Phylogeny of Gelatinodiscaceae (Leotiomycetes) with emphasis on the taxonomy of Ascocoryne in North America

JK Stallman; L Quijada; MC Aime; CA Quandt; DB Raudabaugh; T Melie; I Rimer; S McAdam; D Haelewaters

doi:10.3114/persoonia.2026.56.07

. 2026 May 29;56:420–456. doi: 10.3114/persoonia.2026.56.07

Phylogeny of Gelatinodiscaceae (Leotiomycetes) with emphasis on the taxonomy of Ascocoryne in North America

JK Stallman ^1,^*, L Quijada ^2,³, MC Aime ¹, CA Quandt ⁴, DB Raudabaugh ¹, T Melie ⁵, I Rimer ¹, S McAdam ¹, D Haelewaters ^6,⁷

PMCID: PMC13409176 PMID: 42524026

Abstract

The family Gelatinodiscaceae and genus Ascocoryne represent diverse lineages in Helotiales (Ascomycota, Leotiomycetes) that lack well-sampled phylogenies and taxonomic study. Our research objectives were to 1) determine whether certain genera that lacked DNA sequence data or had uncertain phylogenetic placement — Clathrosporium, Phaeangellina, Skyathea, and Xerombrophila — belonged to Gelatinodiscaceae, 2) estimate species richness and describe new species in Ascocoryne, 3) determine if morphological characteristics delineate intrageneric clades in Ascocoryne, and 4) test the mitochondrial tricarboxylate transport protein (CTP) as an alternative DNA barcode in Ascocoryne. We made 83 Gelatinodiscaceae collections primarily from the midwestern United States, examined 61 collections previously deposited in herbaria, and generated DNA sequence data for seven loci (18S, ITS, 28S, EF1, RPB1, RPB2, CTP) from these collections. We found that Skyathea, Xerombrophila, and up to four other unnamed, putative genera belong to Gelatinodiscaceae, while Clathrosporium and Phaeangellina are excluded. Up to 37 putative undescribed species-level clades or single isolates are present in Ascocoryne, in addition to the nine previously described species and four newly described ones here: Ascocoryne diminutiva, A. fibrosa, A. neoinflata, and A. wilsoniae. Additionally, descriptions for two unnamed species similar to A. cylichnium are given, a preliminary description of the sexual morph of A. laurisilvae is provided, and A. trichophora is synonymized with A. vinosa. Four main clades are found in the genus Ascocoryne. The size of lipid bodies in living ascospores, the arrangement of conidia borne on ascospores, and the presence of calcium oxalate crystals are the most important morphological characteristics for distinguishing among clades. The CTP and ITS loci successfully identified six of six Ascocoryne species tested. This study provides the best-sampled phylogeny of Gelatinodiscaceae and Ascocoryne and highlights unnamed genus- and species-level diversity in this group.

Citation: Stallman JK, Quijada L, Aime MC, Quandt CA, Raudabaugh DB, Melie T, Rimer I, McAdam S, Haelewaters D (2026). Phylogeny of Gelatinodiscaceae (Leotiomycetes) with emphasis on the taxonomy of Ascocoryne in North America. Persoonia 56: 420–456. doi: 10.3114/persoonia.2026.56.07

Keywords: aero-aquatic fungi, aquatic fungi, fungaria, Leotiomycetes, new taxa, new combination, vital taxonomy

INTRODUCTION

Recent reviews and phylogenetic analyses have greatly improved our understanding of Leotiomycetes, but they have also highlighted gaps in our knowledge of the systematic relationships and species richness in this group. For example, 489 Leotiomycetes genera lack multilocus sequence data, 155 of which also have no ITS sequence data available (Johnston & Park 2025). In the most speciose order, Helotiales, as many as 151 genera are considered incertae sedis (Johnston et al. 2019), and only ~5000 of the estimated 70000 species in the order have been described (Hawksworth 2001, Quandt & Haelewaters 2021). Several recent studies focused at the family rank to find stable taxonomic positions for genera and describe new species. For example, the families Arachnopezizaceae, Hyaloscyphaceae, Hyphodiscaceae, and Mollisiaceae have recently been studied (Tanney & Seifert 2020, Kosonen et al. 2021, Quijada et al. 2022), and the family Tricladiaceae was created to separate a group of polyphyletic genera in Solenopeziaceae (Johnston & Baschien 2020). In contrast, little molecular data are available for Gelatinodiscaceae, and therefore the family has received limited modern systematic study.

Gelatinodiscaceae

Gelatinodiscaceae was established for Gelatinodiscus flavidus (Carpenter 1976) and remained monotypic for over 40 years. More recently, Gelatinodiscus was synonymized with Chloroscypha (Baral et al. 2013a) and a broader Gelatinodiscaceae including nine genera was proposed based on phylogenetic and morphological evidence (Jaklitsch et al. 2016). Now the family contains at least ten genera: Ascocoryne, Ascotremella, Chloroscypha (= Gelatinodiscus), Dimorphospora, Helicodendron, Neobulgaria, Ombrophila, Phaeangellina, Skyathea, and Xerombrophila (Baral et al. 2013a, b, Sri-indrasutdhi et al. 2015, Jaklitsch et al. 2016, Ekanayaka et al. 2019, Johnston et al. 2019). Didymocoryne has been considered a distinct genus in the family (Wijayawardene et al. 2018, Ekanayaka et al. 2019) or a synonym of Ascocoryne (Jaklitsch et al. 2016, Hyde et al. 2024). Clathrosporium grouped with the Gelatinodiscaceae p.p. in ITS phylogenies and was placed either in the family (Johnston et al. 2019) or in Helotiales incertae sedis (Johnston & Park 2025), whereas Ekanayaka et al. (2019) placed it in the newly established Hydrocinaceae based on a five-locus phylogeny. Similarly, Xerombrophila was placed in Gelatinodiscaceae based on a 15-locus phylogenetic hypothesis by Johnston et al. (2019), while Ekanayaka et al. (2019) placed it in Hydrocinaceae. Finally, some older lists have included Neocudoniella in Gelatinodiscaceae (e.g., Jaklitsch et al. 2016, Wijayawardene et al. 2018), but in more recent phylogenetic reconstructions it is not confamilial (e.g., Ekanayaka et al. 2019, Johnston & Park 2025).

Species in Gelatinodiscaceae have diverse life histories as endophytes and both terrestrial and aquatic saprotrophs. Many genera and species form apothecia on wood (e.g., Chloroscypha flavida), while others also produce macroscopic asexual structures (e.g., Ascocoryne sarcoides), or are aquatic (e.g., Dimorphospora) or aero-aquatic (e.g., Helicodendron) fungi. Sexual morphs of Gelatinodiscaceae taxa can be differentiated from Helotiaceae taxa (where many current Gelatinodiscaceae genera used to be placed) by the combination of often containing gel and calcium oxalate crystals in excipular tissues with having a different amyloid ascus ring (Jaklitsch et al. 2016), but determining shared morphological characteristics at the family rank remains a challenge in Helotiales (Baral et al. 2015, Quandt & Haelewaters 2021).

Despite recent large-scale phylogenies of Leotiomycetes taxa (Haelewaters et al. 2021, Johnston & Park 2025), sampling in Gelatinodiscaceae has been low, not inclusive of all putative genera, and studies including multilocus sequence data or protein-coding genes are lacking. For example, there are no DNA sequence data available for the genera Phaeangellina and Skyathea, and only two published phylogenies include more than 15 Gelatinodiscaceae isolates: an ITS phylogeny of 27 Ascocoryne isolates (Gao et al. 2017) and an ITS+28S phylogeny of 27 Gelatinodiscaceae isolates from five genera (Ekanayaka et al. 2019). Both phylogenies showed Ascocoryne isolates that were identified as a single species were either not monophyletic or did not have high bootstrap support. An earlier phylogenetic analysis of Ascocoryne showed limited variability in the ITS region in A. sarcoides isolates but found that the mitochondrial tricarboxylate transport protein locus (CTP) successfully resolved species and subspecies relationships in Ascocoryne (Griffin et al. 2010).

Ascocoryne

Species in the genus Ascocoryne make gasoline-like, volatile organic compounds studied as potential biofuels (Gao et al. 2017). Studies focusing on this secondary metabolite production usually generate DNA sequence data but include little apothecial morphological data as these are typically studied from isolates in pure culture (Griffin et al. 2010, Gao et al. 2017). Other studies of Ascocoryne include detailed descriptions of apothecia, but do not include DNA sequence data (Quijada et al. 2017). Therefore, a link between morphology and DNA sequence data is often not available. Adding to the taxonomic challenge of this group, the genus Ascocoryne was established to accommodate members of the genus Coryne with an apothecial sexual morph (Groves & Wilson 1967) and Index Fungorum (2025) lists 86 taxa in the genus Coryne, many of which have not been reevaluated since their original description (Johnston et al. 2014). Some of these taxa may be species of Ascocoryne that should be investigated before describing new species in the genus. Therefore, incorporating both morphology and DNA sequence data while considering older Coryne names is needed when describing new species of Ascocoryne.

Ascocoryne includes nine species of endophytic–saprotrophic taxa found worldwide (Index Fungorum 2025). Most species in the genus have purplish, gelatinous apothecia that grow in association with dead wood, and some also have macroscopic asexual morphs that may grow independently or co-occur with the sexual morph. Many Ascocoryne are xylophiles, living in the bark or wounds of trees (Arhipova et al. 2015, Meyn et al. 2019, Molina et al. 2020). These endophytes may have antagonistic effects on other wood-decaying fungi and are not reported to cause substantial decay to the wood itself (Roll-Hansen & Roll-Hansen 1979a, Vasiliauskas & Stenlid 1998). One species, A. turficola, usually grows from moss in peat bogs instead of wood and is considered rare and of conservation concern throughout much of its range in the Northern Hemisphere (Bunyard et al. 2008, Filippova et al. 2013). A key to Ascocoryne species in Europe suggests the genus can be split into two groups: 1) the A. sarcoides group of which members lack calcium oxalate crystals, have one to a few large lipid bodies (LBs) with smaller and medium LBs in living ascospores, and produce conidia in chains from mature ascospores; and 2) the A. cylichnium group with members that contain calcium oxalate crystals, have living ascospores without large LBs (with only small and medium-sized LBs), and have conidia on mature ascospores that are not produced in chains (Baral 2000).

The most comprehensive taxonomic treatment of Ascocoryne in the Americas (inclusive of North, South, and Central America, including the Caribbean) was a PhD dissertation by Wilson (1962) focusing on species (at the time, in the genus Coryne) in the northeastern USA and southeastern Canada. Wilson listed four known species — C. cylichnium, C. sarcoides, C. striata, and C. urnalis (now considered a synonym of A. cylichnium) — and proposed four new species — C. angustata, C. constrictospora, C. inflata, and C. minima. Descriptions were based on features of ascomata, conidiomata, and cultures, but the descriptions contained no Latin diagnoses and were never validly published. Despite lack of valid publication, Wilson’s Coryne ‘inflata’ (nom. inval., Art. 36) has been used informally and in scientific literature (now as ‘Ascocoryne inflata’) to describe species with capitate paraphyses that occur in several regions, such as Europe (Moyne 2012) and North America (Valenzuela et al. 2021). Species lists of Ascocoryne in North America include A. sarcoides, A. cylichnium, and A. turficola (e.g., Seaver 1951, Roll-Hansen & Roll-Hansen 1979b, Bunyard et al. 2008, Beug et al. 2014). These three species, along with A. albida, A. lilacina, and A. laurisilvae, were originally described from Europe. The only species described from outside America or Europe is A. javanica from Indonesia (Penzig & Saccardo 1902).

Although studied in ecology, forestry, conservation, and biotechnology contexts for decades, there is no well-sampled phylogeny of Ascocoryne and the broader Gelatinodiscaceae with associated morphological, geographical, and host data. Here, we incorporated new and historical collections from fungaria to ask the following questions:

Do DNA sequence data place the genera Clathrosporium, Phaeangellina, Skyathea, and Xerombrophila in Gelatinodiscaceae?
Does Ascocoryne comprise monophyletic species groups distinguishable by traits such as calcium oxalate crystals in the excipulum, LB size in living ascospores, conidia arrangement on ascospores, and an amyloid (blue) reaction of the ectal excipulum in iodine?
What is the species richness of Ascocoryne?
Is the CTP locus a useful DNA barcode in Ascocoryne, and how does it compare to the ITS?

MATERIALS AND METHODS

Collections

We made 79 new collections of Ascocoryne species and four of Neobulgaria species. Two collections were made in the Upper Potaro River Basin of the Pakaraima Mountains in Guyana in 2002 and 2016. Three collections were made in the Merendón Mountains of the Department of Cortés in Honduras in 2022. All remaining collections were made in Illinois, Indiana, Michigan, Washington, and West Virginia, USA, from 2021 to 2024. We also examined 61 other Gelatinodiscaceae collections obtained from fungaria. We generated 234 DNA sequences via Sanger sequencing, Illumina sequencing, and genome mining (Table S1).

Collections from Guyana and Honduras were dried in desiccant while specimens collected in the USA were dried at ~25 °C with a fan. Newly collected specimens were deposited at the Kriebel Herbarium (PUL) and University of Guyana (BRG). Fungarium collections and cultures were requested from the University of Arizona (ARIZ), Canada’s National Mycological Herbarium (DAOM), culture collection of the Westerdijk Fungal Biodiversity Institute (CBS), University of Copenhagen (C), Cornell University (CUP), Harvard University (FH), University of Helsinki (H), Kew Royal Botanic Garden (K), New York Botanic Garden (NY), Royal Ontario Museum Fungarium (TRTC), and Washington State University (WSP).

