C. clavata by forming mostly globose axial vesicles, many of which have several broken pedicels or warts on their surface, while axial vesicles of C. clavata are typically clavate and with no broken pedicels or warts on their surface. In addition, sporophores may branch repeatedly up to seven times in C. clavata, but only up to four times in C. neoverrucosa. Sporangiola of C. clavata are consistently globose, and pedicels are up to 11 μm long, whereas sporangiola of the new species are globose and subglobose, and pedicels are shorter. In addition, chlamydospores are not formed in C. clavata (Zheng & Chen, 2001). To the best of our knowledge, until now, chlamydospores have only been reported in C. saisamornae Suwannar. & Wongputtisin (Suwannarach et al., 2021).
In conclusion, our data demonstrate that C. neoverrucosa is morphologically and genetically different from other Cunninghamella species described so far. Therefore, it is described as new. This study contributes to the knowledge of the taxonomy and diversity of Cunninghamella, a genus of great importance for medicine and biotechnology.
Acknowledgments
The authors express their gratitude to the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES) for the scholarship awarded to Francisca Robervânia Soares dos Santos and the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) for the research grant to André Luiz Santiago. This research was also financed by the project ‘Morphological and molecular approaches to the knowledge of zygosporic fungal communities in the Atlantic Forest of Pernambuco and Paraíba, Brazil’ (FACEPE APQ-1346-2.12/22). This work was also supported by National Research Foundation of Korea (NRF) funded by the Ministry of Science and Information and Communications Technology (2022M3H9A1082984), and the Basic Science Research Program funded by the Ministry of Education (2022R1I1A3068645).
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