Morphology

Field photographs were captured using a Tough TG-6 camera (Olympus, Pennsylvania, USA) or a cell phone camera. Colours are from Kornerup & Wanscher (1978). Stereo microscope photos were captured using a SC30 camera mounted on an Olympus SZ61 stereo microscope, using Olympus cellSens v. 1.18 software to take photographs. Microscopic fungal structures were observed using an Eclipse E600 microscope (Nikon, New York, USA) and photographed using the same camera and cellSens software. Biometric features were measured in Piximètre v. 5.10 (http://ach.log.free.fr/Piximetre/). Micrographs were edited and plates were assembled using Inkscape v. 0.92 (https://inkscape.org/) or Adobe Illustrator. To identify some decorticated tree substrates, sections of wood were made with a Microm HM 430 freezing microtome (Thermo Fisher Scientific, MA, USA), stained with Toluidine Blue O, observed using an Axio Imager 2 Upright Microscope (Zeiss, Jena, Germany), and compared with the Wood Database (https://www.wood-database.com/).

Microscopic characters of specimens were studied considering the distinction between living and dead cells (vital taxonomy, Baral 1992). Apothecia were hand-sectioned and mounted in various reagents: Lugol’s solution (IKI), ~ 5 % potassium hydroxide (KOH), 1 % Congo Red (CR), and Melzer’s reagent (MLZ). All morpho-biometric characters observed after treatment with lethal reagents (e.g., KOH, MLZ) correspond to dead cell traits (indicated with the symbol †). Specimens were initially observed in tap water, allowing the observation of living-cell features (indicated with the symbol *) in fresh collections and some herbarium collections. Living features include the pattern of LBs inside ascospores, the arrangement of ascospores within asci, and the presence of vacuolar bodies (VBs) in paraphyses and excipular cells. The same specimens were then treated with lethal reagents to kill the living cells and enable comparisons with dead herbarium material. A minimum of 50 ascospores and 20 of each of the other microscopic structures (e.g., asci, paraphyses, and excipular cells) were measured to complete the descriptions. Measurements are presented as (a–)b–c(–d), where b and c represent the mean minus/plus standard deviation, and a and d indicate extreme values. For the excipulum, we noted the number of distinguishable layers, texture, gelatinization, cell morphology, and any blue reaction with and without KOH pretreatment in IKI. Ascospores were analyzed thoroughly, considering the lipid content index (LCI), which differentiates guttulate content by percentage and type of guttules (oligoguttulate vs. multiguttulate). Mature ascospores may develop septa and conidia. When present, conidial formation and morphology were studied in detail.

Mature ascomata were placed beneath cover slips to observe the ejected living ascospores. The same technique was applied using inverted Petri dishes with potato dextrose agar (PDA) or Rose Bengal (RB) to initiate cultures following the methods of Karakehian et al. (2021).

Sanger sequencing

Total DNA was extracted from desiccated apothecia using the QIAamp DNA Micro, QIAamp Mini Kit (Qiagen, Stanford, California), or Promega Wizard Genomic Purification Kit (Promega, Madison, Wisconsin) following the manufacturer’s instructions. During sample collection in Honduras and Guyana, fresh apothecia were placed in lysis buffers in the field and the remaining steps of DNA extraction were completed later (3 mo to 1.5 yr) in the laboratory per manufacturer’s instructions. The DNA from cultures < 30 d in age was amplified via colony PCR (sensu Albu et al. 2015) in which a sterile toothpick was used to gather and suspend hyphae from the colony in 50 μL sterile H₂O, microwaved (at 1100 watt power) for 2 min, then centrifuged for 2 min at 18 400 rcf. Approximately 40 μL of the supernatant was retained and 5 μL was diluted in 45 μL sterilized millipore H₂O for subsequent use in PCR.

Amplification reactions were completed in 25-μL reactions with 12.5 μL of 2× MyTaq Mix (Bioline, Swedesboro, New Jersey), 1 μL of DNA extract, 9.5 μL of H₂O, and 1 μL of each 10 μM primer for rDNA loci, or 8.5 μL of H₂O and 2 μL of each 10 μM primer for protein-coding loci. All primers and thermocycler conditions are provided in Table 1. Multiple primers are listed for the same locus because in cases where PCR was unsuccessful, we attempted an alternate primer pair or primers that targeted a shorter portion of the same region.

Table 1.

Primers and thermocycler conditions used for Illumina and Sanger sequencing. Two different cycling conditions were used for RPB1.

Locus and platform	Primer name	Cycle conditions^*				Cycle number	Final extension	Primer sequence	Reference(s)
Locus and platform	Primer name	Initial denaturaturation	Denaturaturation	Annealling	Extension	Cycle number	Final extension	Primer sequence	Reference(s)
Illumina
ITS1	TS-ITS1F	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	ACACTCTTTCCCTACACGACGCTCTTCCGATCTCTTGGTCATTTAGAGGAAGTAA	White et al. (1990) (with Illumina adapter)
ITS1	TS-ITS2R	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCTGCTGCGTTCTTCATCGATGC	White et al. (1990) (with Illumina adapter)
ITS2	TS-ITS3F	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	ACACTCTTTCCCTACACGACGCTCTTCCGATCTGCATCGATGAAGAACGCAGC	White et al. (1990) (with Illumina adapter)
ITS2	TS-ITS4R	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCTTCCTCCGCTTATTGATATGC	White et al. (1990) (with Illumina adapter)
LSU	TS-LR0R	94 °C; 5 min	94 °C; 30 sec	50 °C; 45 sec	72 °C; 1 min	35	72 °C; 7 min	ACACTCTTTCCCTACACGACGCTCTTCCGATCTACCCGCTGAACTTAAGC	https://sites.duke.edu/vilgalyslab/files/2017/08/rDNA-primers-for-fungi.pdf (with Illumina adapter)
LSU	TS-LR3	94 °C; 5 min	94 °C; 30 sec	50 °C; 45 sec	72 °C; 1 min	35	72 °C; 7 min	GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCTCCGTGTTTCAAGACGGG	Vilgalys & Hester (1990) (with Illumina adapter)
Sanger
ITS	ITS1F	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	CTTGGTCATTTAGAGGAAGTAA	Gardes & Bruns (1993)
ITS	ITS2	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	GCTGCGTTCTTCATCGATGC	White et al. (1990)
ITS	ITS3	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	GCATCGATGAAGAACGCAGC	White et al. (1990)
ITS	ITS4	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	TCCTCCGCTTATTGATATGC	White et al. (1990)
ITS	ITS4a	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	CGCCGTTACTGGGGCAATCCCTG	Larena et al. (1999)
ITS	ITS5	94 °C; 5 min	94 °C; 30 sec	48 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	GGAAGTAAAAGTCGTAACAAGG	White et al. (1990)
LSU	LR0R	94 °C; 5 min	94 °C; 30 sec	50 °C; 45 sec	72 °C; 1 min	35	72 °C; 7 min	ACCCGCTGAACTTAAGC	https://sites.duke.edu/vilgalyslab/files/2017/08/rDNA-primers-for-fungi.pdf
LSU	LR5	94 °C; 5 min	94 °C; 30 sec	50 °C; 45 sec	72 °C; 1 min	35	72 °C; 7 min	TCCTGAGGGAAACTTCG	Vilgalys & Hester (1990)
LSU	LR6	94 °C; 5 min	94 °C; 30 sec	50 °C; 45 sec	72 °C; 1 min	35	72 °C; 7 min	CGCCAGTTCTGCTTACC	Vilgalys & Hester (1990)
LSU	LR7	94 °C; 5 min	94 °C; 30 sec	50 °C; 45 sec	72 °C; 1 min	35	72 °C; 7 min	TACTACCACCAAGATCT	Vilgalys & Hester (1990)
LSU	LR3	94 °C; 5 min	94 °C; 30 sec	50 °C; 45 sec	72 °C; 1 min	35	72 °C; 7 min	CCGTGTTTCAAGACGGG	Vilgalys & Hester (1990)
LSU	LIC24R	94 °C; 5 min	94 °C; 30 sec	50 °C; 45 sec	72 °C; 1 min	35	72 °C; 7 min	GAAACCAACAGGGATTG	Miadlikowska & Lutzoni (2000)
SSU	NS1	95 °C; 5 min	95 °C; 30 sec	55 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	GTAGTCATATGCTTGTCTC	White et al. (1990)
SSU	NS2	95 °C; 5 min	95 °C; 30 sec	55 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	GGCTGCTGGCACCAGACTTGC	White et al. (1990)
SSU	NS4	95 °C; 5 min	95 °C; 30 sec	55 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	CTTCCGTCAATTCCTTTAAG	White et al. (1990)
RPB1	RPB1-af	95 °C; 5 min	Denature 1	Annealing 1	Extension 1	1) 5. 2) 30	72 °C; 10 min	GARTGYCCDGGDCAYTTYGG	Stiller & Hall (1997)
RPB1	RPB1-afasc	95 °C; 5 min	Denature 1	Annealing 1	Extension 1	1) 5. 2) 30	72 °C; 10 min	ADTGYCCYGGYCATTTYGGT	Hofstetter et al. (2007)
RPB1	RPB1-cr	95 °C; 5 min	Denature 1	Annealing 1	Extension 1	1) 5. 2) 30	72 °C; 10 min	CCNGCDATNTCRTTRTCCATRTA	Matheny et al. (2002)
RPB2	fRPB2-7cR	95 °C; 4 min	95 °C; 1 min	Annealing 2	72 °C; 1 min	44	72 °C; 10 min	CCCATRGCTTGYTTRCCCAT	Liu et al. (1999)
RPB2	fRPB2-5F	95 °C; 4 min	95 °C; 1 min	Annealing 2	72 °C; 1 min	44	72 °C; 10 min	GAYGAYMGWGATCAYTTYGG	Liu et al. (1999)
EF1	EF1-983F	94 °C; 2 min	Denature 2	Annealing 3	Extension 1	1) 9. 2) 30	72 °C; 10 min	GCY CCY GGH CAY CGT GAY TTY AT	Rehner & Buckley (2005)
EF1	EF1-1018F	94 °C; 2 min	Denature 2	Annealing 3	Extension 1	1) 9. 2) 30	72 °C; 10 min	GAYTTCATCAAGAACATGAT	Stielow et al. (2015)
EF1	EF1-1567R	94 °C; 2 min	Denature 2	Annealing 3	Extension 1	1) 9. 2) 30	72 °C; 10 min	ACH GTR CCR ATA CCA CCR ATC TT	Rehner & Buckley (2005)
EF1	EF1-1620R	94 °C; 2 min	Denature 2	Annealing 3	Extension 1	1) 9. 2) 30	72 °C; 10 min	GACGTTGAADCCRACRTTGTC	Stielow et al. (2015)
CTP	4079F	94 °C; 5 min	94 °C; 30 sec	60 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	CGCCGTTTGGAAAGGCATGG	Griffin et al. (2010)
CTP	4079R	94 °C; 5 min	94 °C; 30 sec	60 °C; 45 sec	72 °C; 45 sec	40	72 °C; 7 min	GCGACGCACACGCAAACGAATTC	Griffin et al. (2010)

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Denature 1: 1) 94 °C; 1 min. 2) 94 °C; 15 sec.

Denature 2: 1) 94 °C; 30 sec. 2) 94 °C; 30 sec.

Annealing 1: 1) 37 °C; 35 sec with .5 °C increase per cycle. 2) 45 °C; 55 sec with .5 °C increase per cycle.

Annealing 2: 50 °C; 1 min with .3 °C increase per cycle.

Annealing 3: 1) 66 °C; 30 sec with 1 °C decrease per cycle. 2) 56 °C; 30 sec.

Extension 1: 1) 72 °C; 1 min. 2) 72 °C; 1 min.

The presence of PCR product was verified using gel electrophoresis at 110 V for 30 min (1 % TBE agarose gel) and SYBR GelRed. The PCR purification and Sanger sequencing were outsourced to GENEWIZ (South Plainfield, New Jersey). Sequencing was completed with the same primers as PCR at 5 μM concentration. Sequence reads were assembled and edited in Sequencher v. 5.4.6 (Gene Codes, Ann Arbor, Michigan) or Geneious v. 9.1.8 (https://www.geneious.com). All sequences generated from this study were deposited in the National Center for Biotechnology Information (NCBI) GenBank sequence database (https://www.ncbi.nlm.nih.gov/genbank/). Accession numbers with associated metadata are provided in Table S1.

High-throughput sequencing and genome mining

Metagenomic sequencing of the ITS and/or beginning of the 28S region was completed for 37 fungarium specimens when Sanger sequencing failed, an approach that has been successful for older samples (Forin et al. 2018, Miller et al. 2022, Olds et al. 2023, Stallman et al. 2024) in which DNA fragmentation and environmental contaminations likely occurred. We followed the protocol from Miller et al. (2022), except that for some isolates, the beginning of the 28S region was also sequenced. Briefly, initial amplification reactions were completed as above and in Table 1. The PCR products were cleaned with exonuclease and shrimp alkaline phosphatase, then sample-specific Illumina indices were ligated onto PCR products in a reaction with 1 μL of PCR product, 1 uL of each forward and reverse index, 9.5 uL of H₂O, and 12.5 μL of Master Mix. Paired samples from the initial PCR product and post-ligation were run in adjacent wells on a 1 % agarose gel at 60 V for 60 min to confirm ligation. Ligated samples were again cleaned with exonuclease and shrimp alkaline phosphatase and pooled for Illumina sequencing on a MiSeq v2 platform using 2 × 250 nucleotide paired-end reads. Raw Illumina reads are deposited in NCBI’s Sequence Read Archive (SRA) database under BioProject PRJNA1018967 and additional metadata are provided in Supplementary File S1 for these isolates.

Reads were processed with DADA2 v. 1.29.0 (Callahan et al. 2016) with forward and reverse amplicons in R v. 4.1.2 (R Foundation for Statistical Computing 2023) using the methodology from Miller et al. (2022). The “filterAndTrim” function was used with the options maxN = 0, maxEE = c(2,2), truncQ = 1, and rm.phix = True, and the “learnErrors” function was used on forward and reverse reads. Forward and reverse reads were then merged with the “mergePairs” function. The “removeBimeraDenovo” function with the method set to consensus was used to remove chimeric sequences. Resulting amplicon sequence variants were compared against the UNITE general FASTA release for Fungi (Abarenkov et al. 2024) using NCBI BLASTn (Altschul et al. 1990). Amplicon sequence variants (ASV) and resulting taxonomic assignments based on the highest BLASTn match were exported to a spreadsheet. For each isolate, the ASV with the most reads that matched Gelatinodiscaceae was used to assign the correct sequence to the collection. In some cases the percentage of matching reads was low (< 1 %) or multiple Gelatinodiscaceae ASVs were present. In these cases, we used additional evidence such as morphology and checked whether reads were unique to a single isolate or present across other samples and controls to determine the correct sequence or discard it due to ambiguous results. Successfully generated sequences were deposited to NCBI (Table S1).

For two collections, CBS 275.75 (DAOM 127045) and CBS 738.84, sequences were derived in full or partly from draft genomes (C.A. Quandt & D. Haelewaters, unpubl. data) assembled using the workflow in Melie (2023). Full collection information for isolates not listed in the Taxonomy section but for which DNA sequence or other data were generated is provided in Supplementary File S2.

Construction of phylogenetic trees

We used T-BAS v. 2.4 (Carbone et al. 2019) to investigate the phylogenetic positions of Phaeangellina empetri and Skyathea hederae, which previously lacked DNA sequence data, and Clathorosporium intricatum and Xerombrophila crystallifera, which had conflicting familial placements between Ekanayaka et al. (2019), and Johnston et al. (2019) and Johnston & Park (2025). We selected “Place Unknowns” for Skyathea [ITS–28S–18S of K (M)1435354], Phaeangellina (ITS–28S–18S of C157023 and 28S–18S of C125384), Clathrosporium (28S of CBS 115285), Filosporella fistucella (18S–ITS–28S–EF1–RPB1 of CCMF-13091), and Hydrocina chaetocladia (18S–ITS–28S of HME4375) to infer their phylogenetic history onto the Leotiomycetes v. 1 reference tree from Haelewaters et al. (2021), which already includes ‘V.’ delicatum (CCMF-19494) and Xerombrophila crystallifera (TAAM 122762). We used the “de novo” option for the IQ-TREE placement, under partitioned models, with GTRGAMMA as substitution model for each partition. Support for each branch was estimated by 1000 ultrafast bootstrap (UFboot2) replicates (Hoang et al. 2018) and the Shimodaira-Hasegawa-like approximate likelihood ratio test (SH-aLRT) (Guindon et al. 2010). Neurospora crassa and Xylaria hypoxylon were used as outgroups.

To investigate the phylogeny of Ascocoryne and Gelatinodiscaceae, newly generated sequences were combined with sequences obtained from GenBank, UNITE, and iNaturalist.org. We focused on isolates with multilocus data and representatives from different named and unnamed species- and genus-level clades. For the genus Ascocoryne, we also focused on isolates for which the CTP locus was sequenced or could be extracted from a genome assembly. Metadata for all sequences used are provided in Table S1.

We constructed maximum likelihood (ML) and Bayesian trees from five different sets of sequences. First, we included all seven loci (18S–ITS–28S–EF1–RPB1–RPB2–CTP) and the entire Gelatinodiscaceae dataset with Arachnopeziza araneosa (Arachnopezizaceae), Blumeria graminis and Golovinomyces cichoracearum (Erysiphaceae), and Lemonniera aquatica and Articulospora tetracladia (Discenellaceae) as outgroup taxa. Second, these same seven loci for only species in the genus Ascocoryne, using A. turficola as the outgroup (Gelatinodiscaceae phylogenies recovered A. turficola basal to the rest of Ascocoryne). Third and fourth, the same datasets above but limited to isolates for which at least two loci were present. Fifth, only species in the genus Ascocoryne using only ITS sequences, with A. turficola as outgroup. In the text, we refer to the first two groups as ‘all-taxa’ trees and the third and fourth groups as ‘2-minimum’ trees.

For the Ascocoryne and Gelatinodiscaceae trees, sequences from each locus were aligned using MAFFT in Geneious (Katoh et al. 2002, Katoh & Standley 2013). The ITS alignment was trimmed to begin after the 5’ starting motif 5’–ATCATTA–3’ and to end at the 5’ start of the 28S at 5’–TGACCT–3’. Similarly, 28S sequences were trimmed to begin at the 5’–TGACCT–3’ motif. After manual trimming, all alignments were further trimmed in the command-line version of TrimAl v. 1.3 (Capella-Gutiérrez et al. 2009) using a gap threshold (−gt) of .60 and minimum coverage (−cons) of .50, with other settings on default. Concatenation of loci was completed in SequenceMatrix (Vaidya et al. 2011). Aligned, trimmed, and concatenated (if relevant) alignments are provided in Supplementary Files S3–S7. Because we primarily applied a phylogenetic species concept (with additional morphological and geographical data) to delimit taxa and estimate species richness in Ascocoryne, we also built single-locus trees to check for discordance between loci, following Genealogical Concordance Phylogenetic Species Recognition (Taylor et al. 2000). We found one instance where the 28S, EF1, and CTP loci placed PULF29998 in the group of Ascocoryne species with inflated paraphyses (where it was inferred to belong based on morphology), while the ITS locus placed it distant to these taxa. After confirming the ITS sequence was correct with reextraction and resequencing, all sequences were included in our concatenated analysis.

Maximum likelihood trees were inferred in IQ-TREE (Nguyen et al. 2015) with partitioned models (Chernomor et al. 2015) and 1000 bootstrap (BS) replicates. Within IQ-TREE, ModelFinder (Kalyaanamoorthy et al. 2017) was used to determine the best model at each partition using the Bayesian Information Criterion (BIC). Bayesian trees were inferred using BEAST2 v. 2.7.7 (Bouckaert et al. 2019) and associated applications (BEAUTi, LogCombiner, TreeAnnotater) and Tracer v. 1.7.2 (Rambaut et al. 2018). In all cases, BEAST2 was run with unlinked site and clock models, and a linked tree model. Ambiguities were considered (box checked), bModelTest (Bouckaert & Drummond 2017) was used for the site model from the model set ‘transitionTransversionsplit’ and mutation rates were estimated. An optimized relaxed clock (Douglas et al. 2021) and the birth death model were used. Two initial runs were completed with a chain length of 10 M, after which the operators were tuned based on suggested outputs. With operator-tuned input files, five independent runs with a chain length of 100 million generations were completed for each tree set. Burn-in was adjusted to optimize effective sample sizes by viewing and editing both individual and combined traces of the log files in Tracer. Trees were combined in LogCombiner and TreeAnnotator was used to generate the maximum clade credibility tree using the mean for node heights. Tree topologies of ML and Bayesian trees were visualized in FigTree v. 1.4.3 (http://tree.bio.ed.ac.uk/software/figtree/) and edited in Inkscape v. 1.2.2. Information on alignments and models selected for each methodology is provided in Table S2.

Barcode analysis

We used three different DNA barcoding methodologies to compare the ITS locus to the CTP locus in identifying a priori-identified Ascocoryne species: distance-based (e.g., barcode gap), tree-based (e.g., phylogenetic), and character-based (e.g., diagnostic nucleotide positions in alignment). We examined six species for which we had at least two sequences of both ITS and CTP loci: A. albida (2 sequences), A. aff. cylichnium 1 (9), A. aff. cylichnium 2 (3), A. fibrosa (3), A. sarcoides (7), and A. wilsoniae (5). We included all other Ascocoryne CTP sequences available and their corresponding ITS sequences, resulting in 37 total sequences of 14 different species at each locus. For each method, we first aligned all the sequences at each locus using MAFFT in Geneious (Katoh et al. 2002, Katoh & Standley 2013). We trimmed any regions outside the two flanking primers for the CTP locus and trimmed the ITS as described above for the phylogenetic analysis. Finally, we removed any sequences less than one half the length of the total alignment, which resulted in the removal of two A. aff. cylichnium 1 sequences (PULF29777 and PULF29778) from the ITS dataset; no sequences were removed from the CTP dataset. The CTP alignment (Supplementary File S8) was 698 nucleotides in length and the ITS alignment (Supplementary File S9) was 531 nucleotides in length.

For the distance-based method, we used the MEGA v. 11 (Tamura et al. 2021) distance estimation tool, selecting ‘none’ for variance estimation method, ‘p-distance’ for the model/method, “d: Transitions + Transversions” for the substitutions to include, ‘Uniform Rates’ for rates among sites, and “Pairwise Deletion for ‘Gaps/Missing Data Treatment.” If species had a ‘barcode gap’ (Meyer & Paulay 2005), or a minimum interspecific distance greater than the maximum intraspecific distance, they were considered successfully identified via this method (Hebert et al. 2004).

For the tree-based method, we used IQ-TREE v. 1.6.12 (Nguyen et al. 2015) and the built-in ModelFinder Plus (Kalyaanamoorthy et al. 2017) to construct ML phylogenies with 1000 ultrafast bootstrap replicates (Hoang et al. 2018). If species were recovered in supported (> 90 BS), monophyletic clades, they were considered successfully identified via this method. For the character-based method, we manually examined alignments and counted the number of diagnostic positions, or sites where a single nucleotide or gap was shared by all members of the same species but not by any other species in the alignment. Species were considered successfully identified via this method if they had at least one diagnostic position (Jaén-Molina et al. 2015).

RESULTS

High-throughput sequencing

We obtained correct ITS1, ITS2, and 28S sequences in 32/115 (28 %) attempts from 37 isolates via Illumina sequencing. In some cases, a second elution from the same extraction, or separate extractions from sporocarps and cultures were both used, leading to more than one attempt per isolate. For individual loci, the success rate of ITS1 was 11/41 (27 %), ITS2 16/44 (36 %), and 28S 5/28 (18 %). For different collection types, we obtained correct sequences from extractions of desiccated cultures in 2/15 (13.3 %) attempts and from extractions of apothecia in 30/100 (30 %) attempts. In the 83 out of 115 (72 %) unsuccessful attempts, 64 (77 %) attempts did not contain Gelatinodiscaceae sequences, and 19 (23 %) attempts contained Gelatinodiscaceae sequences, but they were not used because there were multiple or conflicting ASVs, the ASV did not match the morphology of the specimen, or the ASV was present among multiple samples of putatively different species (i.e., potentially contaminants).

Phylogenetic placement of Clathrosporium, Phaeangellina, Skyathea, and Xerombrophila

In our Leotiomycetes-wide phylogeny (Fig. S1), the two Phaeangellina isolates were not in Gelatinodiscaceae and formed a supported (SH-aLRT 100 % / UFboot2 100 %) clade with Cenangium acuum, Piceomphale bulgaroides, and Moellerodiscus pinicola, sister to the Sclerotiniaceae and Rutstroemiaceae, consistent with Johnston & Park (2025). The Skyathea and Xerombrophila isolates were supported (SH-aLRT 100 % / UFboot2 100 %) in the family Gelatinodiscaceae, with isolates of Ascocoryne, Chloroscypha, and Neobulgaria. The Clathrosporium and Hydrocina isolates formed a supported (SH-aLRT 98 % / UFboot2 99 %) clade with Filosporella fistucella and Varicosporium delicatum outside Gelatinodiscaceae. Therefore, in our phylogenetic analyses of Gelatinodiscaceae, Skyathea and Xerombrophila isolates were included, while Clathrosporium and Phaeangellina isolates were excluded.

Gelatinodiscaceae phylogenies

The topologies for the all-taxa tree and 2-minimum tree for Gelatinodiscaceae were nearly identical using both ML and Bayesian methodologies (ML topology of all-taxa tree Fig. 1; 2-minimum tree Fig. S2; unedited trees Supplementary Files S10–S13). All four analyses supported a monophyletic Gelatinodiscaceae and the following clades: 1) the genera Dimorphospora, Helicodendron, Ombrophila, Skyathea, and Xerombrophila, including four isolates — CBS 301.50 2, NY03416357, 203_OA_2013, and ZML09_3 — that occurred in three clades with uncertain generic placement; 2) the genera Ascotremella, Neobulgaria, and AFTOL_ID1259 with an uncertain generic placement; 3) the genera Ascocoryne and Chloroscypha. In all four analyses, isolates of Ascocoryne turficola formed a separate clade between all other Ascocoryne isolates and Chloroscypha.

Fig. 1 — Seven-locus phylogenetic analysis of *Gelatinodiscaceae* (all-taxa tree). The maximum-likelihood topology is displayed with bootstrap support values ≥ 70 and Bayesian posterior probability values ≥ .90 when nodes are shared between trees. Within *Ascocoryne*, clades or individual isolates with morphological data are highlighted and ascospore lipid bodies (LBs) and conidia type, whether paraphyses apices are apically inflated, whether the ectal excipulum is amyloid, and whether calcium oxalate crystals are present is shown. The scale bar indicates the number of expected substitutions per site.

Ascocoryne phylogenies — intrageneric clades and morphology

The four multilocus analyses retrieved a monophyletic, supported Ascocoryne, excluding A. turficola, which was used as an outgroup after Gelatinodiscaceae phylogenies recovered it as basal to the rest of Ascocoryne (ML topology of all-taxa tree Fig. 2; 2-minimum tree Fig. S3; unedited trees Supplementary Files S14–S17). The ITS-only phylogeny (Fig. S4) generally lacked high support values at deeper nodes. It will not be used to discuss clades above the species rank and the individual ML and Bayesian tree files can be found in Supplementary Files S18 and S19.

Fig. 2 — Seven-locus phylogenetic analysis of *Ascocoryne* (all-taxa tree). The maximum-likelihood topology is displayed with bootstrap support values ≥ 70 and Bayesian posterior probability values ≥ .90 when nodes are shared between trees. The four large clades mentioned in the text are shown and clades or individual isolates with morphological data are highlighted and ascospore lipid bodies (LBs) and conidia type, whether paraphyses apices are apically inflated, whether the ectal excipulum is amyloid, and whether calcium oxalate crystals are present is shown. The scale bar indicates the number of expected substitutions per site.

Ascocoryne lilacina was recovered in an unsupported sister position to the rest of the taxa with soil isolates from Estonia and Greenland in our all-taxa trees. All four analyses recovered a clade of A. albida, A. ‘inflata’, A. neoinflata sp. nov., and A. sarcoides, along with several other isolates (clade 1), but this clade only had support in the Bayesian analyses. Morphologically, all these species have conidia that form in chains (or conidia type is unobserved) from ascospores that contain at least one large LB, do not have an amyloid reaction of the ectal excipulum, and do not contain calcium oxalate crystals. Isolates with consistently inflated paraphyses were recovered in a single subclade within clade 1, but this clade was also only supported in Bayesian analyses.

Apart from A. lilacina and clade 1, all remaining taxa in the genus occurred in three clades of a supported (2-minimum tree) or unsupported (all-taxa tree) larger clade, except AJMH-2010, which was retrieved in an unsupported position alone. Clade 2 was supported in all phylogenies and consisted of A. laurisilvae, A. wilsoniae sp. nov., and several other isolates. Shared morphological characteristics of species in this clade are conidia that occur in chains from mature ascospores, the presence of small calcium oxalate crystals, an inamyloid ectal excipulum, and paraphyses without inflated apices.

Clades 3 and 4 formed a larger, supported group in the 2-minimum tree and Bayesian analysis of the all-taxa tree. All apothecial morphs in this group have ascospores with small to medium LBs only and conidia that form directly on the ascospore wall, not in chains. Clade 3 was unsupported in all analyses and isolates recovered in this clade varied in different analyses. In the 2-minimum tree, clade 3 included A. diminutiva sp. nov. and isolates from Guadeloupe, Indonesia, New Zealand, and USA (Hawaiʻi) that have an amyloid ectal excipulum and variable presence of calcium oxalate crystals. In the all-taxa tree, A. trichophora also was recovered in clade 3 with more New Zealand isolates and a collection from Guyana that does not have an amyloid ectal excipulum. In this tree, the Guadeloupe, Indonesia, and USA isolates were placed in clade 4.

Clade 4 was supported in the 2-minimum tree and consisted of A. fibrosa, A. cf. cylichnium, A. aff. cylichnium 1, A. aff. cylichnium 2, along with several other, single isolates. Ascocoryne fibrosa, which does not have crystals and has a variable amyloid reaction of the ectal excipulum (among other divergent characters), was placed sister to the rest of the taxa that all contain crystals and do not have an amyloid ectal excipulum. In the all-taxa tree, clade 4 was unsupported and included the addition of the Guadeloupe, Indonesia, and USA isolates mentioned above. The subclade of A. cf. cylichnium, A. aff. cylichnium 1 and A. aff cylichnium 2 (excluding A. fibrosa and isolates with varying topologies) was supported in the Bayesian analysis.

Ascocoryne — support for described species and species richness

Our four Ascocoryne-wide phylogenies (including all-taxa trees and 2-minimum trees using both ML and Bayesian methodologies) had largely congruent topologies, particularly at supported branches and terminal clades. All described species of Ascocoryne, including those described here and those without formal recognition (A. ‘inflata’, A. aff. cylichnium 1 and 2, and A. cf. cylichnium) were supported in most trees, except for A. albida and A. sarcoides, which were supported in one of four and two of four trees, respectively (Table S3). Depending on the number of isolates used in the phylogeny, anywhere from ~33 (2-minimum tree) to ~50 (all-taxa tree) species-level clades or single isolates were present in Ascocoryne (Figs 1, 2, Supplementary Files S11 and S16)

ITS and CTP barcode analysis

All six Ascocoryne species could be identified by at least one method at the CTP and ITS loci using our barcoding methods (Table 2). Four of six species at the CTP locus and six of six species at the ITS locus were identified by the distance-based method. The average p-distance at the CTP locus was higher (0.046) than at the ITS locus (0.014). CTP and ITS alignments are in Supplementary Files S8 and S9 and MEGA distance matrices for these alignments are in Supplementary Files S20 and S21. Five of six species at the CTP locus and three of six species at the ITS locus were identified using the tree-based method. Tree files for the CTP and ITS phylogenies are in Supplementary Files S22 and S23. Six of six species at the CTP locus and five of six species at the ITS locus were identified using the character-based method. We found each of the six species considered in our analysis had 1–23 (.14–3.30 %) diagnostic characters in the alignment at the CTP locus and 0–25 (.00–4.71 %) at the ITS locus. On average, the CTP locus had 1.65 % of positions in the alignment containing diagnostic characters and the ITS locus had 1.13 %. Overall, the ITS locus worked in 14 of 18 barcoding attempts and the CTP locus in 15 of 18.

Table 2.

Results of ITS and CTP barcoding analysis.

Method	Description	A. albida	A. aff. cylichnium 1	A. aff. cylichnium 2	A. fibrosa	A. wilsoniae	A. sarcoides	Mean
Distance	Barcode gap ITS?	Yes	Yes	Yes	Yes	Yes	Yes	n/a
	p-distance gap ITS	0.006	0.017	0.034	0.015	0.008	0.004	0.004
	Barcode gap CTP?	Yes	No	Yes	Yes	Yes	No	n/a
	p-distance gap CTP	0.072	−0.017	0.059	0.074	0.089	−0.002	−0.002
Phylogeny	Monophyletic and, supported ITS?	No	Yes	Yes	No	Yes	No	n/a
	Monophyletic and supported CTP?	Yes	No	Yes	Yes	Yes	Yes	n/a
Character	> 1 diagnostic character ITS?	Yes	Yes	Yes	Yes	Yes	No	n/a
	Total diagnostic characters ITS	1	1	7	2	25	0	6
	Percent diagnostic in alignment ITS	0.19 %	0.19 %	1.32 %	0.38 %	4.71 %	0.00 %	1.13 %
	> 1 diagnostic character CTP?	Yes	Yes	Yes	Yes	Yes	Yes	na
	Total diagnostic characters CTP	3	1	10	12	20	23	11.5
	Percent diagnostic in alignment CTP	0.43 %	0.14 %	1.43 %	1.72 %	2.87 %	3.30 %	1.65 %

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TAXONOMY

Ascocoryne J.W. Groves & D.E. Wilson, Taxon 16: 40. 1967.

Apothecia *0.8–30 mm diam. × 0.5–6.0 mm high, scattered to gregarious, cespitose or not, sessile to short-stipitate, rarely long-stipitate (A. turficola) or stipe partially buried in substrate (A. aff. cylichnium 2), discoid, cupulate, pulvinate, or turbinate; disc smooth, light to dark pink, purple, or brown, rarely reddish (A. neoinflata) or greenish (A. turficola); margin and receptacle ± concolourous or appearing slightly lighter or darker, with or without a dusty white covering or rarely purplish fibres (A. fibrosa), smooth to wavy, wrinkled, or furrowed, often more pronounced in desiccation. Asci †70–184 × 5.9–12.8 µm, cylindric–clavate, 8-spored, apex rounded to subconical, slightly thick-walled, pore blue in iodine with or without KOH-pretreatment, base arising from croziers, or rarely not (A. cf. cylichnium). Ascospores †6–28.8 × 3.0–6.1 µm, hyaline, cylindrical, ellipsoid, or fusoid, straight to inequilateral or slightly curved, 1–12-septate, or rarely without septa (A. lilacina), rarely with a gelatinous sheath (A. turficola), *containing LBs, 1–2 large (> 2 µm diam., sometimes smaller in A. lilacina) and/or numerous small (< 2 µm diam.), generally filling more than 50 % of the cytoplasm; with or without conidia budding individually from ascospore walls, or in chains from their poles. Conidia †1.3–5.3 × 1.0–3.6 µm, hyaline, globose, ovoid, or subellipsoid, usually with one LB. Paraphyses cylindrical, septate, often branching in lower cells; terminal cell †7.9–41.6 × 1–6 µm uninflated cylindrical to clavate or capitate, *with VBs in the apical cell(s), numerous, small and globose, or fewer, long and cylindrical. Medullary excipulum composed of textura intricata of hyaline hyphae †1–5.5 µm diam., immersed in a thin to abundant gel, with or without calcium oxalate crystals and druses. Ectal excipulum composed of t. globulosa–angularis of brown to purple cells †3.0–27.3 × 2.6–26.3 µm diam., rarely a gelatinized t. intricata (A. fibrosa), with or without calcium oxalate crystals and druses, usually releasing purplish-brownish pigments in KOH, sometimes blue in iodine with or without KOH-pretreatment. Base of ectal excipulum often connected to substrate by a layer of hyaline t. intricata, immersed in abundant gel, sometimes blue in iodine with or without KOH pretreatment.

Conidiomata generally gregarious, often cespitose, sessile to stipitate, discoid, cerebriform, or synnamatous with subglobose to irregular, usually convex apex; smooth, whitish, cream, reddish, pinkish, or light to dark purplish, occurring interspersed with, or beside the sexual morph, or not; covered in conidiophores completely, or only at the differentiated apex; interior cells t. intricata immersed in a thin to abundant gel; sterile exterior cells t. globulosa–angularis. Conidiophores usually branching and containing straight to curved metulae and phialides 5–22 × 1–4 µm to neither (A. albida). Conidia †1.5–4.0 × 1.0–3.5 subglobose to ovoid, elliptical, or allantoid.

Cultures often polymorphic over time and after subculturing; white or purplish (and turning the media purplish), less often beige, cream, or greenish-brown; texture powdery, smooth, slimy, or fluffy; hyphae *1.4–7.0 µm diam., cylindrical, hyaline to purplish, forming branching conidiophores; metulae and phialides *5–18 × 0.9–4 µm, subconical, fusiform, lageniform, or inflated cylindrical; conidia *2–6(–10) × 0.6–3.2 µm, globose, ovoid, allantoid, or cylindrical, sometimes multiple types within a single culture, with 0–2 LBs, sometimes budding directly from hyphae in chains.

Notes: Growth on dead wood, usually well-decayed, rarely on Sphagnum spp. (A. turficola) or leaves (FH00374851), in the autumn in the temperate or boreal northern hemisphere. Broadly distributed in boreal, temperate, and tropical ecosystems.

Ascocoryne aff. cylichnium 1. Figs 3, 4.

Fig. 3 — Morphology of *Ascocoryne* aff. *cylichnium* 1 († and from PULF29781 unless otherwise stated). A. *Ascomata, in-situ. B. *Ascomata in-situ (PULF29777). C. *Ascomata under dissecting microscope in laboratory (PULF29777). D. Section in H₂O. E. *Section in H₂O (PULF29778). F. *Section in H₂O (PULF29777). G. Ectal excipulum margin in KOH. H. Ectal excipulum margin in KOH+CR. I. Ectal excipulum upper flanks in KOH. J. Ectal excipulum upper flanks in KOH+CR. K. Ectal excipulum lower flanks in KOH. L. Ectal excipulum lower flanks in KOH+CR. M. Ectal excipulum base in KOH+MLZ. N. Ectal excipulum base and hyphae connecting to substrate in MLZ. O. Basal hyphae connecting to substrate in KOH+CR. P. Medullary excipulum in KOH+MLZ (PULF29777). Q. *Ascus in H₂O (PULF29777). R. *Ascus in H₂O. S. *Ascus in H₂O (PULF29778). T. Asci in MLZ. U. Asci in MLZ (PULF29777). V. Croziers in KOH+CR. W. *Paraphyses in H₂O (PULF29992. X. *Paraphyses in H₂O (PULF29778). Y. *Paraphysis in H₂O (PULF29774). Z. Paraphysis in KOH. **AA.** Paraphyses in KOH (PULF29774). **AB.** *Ascospores in H₂O. **AC.** *Ascospore in H₂O (PULF29777). **AD.** Ascospores in H₂O (PULF29777). **AE.** Ascospores in MLZ (PULF29777). **AF.** Ascospores in MLZ. Scale bars: A–C = 1 mm; D–F = 100 µm; G–AF = 10 µm.

Fig. 4 — Cultures of *Ascocoryne fibrosa* (A–I; PULF29773), *Ascocoryne wilsoniae* (J–S; PULF29771), and *Ascocoryne* aff. *cylichnium* 1 (T–AB; PULF29774) after 30 d on PDA unless otherwise stated. **A, B.** *Culture colony. **C, D.** *Conidiophores in H₂O. E. *Conidiophore in IKI (50 d). F. †Hyphae of colony in KOH. G. *Conidia in H₂O. H. *Conidia in IKI (50 d). I. †Conidia in KOH. **J, K.** *Culture colony. **L, M.** †Conidiophores in KOH (50 d; Rose Bengal). N. *Conidiophores in H₂O (50 d; Rose Bengal). O. *Hyphae of colony in H₂O (d). P. *Conidia from conidiophores in H₂O. Q. *Conidia from conidiophores in H₂O (50 d on Rose Bengal). **R, S.** *Conidia budding directly from hyphae in H₂O. **T, U.** *Culture colony. **V, W.** *Conidiophores in H₂O (44 d, PULF29785). X. *Conidiophore in IKI (44 d). Y. *Hyphae of colony hyphae in H₂O. **Z–AB.** *Conidia in H₂O (AD and AE 50d on Rose Bengal). Scale bars: A, B, J, K, T, U = 10 mm; C–I, L–S, V–AB = 10 µm.

Apothecia *2–12.2 mm diam. × 2–6 mm high, scattered to gregarious, often cespitose, cupulate-stipitate to turbinate, subsessile, usually broadly attached to substrate; disc smooth, light purple (11E–F5 to 12F5–7), margin differentiated, often with conspicuously darker purple line; receptacle ± concolourous to disc, often appearing lighter due to whitish dusky surface, particularly closer to margin of the receptacle. Asci †(141–)147–168(–176) × (7.4–)8.1–9.7(–10.8) µm, cylindric-clavate, 8-spored, ascospores *biseriate, apex rounded to subconical, slightly thick-walled, pore 1.6–2.6(–3.6) × 1.1–2.0 µm, blue with or without KOH-pretreatment, base arising from croziers. Ascospores *(20.3–)22.8–26.8(–30.5) × (4.4–)5.1–6.1(–7.0) µm, †(18.9–)20.7–24.4(–27.6) × (3.3–)4.2–5.2(–5.6) µm, hyaline, cylindrical-fusiform to fusoid, inequilateral, slightly helicoid, containing numerous small (*< 2 µm) LBs completely filling the cytoplasm (LCI 5) coalescing into large guttules in dead ascospores, when mature up to 5-septate, overmature ascospores budding conidia from ascospore walls. Conidia *(1.3–)1.7–2.6(–3.1) × (1.2–)1.5–2.1(–2.2) µm, †1.3–2.0(–2.3) × (1.0–)1.2–1.7(–1.9) µm, globose to subellipsoid with one LB, hyaline. Paraphyses cylindrical, apically slightly to medium clavate, septate, terminal cell †(21.2–)23.7–35.0(–41.6) × (2.0–)2.2–2.7(–2.9) µm, lower cells †(15.0–)18.0–24.4(–27.5) × 1.1–1.7(–2.1) µm *with grey or colourless VBs in the final, or sometimes final two apical cells, usually numerous, globose, and small (0.4–2.0 µm diam.), less commonly cylindrical (up to 6 µm in length). Medullary excipulum of hyaline t. intricata, †139–723 µm thick, hyphae †1.0–2.8 µm, *2.7–4.0(–7.0 µm) diam., slightly gelatinized, with small to usually large calcium oxalate crystals throughout, (9–)12–56 µm diam., often aggregating into clusters up to 104 µm diam. Ectal excipulum a purplish t. globulosa–angularis, †51–145 µm thick, cells †4.8–12.6(–17.6) × (3.3–)3.9–9.3(–13.6) µm, generally larger at base and interior region, smaller in exterior region and towards margins where a palisade of cylindrical to angular hyphae is formed terminating in clavate to subcapitate cells, with crystals as the medullary excipulum; pigments dissolving in KOH leaving ectal excipulum light brownish-yellowish. Base of ectal excipulum covered by another layer of t. intricata composed of loosely packed hyphae, strongly gelatinized, that connects the ascomata to the substrate, this layer is blue in IKI with or without KOH pretreatment. Asexual morph unknown.

Culture characteristics: On PDA growing slowly, 10–20 mm diam. after 30 d, greenish brown (3C4–4D4) towards center, cream then white towards leading edge, center fluffy in appearance, more slimy towards edge; hyphae *1.4–3 µm diam., cylindrical, hyaline, producing ellipsoid-subglobose conidia *(2.0–)2.4–3.2(–4.4) × (1.4–)1.7–2.4(–3.2) µm, usually containing one small (< 1 µm) LB, borne on subconical to inflated cylindrical phialides, *5.1–10.8(–18.0) µm long, arising from a bifurcating, hyphal base, with or without inflated cylindrical metulae, similar sized to phialides.

Distribution: Known from the midwestern USA (Indiana, Michigan), eastern Canada, and India.

Materials examined: Canada, Nova Scotia, Bathurst, on Abies balsamea, ‘butt rot, entered by roots white feather’, 13 Aug. 1951, V. Nordin B1641-1 (DAOM33818). India, Himachal Pradesh, Kulu, Pulga on dead angiosperm stump, 30 Sep. 1965, K.S. Thind 3124 (K-M1434234). USA, Michigan, Waterloo Recreation Area, Spring Pond Trail, on decorticated wood, 2 Oct. 2021, J.K. Stallman 376 (PULF29765); Michigan, Waterloo Recreation Area, Spring Pond Trail, on decorticated wood, 2 Oct. 2021, J.K. Stallman & J. Liu 377 (PULF29766); Indiana, Parke County, on decorticated wood, 16 Oct. 2021, J. Liu 381 (PULF29767; Indiana, Tippecanoe County, Ross Biological Reserve, on decorticated angiosperm wood, 23 Oct. 2021, J. Peterson RR307 (PULF29769; Indiana, Brown County, Yellowwood State Forest, 39.1324, −86.3378, on decorticated wood, 1 Oct. 2022, J.K. Stallman & C. Brito 409 (PULF29774), culture DAOMC 252831; ibid., on decorticated angiosperm wood, 15 Oct. 2022, 413 (PULF29777). Ibid., 414 (PULF29778); ibid., 418 (PULF29781); ibid., 419b (PULF29783); Indiana, Brown County, Yellowwood State Forest, on decorticated angiosperm wood, 39.1329, −86.3401, 7 Oct. 2023, J.K. Stallman 427 (PULF29784); ibid., 428 (PULF29785); ibid., 429 (PULF29786); ibid., 430 (PULF29787); ibid., 431 (PULF29788); ibid., on decorticated Platanus occidentalis, 12 Oct. 2024, 454 (PULF29989); ibid., on decorticated Quercus, 455 (PULF29990); ibid., on angiosperm wood, 456 (PULF29991); ibid., 457 (PULF29992); ibid., 458 (PULF29993); Indiana, Tippecanoe County, Purdue Horticulture Park, 40.4279, −86.9352, on decorticated Quercus sp., 19 Oct. 2024, J.K. Stallman 460 (PULF29994); ibid., 461 (PULF29995); ibid., 462 (PULF29996); Indiana, Brown County, Yellowwood State Forest, 39.1321, −86.3410, on decorticated Quercus alba, 6 Nov. 2024, J.K. Stallman 464 (PULF29997); ibid., on angiosperm wood, 467 (PULF30000); ibid., 468 (PULF30001); ibid., 469 (PULF30002); ibid., 470 (PULF30003); ibid., on decorticated Quercus alba, 471 (PULF30004); ibid., on angiosperm wood, 472 (PULF30005); ibid., 39.1316, −86.3382, 476 (PULF30009); ibid., 39.1321, −86.3410, on decorticated Quercus alba, 477 (PULF30010); ibid., on decorticated Platanus occidentalis, 478 (PULF30011); ibid., on decorticated Acer saccharum 479 (PULF30012); ibid., on decorticated Quercus alba, 480 (PULF30013); ibid., 39.1316, −86.3382, on decorticated angiosperm wood, 480 (PULF30014); Illinois, Vermillion County, Kickapoo State Recreation Area, 40.1547, −87.7366, on decorticated Quercus sp., 9 Nov. 2024, J.K. Stallman 484 (PULF30017); ibid., on decorticated Quercus alba, 485 (PULF30018); ibid., 486 (PULF30019); on decorticated Catalpa speciosa 487 (PULF30020); Indiana, Tippecanoe County, Ross Biological Reserve, 40.409, −87.0654, on decorticated angiosperm wood, 1 Nov. 2024, J.K. Stallman 490 (PULF30023); ibid., 491 (PULF30024); ibid., 492 (PULF30025); ibid., 493 (PULF30026; ibid., 497 (PULF30027; ibid., 499 (PULF30028).

Notes: Ascocoryne aff. cylichnium 1 is consistent with the broad concept of A. cylichnium: it has long ascospores filled with medium to small LBs when living that develop many septa when overmature. Conidia bud directly from the ascospore wall, not in chains, the medullary excipulum is not particularly gelatinized, and calcium oxalate crystals are abundant. It is the most common Ascocoryne species found in our collecting locations in Illinois and Indiana and is also found in Michigan. Until the name A. cylichnium can be stabilized with type studies and detailed morphological and/or molecular data, we do not suggest describing new Ascocoryne species with overlapping biometric characteristics with this taxon (see Discussion).

Ascocoryne aff. cylichnium 2. Fig. 5.

Apothecia *3–9 mm diam. × 2–5 mm high, gregarious, usually cespitose, discoid-subcupulate or turbinate–petuloid, particularly when growing in clusters, subsessile, usually broadly attached to substrate and with rudimentary stipe immersed in substrate; disc smooth, pits or folds developing in age, light purple (11E–F5 to 12F5–7) to purple, margin differentiated, often with conspicuously darker purple line at margin; receptacle ± concolourous to disc, often appearing lighter due to whitish dusky surface, particularly closer to margin of the receptacle. Asci †(149–)156–172(–177) × (7.7–)8.5–9.9(–11.3) µm, cylindric-clavate, 8-spored, ascospores †obliquely biseriate to uniseriate in lower part, apex subconical, sometimes subtruncate, slightly thick-walled, pore †(1.3–1.6–2.4(–2.8) × (0.9–)1.1–1.5 µm, blue with or without KOH-pretreatment, base arising from croziers. Ascospores *(20.1–)25.2–29.9(–31.7) × (4.6–)5.2–6.0(–6.6) µm, †(17.3–)22.2–26.3(–28.8) × (4.2–)4.7–5.5(–6.0) µm, hyaline, cylindrical-fusiform to fusoid, inequilateral, containing numerous small (*< 2 µm) LBs completely filling the cytoplasm (LCI 5) coalescing into large guttules in dead ascospores, when mature up to 5-septate, overmature ascospores budding conidia from ascospore walls. Conidia †(1.5–)1.8–2.6(–3.4) × (1.0–)1.3–2.0(–2.6) µm, globose to ellipsoid or ovoid with one LB, hyaline. Paraphyses uninflated cylindrical, rarely apically slightly clavate, septate, terminal cell †(13.8–)14.6–37.7(–52.1) × (1.6–)1.7–2.5(–2.8) µm, lower cells †(7.9–)11.5–24.1(–26.2) × (0.9–)1.0–1.7(–2.6) µm, usually simple, but sometimes branching, slightly exceeding the asci. Medullary excipulum of hyaline t. intricata, †85–363 µm thick, lower regions near base with thinner hyphae 1.2–3.8 µm diam., somewhat gelatinized, upper regions of thicker hyphae 2.7–6.8 µm diam., not gelatinized, with small to large (2–)5–19(–40) µm diam. calcium oxalate crystals throughout, often aggregating into clusters up to 96 µm diam. Ectal excipulum composed of a brownish t. globulosa–angularis, †41–153 µm thick, cells †(3.0–)4.5–9.7(–12.7) × (2.6–)3.3–5.8(–8.1) µm, containing calcium oxalate crystals, at margin a palisade of cylindrical to angular hyphae with a cortical layer formed of slightly protruding clavate cells with crystals; pigments mostly dissolving in KOH leaving ectal excipulum light brownish. Asexual morph unknown. Culture characteristics unknown.

Distribution: Thus far only known from Illinois, USA.

Materials examined: USA, Illinois, Vermilion County, Kickapoo State Park, 40.145, −87.734, on decorticated angiosperm wood among moss, 21 Oct. 2023, J.K. Stallman & C. Brito 440 (PULF29791); ibid., 441 (PULF29792); ibid., 40.144, −87.7333, 447 (PULF29795).

Notes: Ascocoryne aff. cylichnium 2 is genetically differentiated from A. aff. cylichnium 1 (clade 4, Fig. 2) although many biometric features such as asci, ascospores, and paraphyses overlap. The submerged stipe, larger conidia, smaller excipular cells on average, and perhaps additional cultural or vital taxonomic features may be useful in differentiating A. aff. cylichnium 2 from A. aff. cylichnium 1, but more data are needed. Ascocoryne aff. cylichnium 2 is only known from three collections made on the same day in 2023 from Kickapoo State Park in Illinois, USA. Due to the resemblance to A. aff. cylichnium 1, this species was only recognized to be distinct after DNA sequencing and thus detailed microscopy of living cells and culturing were not performed. Attempts to recollect this taxon and obtain additional data in 2024 failed. See the Discussion for further information on A. cylichnium and similar species.

Ascocoryne diminutiva Stallman, Quijada, & Aime, sp. nov. MB 861572. Fig. 6.

Etymology: The specific epithet refers to the small size of the apothecia and spores.

Diagnosis: Differs from A. cylichnium and A. javanica by smaller ascospores and ascomata. Additionally, differs from A. cylichnium by the blue ectal excipulum in IKI and from A. javanica by having few, small calcium oxalate crystals, not present in subhymenium.

Apothecia 1.0–3.0(–5.1) mm diam. × 0.5–2.0(–3.8) mm high, solitary or scattered to gregarious and cespitose, when immature discoid or slightly concave, later shallowly cupulate to turbinate, slightly gelatinous, disc smooth, translucent, whitish-pink (12A2–B2) to light purple-pink (12B3–C3); margin round to slightly wavy, concolourous, smooth to slightly crenate; receptacle concolour to light pinkish-purple (lighter than 11A2–12B2) or light purplish brown (12C3–12E4); shortly to medium stipitate, stipe 0.3–3.2 × 0.3–3.0 (wider if fused with adjoining apothecia), tapering toward the base or not. Asci †(89–)101–119(–124) × (6.3–)7.0–9.7(–12.2) µm, cylindric-clavate, 8-spored, spores obliquely biseriate to uniseriate in lower part; apex rounded to conical, slightly to thick-walled, pore (1.3–)2.0–3.2(–4.0) × 1.2–1.8 µm, blue with or without KOH-pretreatment, base arising from croziers. Ascospores †(11.9–)13.2–15.3(–16.5) × (3.3–)3.7–4.4(–5.0) µm, hyaline, subcylindrical to fusiform-clavate, inequilateral, slightly curved in side view, 0–3-septate, containing numerous small LBs (LCI 5) coalescing into large guttules when dead, overmature spores budding conidia directly from the ascospore wall. Conidia †1.6–2.2 × 1.2–1.8 µm, globose-subglobose to ovoid, containing one guttule (LB). Paraphyses cylindrical, apically medium clavate to slightly moniliform, septate, terminal cell †(16–)20–27.5(–30) × (3–)3.5–4.5 µm, lower cells †(8–)10.5–16.5(–22) × 1–2 µm, hyaline, simple or branching near lower cells. Medullary excipulum of t. intricata, †160–275 µm thick, hyaline, pinkish close to the subhymenium, cells loosely packed, mildly gelatinized, cells †1–2(–3.5) µm diam., not reacting in IKI, small (2–5 µm diam.) calcium oxalate crystals present, more frequent at base near ectal excipulum, not aggregating into large clusters. Ectal excipulum yellowish–pink, pigments dissolving in KOH colouring medium purplish; weakly to medium blue in IKI. From base to mid flanks made of t. globularis–angularis–prismatica, 81–175 µm thick, cells (10–)11.5–14.5(–18.5) × (6.5–)7–9(–12) µm, containing a few to many calcium oxalate crystals 2–5(–15) µm diam., not aggregating; outermost layer from margin toward upper flanks of t. globularis–angularis, 27–60 µm thick, cells (5–)5.5–6.5(–7) × (4–)4.5–6.5(–7) µm, usually with protruding, globose-clavate, thin-walled cells and covered by hyaline or yellowish amorphous resin; inner cells with yellowish walls up to 1.2 µm thick. Base of ectal excipulum covered by another layer of t. intricata composed of loosely packed hyphae, strongly gelatinized, that connects the ascomata to the substrate, this layer reacts blue in IKI with or without KOH pretreatment. Asexual morph unknown. Culture characteristics unknown.

Distribution: Known from Guyana and Puerto Rico.

Typus: Guyana, Upper Potaro River Basin, Pakaraima Mountains, base camp, 5.3013, −59.9112, ~710 m a.s.l., on unknown wood, 16 Jun. 2016, M.C. Aime 6191 (holotype BRG43301; isotype PULF29799). GenBank: ITS = PQ483814, 28S = PQ483853, 18S = PQ483887, EF1 = PQ505780, RPB1 = PQ505758, RPB2 = PQ505752.

Additional materials examined: Guyana, Upper Potaro River Basin, Pakaraima Mountains, Dicymbe Plot 1, 5.2763, −59.9161, ~710 m a.s.l., on unidentified wood, 30 Jun. 2002, M.C. Aime 2180 (BRG43302, PULF29798). USA, Puerto Rico, Maricao, Maricao Fish Hatchery, ~460 m a.s.l., on rotten log, 16 Jun. 1970, R.P. Korf et al. (CUP4158); Río Grande, El Yunque, Bisley Trail, on angiosperm wood, 7 Jul. 2002, K.T. Hodge et al. (CUP-PR-004327).

Notes: Ascocoryne diminutiva morphologically fits into the broad concept of A. cylichnium with ascospores containing smaller LBs only and conidia budding directly from the ascospore wall, not in chains, but differs in several other characteristics. Ascocoryne diminutiva has small calcium oxalate crystals that do not aggregate into large clusters, smaller ascospores that are 0–3-septate, and a blue reaction of the inner layer of the ectal excipulum in IKI which is composed of yellowish thick-walled cells. Descriptions of A. cylichnium have larger crystals, overmature ascospores with five or more septa, and do not note an amyloid ectal excipulum (e.g., Quijada et al. 2017). Ascocoryne diminutiva differs from A. javanica, which also has an amyloid ectal excipulum (per notes from Richard P. Korf associated with syntype collection CUP-K-4733, T. Iturriaga pers. comm.), by lacking large crystals in general and any crystals in the subhymenium and upper medullary excipulum (vs abundant crystals in A. javanica, Penzig & Saccardo 1902). Additionally, A. diminutiva has smaller ascospores [†13.2–15.3(–16.5) µm vs †19–21 µm] and ascomata (apothecia up to *5.1 mm diam. vs up to 20 mm diam.). Ascocoryne lilacina has an amyloid ectal excipulum (L. Quijada, pers. obs.), but the ascospores are shorter [†6–8(–9) µm] and do not become septate when mature (Dennis 1971).

Bulgaria similis was described from Cuba (Berkeley & Curtis 1868) and later synonymized with Ascocoryne sarcoides (Dennis 1954). Notes from Kent Parsons Dumont associated with slides and preserved sections of the ex-K holotype of B. similis (Wright 189), NY03417477, state the spores are 12–13 × 4–5 µm (vs. 10 × 5 µm long in Berkeley & Curtis 1868 and 13–17 µm long in Dennis 1954), and 1-septate. Examination of prepared slides of this collection revealed degraded tissue that did not yield useful morphological characteristics. The spores of A. diminutiva differ in being up to 3-septate and having conidia of the cylichnium-type, budding from the ascospore walls, not in chains. In synonymizing B. similis with A. sarcoides, Dennis (1954) stated he was, ‘unable to separate this [B. similis] from European material of C. sarcoides.’ Because Dennis (1956) considered conidia budding from the ascospore wall to be a key feature of A. cyclichnium, it is unlikely he would make this statement if B. similis had spore characteristics that more resembled A. cyclichnium than A. sarcoides. Therefore, we do not believe B. similis is an earlier synonym of A. diminutiva. Destructive sampling was not pursued of any material at NY, FH, or K because the holotype collection at K has already been sampled for genome sequencing (L. Davies pers. comm) that, if successful, will help future work on this species.

Ascocoryne fibrosa Stallman & Quijada, sp. nov. MB 861571. Figs 4, 7.

Fig. 7 — Morphology of *Ascocoryne fibrosa* (from PULF29773 unless otherwise stated). A. *Ascomata in-situ (PULF29772). B. *Ascomata in-situ. C. *Ascomata under dissecting microscope (PULF29772). D. †Ascomata desiccated under dissecting microscope (PULF29772). E. †Ascomata rehydrated under dissecting microscope (PULF29772). F. †Section in H₂O. G. †Section in KOH. H. †Section of outer ectal excipulum and receptacle (PULF29770). I. †Ectal excipulum margin in KOH+MLZ. J. †Ectal excipulum margin in KOH. K. *Ectal excipulum upper flanks in H₂O (PULF29772). L. †Ectal excipulum upper flanks in KOH+MLZ. M. †Ectal excipulum mid flanks in MLZ (PULF29779). N. †Ectal excipulum lower flanks in KOH+MLZ. O. †Ectal excipulum base in H₂O. P. †Ectal excipulum base in MLZ (PU F29772). Q. †Medullary excipulum in KOH. R. *Medullary excipulum in H₂O (PULF29772). S. *Asci in H₂O (PULF29779). T. †Ascus in MLZ (PULF29772). U. †Asci in MLZ. V. †Ascus in KOH+CR. W. †Croziers in KOH. X. †Croziers in KOH+CR. Y. *Paraphyses in H₂O. Z. †Paraphyses in KOH (PULF29772). **AA.** †Paraphyses in KOH+CR **AB.** *Ascospores in H₂O (PULF29772). **AC.** †Conidia in KOH. **AD.** †Ascospore in H₂O. **AE.** †Ascospore in KOH. **AF.** †Ascospore in (PULF29779). **AG.** †Ascospore in KOH+MLZ. Scale bars: A–E = 1 mm; F–H = 100 µm; I–AA = 10 µm; AB–AG = 5 µm.

Etymology: The specific epithet refers to the ectal excipulum covering of purplish-brown fibers.

Diagnosis: Differs from other Ascocoryne species by its ectal excipulum composed of cylindrical hyphae immersed in a gel and a receptacle covered with groups of protruding cells that macroscopically look like fibers.

Apothecia *2–5.5 mm diam. × 2.5–3.5 mm high, scattered to gregarious (cespitose), cupulate to flat or slightly turbinate when mature, substipitate to shortly stipitate, fleshy to slightly gelatinous; disc light purple to purple (12F8, 14F4–5 to 16F4–5); margin concolourous to lighter with whitish tinge, smooth to broadly undulating; receptacle concolourous or with lighter, purplish–orange colouration (~9D6), often darkening at base, covered in fibres, often aggregating into clumps, sometimes disappearing when overmature; stipe 0.1–1.2 × 0.5–1 mm tapering towards base. Asci †(124–)139–154(–165) × (5.9–)7.4–8.8(–10.0) µm, cylindric-clavate, 8-spored, *obliquely biseriate, apex rounded to subconical, slightly thick-walled, pore 0.9–1.4 × 1.1–1.8 µm, blue with or without KOH-pretreatment, base arising from croziers. Ascospores *(14.1–)16.2–19.2(–22).0 × (4.6–)5.3–6.1(–6.8) µm, †(10.8–)11.7–14.6(–16.9) × (2.8–)3.3–4.1(–4.8) µm, hyaline, ellipsoid to cylindrical-fusoid, inequilateral, multiguttulate, LBs numerous, mostly small (*< 1 µm) and a few medium (*up to 2.4 µm), filling the cytoplasm (LCI 5), coalescing into large LBs in dead spores, 0–3-septate, overmature spores usually forming conidia only at the poles, not in chains. Conidia* 2.1–3.1 × 1.7–2.4 µm, †1.7–2.4 × 1.2–1.7 µm, globose–subglobose to ovoid, hyaline, containing one LB. Paraphyses cylindrical uninflated, straight to slightly wavy, septate, terminal cell †(22.6–)27.7–44.0(–44.9) × 1.8–2.4(–2.7) µm, lower cells †(11.2–)14.1–24.4(–33.9) × (1.2–)1.4–2.0(–2.4) µm, hyaline, usually simple but sometimes branching at lower cells, *with globose (1–2 µm diam.) to cylindrical grey or colourless VBs (–4 µm diam.). Medullary excipulum of t. intricata †181–416 µm thick, mildly to moderately gelatinized, hyphae hyaline to light brownish, †1.0–2.3(–3.5) µm diam., becoming darker and packed tighter closer to subhymenium and looser towards ectal excipulum, not reacting in IKI, releasing purplish pigment in KOH, calcium oxalate crystals absent. Ectal excipulum of t. intricata †62–108 µm thick, thicker near base and tapering to margins, hyphae hyaline, weakly amyloid with or without KOH pretreatment (reaction not always observed), very loosely packed, cells cylindrical, †0.8–1.6(–2.5) µm diam., strongly gelatinized, structure consistent from margin to base, often poorly differentiated from medullary excipulum, particularly near base. Outermost layer differentiated, cells packed tighter, †1.0–2.2 µm diam., ± parallel to apothecial margin, more pigmented and strongly gelatinized, sometimes aggregating into tangled masses up to 40 µm diam. of protruding groups of cells, releasing purplish pigments in KOH; base of ectal excipulum covered by another layer of t. intricata composed of loosely packed hyphae, strongly gelatinized, that connects the ascomata to the substrate, this layer is blue in IKI with or without KOH pretreatment. Asexual morph unknown.

Culture characteristics: On PDA growing slowly, 8–15 mm diam. after 30 d, beige to tan (5B3–5C4) at center, lightening to white at leading edge, colony smooth, firm but with slimy appearance; hyphae *1.4–3.6 µm diam., cylindrical, hyaline; producing variable-shaped conidia, primarily allantoid *(3.8–)4.2–5.1(–5.9) × (0.9–)1.1–1.5(–1.8) µm or ellipsoid–ovoid–oblong *(3.2–)3.7–4.7(–5.7) × (1.6–)1.9–2.5(–2.9) µm, containing several small (≤ 1.1 µm) LBs, borne on inflated cylindrical to subconical phialides (5.7–)6.4–10.7(–14.4) µm long, with or without inflated cylindrical metulae (5.1–)5.8–8.2(–8.6) µm long.

Distribution: Known from Illinois and Indiana, USA.

Typus: USA, Indiana, Yellowwood Forest, 39.1324, −86.3378, on decorticated angiosperm wood, 1 Oct. 2022, J.K. Stallman 408 (holotype PULF29773; ex-type culture DAOMC 252830). GenBank: ITS = PQ483805, 28S = PQ483845, 18S = PQ483882.

Additional materials examined: USA, Indiana, Tippecanoe County, Ross Biological Reserve, on decorticated angiosperm wood, 23 Oct. 2021, K. Maley RR314 (PULF29768); Indiana, Parke County, on angiosperm stump, 7 Nov. 2021, J.K. Stallman 389 (PULF29770); Indiana, Brown County, Yellowwood State Forest, 39.1325, −86.3375, on decorticated angiosperm wood, 1 Oct. 2022, J.K. Stallman 407 (PULF29772); ibid., 39.1324, −86.3378, 15 Oct. 2022, 415 (PULF29779); Illinois, Vermillion County, Kickapoo State Recreation Area, 40.1540, −87.7364, on decorticated Quercus sp., 9 Nov. 2024, J.K. Stallman 482 (PULF30015).

Notes: Ascocoryne fibrosa is characterized by a gelatinized ectal excipulum and receptacle with brownish-purplish fibers. The ascospores have small LBs only and conidia are usually produced from the ascospore poles, but not in chains. Crystals are absent. Although we found the amyloid reaction of the ectal excipulum (present in PULF29779, absent in all other collections) to be variable, the gelatinized ectal excipulum of t. intricata is completely unique within Ascocoryne. Coryne hirneolus is described as having a furfuraceous-granular exterior that might resemble the receptacle fibers of A. fibrosa, but the ascospores of C. hirneolus are wider (and on average longer) than A. fibrosa, even when extreme values are included: †15–16 × 5–6 vs 10.8–16.9 × 2.8–4.8 µm (Saccardo et al. 1889).

Ascocoryne laurisilvae A. Mateos & De la Peña, Fung. Syst. Evol. 13: 204. 2024. Fig. 8.

Apothecia *1.2–6 mm diam. × 0.6–2.5 mm high, scattered, single to subgregarious, discoid to slightly turbinate, sessile to shortly stipitate; disc smooth, sometimes with eroded areas, light pink–purple (9B6–9D6); margin darker (~8D8), smooth; receptacle concolourous to disc or lighter (~8A6), smooth. Asci †124–149 × 7.5–9.2 µm cylindric–clavate, 8-spored, apex rounded to subconical, pore blue with or without KOH-pretreatment, base arising from croziers. Ascospores †(12.7–)13.7–15.9(–17.9) × (3.4–)4.0–4.6(–4.7) µm hyaline, ellipsoid to cylindrical, rarely subfusiform, 0–1-septate with †0–4 large LBs, overmature spores forming conidia at the poles in chains. Conidia †2.9–4.8 × 2.1–3.6 µm, globose–subglobose to ovoid, hyaline, usually containing one LB. Paraphyses cylindric uninflated to slightly enlarged at apex, straight to wavy, septate, terminal cell †(1.8–)2.0–2.7(–3.2) µm diam., hyaline. Medullary excipulum of t. intricata †98–177 µm thick, mildly to moderately gelatinized, with small (1.3–5.5 µm diam.) calcium oxalate crystals not aggregating to druses, hyphae hyaline to light pinkish–brown, †1.8–2.5 µm diam., releasing purplish pigment in KOH. Ectal excipulum made of brown t. angularis 87–125 µm thick, cells †6.1–13.7 µm diam., releasing some pigment in KOH but remaining light brown. Asexual morph see Crous et al. (2024). Culture characteristics unknown.

Materials examined: USA, Hawaiʻi, Hawaiʻi Island, Puʻu Makaʻala Natural Area Reserve, on Cibotium glaucum, May 1996, D.E. Hemmes 1099 (ARIZ-M-AN00673).

Distribution: Known from Hawaiʻi, Germany, and Portugal.

Notes: Ascocoryne laurisilvae was recently described from a single collection of the asexual morph from Portugal (Crous et al. 2024). Here, we provide a preliminary description of the sexual morph based on ARIZ-M-AN00673, collected in Hawaiʻi, USA. The isolate of this collection forms a supported, monophyletic clade with A. laurisilvae in our 2-minimum analyses (Figs S2 and S3), and with A. laurisilvae and the isolate ‘MF206’ from Germany in our all-taxa analyses (Figs 1, 2, and S4).

Ascocoryne laurisilvae belongs to clade 2 with A. wilsoniae. Both species have ascospores with conidia in chains, lack apically inflated paraphyses, have inamyloid ectal excipula, and contain small calcium oxalate crystals (which may be absent or inconspicuous in A. wilsoniae). More data are needed for a complete comparison between A. wilsoniae and A. laurisilvae, but at this time, A. wilsoniae has ascospores more fusiform in shape that are up to 3-septate, while A. laurisilvae generally has ellipsoid–cylindrical ascospores up to 1-septate.

Collection ARIZ-M-AN00673 consisted of one large and two medium-sized apothecia. Due to the small size of the collection, only tissue from the edge of the large apothecium was destroyed for DNA extraction and morphological analysis. Upon examination in 2022 before A. laurisilvae was published, it was considered a singleton in our dataset and was not destructively sampled further to pursue a formal description before it was returned to ARIZ. Colour codes in the description are based on a digitized slide.

Ascocoryne neoinflata Stallman, sp. nov. MB 861570. Fig. 9.

Etymology: The specific epithet refers to the apically enlarged paraphyses and indicates a different taxon than the informally used epithet ‘inflata.’

Diagnosis: Differs from other Ascocoryne species by its reddish-purple and discoid–pulvinate apothecia and capitate paraphyses.

Apothecia *0.8–3 mm diam. × 0.5–1.5 mm high, scattered to gregarious, solitary or cespitose, convex-discoid when young, later discoid-flat to pulvinate with broad attachment to substrate, sessile to subsessile, disc smooth developing pits or folds in age, light to dark purplish–red (8B8, 10B7, 11A7–11C7); margin undifferentiated; receptacle glabrous, concolourous to slightly lighter. Asci †(103–)107–123(–135) × (5.9–)7.0–8.9(–10.0) µm, cylindric–clavate, 8-spored, spores obliquely biseriate to uniseriate in lower part, apex rounded to subconical, pore 1.5–2.0 × (1.2–)1.3–2.0 µm, blue with or without KOH-pretreatment, base arising from croziers. Ascospores †(12.1–)13.9–17.3(–20.2) × (4.2–)4.7–5.4(–6.1) µm, hyaline, ellipsoid–fusoid, with †two medium to large (1.3–3.0 µm diam.) LBs, overmature spores 1(–3)-septate and sometimes forming germ tubes or conidia in chains from ascospore wall. Conidia †4.8–5.3 × 2.3–2.6 µm, ovoid or fusiform, hyaline. Paraphyses strongly to very strongly clavate–capitate, rarely submoniliform, septate, terminal cell †(7.1–)12.0–25.3(–30.6) × (3.0–)3.7–5.2(–6.0) µm, lower cells †(12.7–)13.2–23.6(–32.7) × (1.0–)1.3–1.9 µm, hyaline, rarely branched, slightly exceeding the asci. Medullary excipulum of t. intricata †69–596 µm thick, hyaline, cells †1.4–2.9 µm diam., loosely packed, embedded in a colourless gel, calcium oxalate crystals absent. Ectal excipulum †25–87 µm thick, differentiated in three layers, innermost layer pigmented, inner layer hyaline, cortical layer glassy-like with slightly protruding thick-walled cells covered by a layer of gel. All layers are made of t. globulosa–angularis with cells †(4.9–)5.7–12.9(–27.3) × (3.0–)4.4–10.3(–26.3) µm, larger at base and diminishing in size towards margin, pigments dissolving in KOH; base of ectal excipulum covered by another layer of t. intricata composed of loosely packed hyphae, strongly gelatinized, that connects the ascomata to the substrate. Asexual morph unknown. Culture characteristics unknown.

Distribution: Thus far only known from Cusuco National Park in northwestern Honduras.

Typus: Honduras, Department of Cortés, Cusuco National Park, 15.50640, −88.24284, ~2048 m a.s.l., on corticated wood, 15 Jul. 2022, J.K. Stallman HONDURAS22-F379 (holotype PULF29763). GenBank: ITS = PQ483772, 28S = PQ483860, 18S = PQ483872, EF1 = PQ505765.

Additional materials examined: Honduras, Department of Cortés, Cusuco National Park, 15.50879, −88.23322, on decorticated wood, 11 Jul. 2022, J.K. Stallman HONDURAS22-F328 (PULF29762); ibid., 15.50640, −88.24284, on corticated wood, 15 Jul. 2022, J.K. Stallman HONDURAS22-F379 (PULF29764).

Notes: Ascocoryne neoinflata is part of the sarcoides group (clade 1, Fig. 2) with conidia budding in chains and presumably large LBs in living specimens, although we did not find living spores in our collections. It differs from other described species in clade 1 by its capitate paraphyses, but occurs in a subgroup with several other undescribed species and C. ‘inflata’ (nom. inval.) which share this feature. The name Coryne ‘inflata’ was used by Wilson (1962) for a species with capitate paraphyses from northeastern North America. Coryne ‘inflata’ sensu Wilson has larger apothecia (up to 10 mm diam. vs 3 mm), longer ascospores [(†14–)18–24(–28) vs (12–)14–17(–20) µm], and larger asci [†125–160 × 8.5–12 vs (103–)107–123(–135) × (5.9–)7–8.9(–10.) µm] than A. neoinflata. A collection identified as A. ‘inflata’ from France has similar ascospore sizes (†18–23 × 4.5–5.5 µm) to C. ‘inflata’ sensu Wilson, although the asci are larger (†150–180 × 12–15 µm) (Moyne 2012). A collection identified as A. ‘inflata’ from Mexico has larger apothecia (4–20 mm) than A. neoinflata and overlapping ascospore sizes (†10–20 × 4–6 µm), although a complete description is absent and the asci measurements (†2.3–170 × 1.7–2.2 µm) appear to be erroneous (Valenzuela et al. 2021). During field collecting over three years, we found C. ‘inflata’ sensu Wilson only once (PULF29998, from Indiana, USA on Quercus). Others are studying this taxon with access to additional collections and vital taxonomic data with plans for a formal description (H.O. Baral pers. comm.); therefore, we do not formally describe A. ‘inflata’ at this time.

In addition to C. ‘inflata’ sensu Wilson and A. neoinflata, we examined three other Ascocoryne taxa with inflated paraphyses, all known from single collections. Ascocoryne neoinflata has longer ascospores [(†12.1–)13.9–17.3(–20.2) µm] than FH374803 [(9.2–)11.0–14.1(–17.7) µm] from Guadeloupe, and shorter ascospores than PULF29796 [(†16.1–)18.1–22.4(–25.1) µm] from West Virgina, USA. Ascocoryne neoinflata has overlapping ascospore and ascus sizes with a collection from Hawaiʻi, USA (CUP063507). The paraphyses of A. neoinflata are generally wider [(†3.0–)3.7–5.2(–6.0) µm] than those of CUP063507 [(†1.3–)2.6–4.1(–4.5) µm], but CUP063507 may represent a less-mature collection as the incidence of capitate paraphyses is low and conidia forming from ascospores or released into the mounting liquid could not be found. Overall, the different phylogenetic positions and ranges support distinct species. An (over)mature collection of the taxon from Hawaiʻi is needed for a comprehensive morphological comparison to include conidia and the colour and shape of the fresh ascomata. Finally, Coryne tasmanica can have small (1–5 mm), discoid–flat apothecia, but this species is likely darker in colour and has substantially longer asci (170 µm vs. up to 135 µm) than A. neoinflata (Dennis 1958).

Ascocoryne vinosa (Berk. & M.A. Curtis) Stallman, comb. nov. MB 861573. Fig. 10.

Basionym: Coryne vinosa Berk. & M.A. Curtis, Bot. J. Linn. Soc. 10: 341. 1868.

Synonyms: Ascocoryne trichophora (A.L. Sm.) Seifert, IMA Fungus 5: 95. 2014.

Coryne trichophora (A.L. Sm.) Seifert, Stud. Mycol. 31: 159. 1989.

Heydenia trichophora A.L. Sm., Bot. J. Linn. Soc. 35: 13. 1901.

Conidiomata (desiccated) †0.7 mm diam. × 1.7–4 mm high, purplish-black, capitulum lightening to pinkish-purple on rehydration; stipe 0.6 mm diam. at base, 0.3 mm diam. at apex, with a cortical layer of t. angularis–globulosa †24–69 µm thick; cells (4.9–)6.7–12.7(–16.6) × (3.5–)4.1–7.6(–9.4) µm, brownish-grey, releasing pigment in KOH, inner layers made of gelatinized t. intricata 215–327 µm thick; cells 1.9–3.7 µm diam., brownish closer to the outer region, hyaline closer to center, retaining pigments in KOH; both regions with amorphous, dark black elements; capitulum †0.7 mm diam. × 0.8 mm high, with long, cylindrical, sterile hyphae 113–155 × 1.5–2 µm exceeding the conidiophores by 70–150 µm. Conidiophores located only in capitulum, hyaline, branching 2–4 times. Metulae (†9.2–)9.9–14.2(–17.2) × (1.2–)1.4–1.9(–2.4) µm, cylindrical. Phialides (†10.4–)13.5–22.4(–27.5) × 0.9–1.5(–1.8) µm, cylindrical, curved, usually tapering at apex, sometimes at base. Conidia (†1.2–)1.6–2.0(–2.2) × 0.9–1.6 µm, ellipsoid, ovoid, or subglobose.

Distribution: Known from Cuba, Dominica, and Ecuador. Likely widespread throughout tropical America based on observations from online citizen science platforms.

Notes: During our review of old names published in the genus Coryne, we observed that C. vinosa had similar features to A. trichophora. Coryne vinosa was described as a dark-reddish, wrinkled, spindle-shaped structure, “2-lines high” (~4.2 mm) with an enlarged apex and minute conidia, growing singly or in clusters on rotten wood in Cuba (Berkeley 1868). Macroscopically, this is undifferentiable from the original description of A. trichophora (as Heydenia trichophora) from Dominica (Smith 1901) and a modern description of the lectotype K-M121466 (Seifert 1989).

The type of C. vinosa, K-M1434236, consists of two pieces of wood glued to the collection packet. The wood contained nine conidiomata, although some appeared to lack a defined apex and may have been immature, damaged, or destructively sampled. In several cases the conidiomata were twisted and lying nearly parallel to the substrate, obscuring how mature or intact they were. One conidiomata was destructively sampled for microscopy and another was destructively sampled for DNA extraction. Attempts to generate DNA sequence data failed via both Sanger and Illumina sequencing.

Examination of the C. vinosa holotype revealed features consistent with the lectotype of A. trichophora (K-M121466) reported by Seifert (1989): conidia 1.6–2.0 × 1.0–1.4 µm (vs 1.5–2 × 1–1.5) borne on simple, branching conidiophores present at the apex of the conidiomata with sterile hyphae extending beyond the hymenium and a stipe composed of t. intricata in the center and t. angularis at the margins. Although we were unable to obtain a DNA sequence of the holotype for comparison, the morphological evidence shows that C. trichophora is a later synonym of C. vinosa. The original description of A. trichophora lists two specimens collected by W.R. Elliott, 1447 (K-M121466) and 1457 K-M121465). Seifert (1989) stated only Elliott’s collection 1447 was at Kew at the time of his study, but both collections were located by Kew for our study.

Our phylogenetic reconstruction recovers the lectotype and paralectotype of A. trichophora in a clade with modern collections identified as A. trichophora from Ecuador (Vandegrift et al. 2023) in the unsupported clade 3 (Fig. 2). The broader group of clades 3 and 4 is supported in our Bayesian phylogenetic analysis (Fig. 2) and contains all Ascocoryne species with ascospores lacking large LBs and conidia not occurring in chains. Therefore, these features are hypothesized to occur in the sexual morph of A. vinosa. A preliminary description of the apothecia of A. vinosa is only known from Seifert (1989), but LBs and conidia arrangement are not noted.

Ascocoryne wilsoniae Stallman & Quijada, sp. nov. MB 861569. Figs 4, 11.

Fig. 11 — Morphology of *Ascocoryne wilsoniae* (from PULF29780 unless otherwise stated). A–C. *Ascomata in-situ (B. PULF29782; C. PULF29771). D. †Apothecium rehydrated (PULF29776). **E–G.** †Sections in KOH (F. PULF29775; G. PULF29771). H. †Sections in KOH+CR. I. †Sections in H₂O. **J, K.** †Ectal excipulum margin in KOH+CR. **L, M.** †Ectal excipulum upper flanks in KOH+CR. N. †Ectal excipulum lower flanks in H₂O. O. †Ectal excipulum lower flanks in KOH+CR. P. †Ectal excipulum base in KOH+CR. Q. †Ectal excipulum base in KOH. R. †Ectal excipulum lower flanks interior region in KOH+CR. **S, T.** †Ectal excipulum lower flanks interior region with crystals in KOH (PULF29775). U. †Medullary excipulum in KOH+IKI. V. †Medullary excipulum in KOH+MLZ. W. †Medullary excipulum in KOH+CR. X. †Asci in H₂O (PULF29782). Y. †Asci in KOH. Z. †Asci in KOH+CR. **AA.** †Asci in KOH+MLZ. **AB.** †Asci in KOH+CR. **AC.** *Paraphyses in H₂O (PULF30022). **AD.** *Paraphysis in H₂O (PULF30006). **AE.** †Paraphyses in KOH (PULF29771). **AF.** †Paraphyses in KOH+CR. **AG.** *Ascospores (†small spores) in H₂O (PULF29771). **AH.** *Ascospore (†bottom spore) in H₂O. **AI.** *Ascospores in H₂O (PULF29771). **AJ.** †Ascospores and conidia in H₂O (PULF29771). **AK.** †Ascospore germinating in H₂O (PULF29771). **AL.** †Ascospore in KOH (PULF29782). **AM.** †Ascospore in KOH. **AN.** †Ascospore in KOH+MLZ. **AO.** †Ascospores in MLZ, Scale bars: A–D = 5 mm; E–I = 100 µm; J–Q = 50 µm; R–AO = 10 µm

Etymology: The specific epithet honours Doreen E. Wilson (née Wells), who studied the genus Coryne (=Ascocoryne) in North America in the mid-20^th century.

Diagnosis: Differs from A. sarcoides by its predominantly fusiform ascospores, non to weakly gelatinized medullary excipulum, and ectal excipulum with an inner region of thick t. intricata.

Apothecia *4.5–10(–30) mm diam. × 2–6 mm high, scattered and solitary to subgregarious, cupulate when immature, later discoid, depressed at center or slightly turbinate, sessile to subsessile with broad attachment, sometimes poorly developed stipe immersed in substrate; disc smooth or sometimes becoming furrowed in age, appearing subcerebriform, purple to dark purple when young (15E6, 15F5, 17F4), potentially lightening to pinkish-purple when mature (~14C5); margin marked on immature ascocarps, smooth or slightly lobate, concolourous to slightly lighter; receptacle concolourous, glabrous. Asci *(163–)177–208(–218) × (8.1–)9.9–13.4(–14.4) µm, †(133–)141–165(–184) × (6.2–)7.7–10.4(–12.8) µm, cylindric-clavate, 8-spored, obliquely biseriate, apex rounded to subconical, slightly †thick-walled, pore 1.2–2.1 × 1.0–1.6, blue with or without KOH-pretreatment, base arising from croziers. Ascospores *(15.7–)17.4–20.6(–25.0) × (4.4–)5.7–7.4(–9.2) µm, †(12.3–)14.3–17.5(–19.1) × (3.5–)4.2–5.4(–6.1) µm, hyaline, fusoid to slightly fusiform-clavate, with *2 large (3–6 µm diam.) LBs and numerous smaller (< 1 µm diam.) LBs filling the remaining cytoplasm (LCI 5), 1–3(–4)-septate, overmature spores forming germ tubes or conidia chains from one or both poles. Conidia †1.9–3.5 × 1.6–3 µm, globose to broadly ellipsoid, hyaline, with one small LBs in the center. Paraphyses uninflated cylindrical to slightly clavate, straight or slightly sinuous, septate, terminal cell †(12.3–)14.7–36.4(–40.3) × 2.0–2.8 µm, lower cells †(9.6–)12.7–26.9(–38.7) × (1.3–)1.5–2.0(–2.4) µm, hyaline, branching not observed, with grey or colourless VBs in the apical or secondary cells, from long and cylindrical (up to 32 µm long) to small and globose (< 1 × 1 µm). Medullary excipulum of t. intricata †154–750 µm thick, hyaline to light brown, not particularly gelatinized, without IKI reaction, cells †1.8–3.5(–5.0) µm diam., loosely packed, embedded in a colourless gel, thickening and darkening in transition to ectal excipulum, containing (PULF29771, PULF29775, PULF29776) to not containing (all other collections) a few to many small to medium (4.0–13.1 µm diam.) calcium oxalate crystals, not to rarely aggregating into clusters up to 23 µm diam. Ectal excipulum †55–172 µm thick, generally composed of two regions. Inner region of brownish-purple t. intricata †38–98 µm thick, thinning to disappearing towards margin, cells †(2.4–)3.4–6.4 µm diam., well-packaged without spaces among cells, pigments somewhat dissolving in KOH but remaining light brownish-yellow; outer region lighter in colour, made of t. globulosa–angularis †26–94 µm thick, cells †3.5–9.5(–16.0) µm diam., with inflated ellipsoid, protruding cells clavate to subcapitate, with sparse yellowish amorphous resin, pigments mostly dissolving in KOH, sometimes with crystals as the medullary excipulum; base of ectal excipulum covered by another layer of t. intricata composed of loosely packed hyphae, strongly gelatinized, that connects the ascomata to the substrate, this layer is blue in IKI with or without KOH pretreatment. Asexual morph unknown.

Culture characteristics: On PDA growing slowly, 5–15 mm diam. after 30 d, white to off white, colony sparse, powdery in appearance; on Rose Bengal growing slowly, 8–20 mm diam. after 50 d, deep purple (~17f3) with sparse covering of white hyphae, in places aggregating into erect strands up to 2 mm high from a shared base; hyphae *(1.5–)2.4–5.4(–7) µm diam., cylindrical, hyaline; on both PDA and RB producing hyaline ellipsoid–ovoid conidia *(2.3–)2.8–3.9(–4.5) × (1.4–)1.7–2.3(–2.4) µm, containing 0–2 small (≤ 1 µm) LBs, borne on fusiform, subconical, or less-commonly lageniform phialides, *(5.1–)6.9–12.6(–14.9) µm long, on inflated cylindrical metulae *(4.0–)6.0–10.1 µm long; on PDA also producing globose conidia *2.5–4.3 µm diam. directly from hyphae walls in chains, sometimes branching, first cell in chain sometimes elongated.

Distribution: Known from Indiana and Vermont, USA.

Typus: USA, Indiana, Yellowwood State Forest, 39.1324, −86.3378, ~250 m a.s.l., on decorticated angiosperm wood, 15 Oct. 2022, J.K. Stallman & C. Brito 417 (holotype PULF29780). GenBank: ITS = PQ483800, 28S = PQ483841, EF1 = PQ505774, RPB2 = PQ505750.

Additional materials examined: USA, Vermont, Windsor County, Ascutney, 43.4070, −72.4070, ~125 m a.s.l., on moss-covered gymnosperm wood, Sep. 1996, S. Flynn (CUP063597); Indiana, Brown County, Yellowwood State Forest, 39.1325, −86.3375, on decorticated angiosperm wood, 1 Oct. 2022, J.K. Stallman & C. Brito 406 (PULF29771), culture DAOMC 252829; ibid., 410 (PULF29775); ibid., 411 (PULF29776); ibid., 39.1324, −86.3378, 15 Oct. 2022, 419a (PULF29782); ibid., 39.1323, −86.3380, 7 Oct. 2023, J.K. Stallman 432 (PULF29789); ibid., 433 (PULF29790); ibid., 39.1316, −86.3382, 6 Nov. 2024473 (PULF30006); ibid., 39.1321, −86.3410, on corticated and decorticated Quercus alba, 489 (PULF30022).

Notes: Ascocoryne wilsoniae has ascospores with large LBs and conidia that form in chains. It differs from A. sarcoides by its fusiform ascospores, region of thickened t. intricata in the interior of the ectal excipulum, and lacking a strongly gelatinized medullary excipulum. Conidia produced by our cultures of A. wilsoniae were ellipsoid, ovoid, or subglobose, differing from the predominantly cylindrical or allantoid conidia produced by A. sarcoides (Wilson 1962, as C. constricta and C. striata). Ascocoryne wilsoniae has larger ascospores (12–19 × 3.5–6 vs 9–15 × 3.5–4 µm) and different apothecium colours (purple vs brown) than A. albida (Dennis 1971, as A. solitaria)

The inner ectal excipulum of A. wilsoniae maintains a brownish colouration in KOH compared with other species examined here where pigments fully dissolve. Three of the ten collections examined (PULF29771, PULF29775, PULF29776) had crystals present in the medullary and ectal excipulum but we did not observe crystals in others, or there were so few (< 5 total) it was not possible to determine if these were inherent to the ascomata, or exogeneous in origin. Ascocoryne striata has crystals and an ectal excipulum with differentiated cortical cells, but these cells differ by branching and A. striata has ascospores with small LBs (Kučera & Lizon 2005).

DISCUSSION

Based on 234 new sequences generated from 144 collections from recent fieldwork and fungaria studies, we clarify the genera belonging to the family Gelatinodiscaceae and propose four new species and one new combination in Ascocoryne.

Metagenomic sequencing of fungaria specimens recovered ITS1, ITS2, or 28S sequences from the target organism 32 out of 115 times (27.8 %) for the 37 collections attempted. Results were variable, with several successes in collections made during the 1800s (e.g., H6036559 from 1860, K-M121465 and K-M121466 from (≤ 1867) while some collections made as recently as 1989 (NY3416360) did not yield a sequence from the target organism. Our 27.8 % success rate compares to 64 % from 36 specimens collected in the late 1800s identified as Peziza (Forin et al. 2018), 25 % for 20 newer and older specimens across Ascomycota (Miller et al. 2022), and 50 % for four Chlorosplenium specimens (Stallman et al. 2024).

Genera in Gelatinodiscaceae

Gelatinodiscaceae is recovered as a monophyletic, supported family including the 9 genera Ascocoryne, Ascotremella, Chloroscypha, Dimorphospora, Helicodendron, Neobulgaria, Ombrophila, Skyathea, and Xerombrophila (Figs 1, S1, S2). Phaeangellina is excluded from the family and is recovered in a supported clade with Piceomphale bulgarioides and ‘Cenangium’ acuum (Fig. S1) that has previously been recovered in the polyphyletic Rutstroemiaceae and Sclerotineaceae families (e.g., Pärtel et al. 2017). We were unable to sequence both the type of Cenangium empetri nom. illegit. (K32261), later named Phaeangella empetri and then combined in a new genus as Phaeangellina empetri (Dennis 1955), and other collections close to the type locality in the UK (IMI269374 and IMI262167). We did obtain DNA sequence data from two collections from Denmark (C125384 and C157023) and another from Canada (WSP56939) identified as P. empetri that matched its morphological features and were found on its host, Empetrum nigrum (Ericaceae). ITS sequences between the collection from Canada and one collection from Denmark shared 99.8 % pairwise identity.

The 28S isolate of Clathrosporium intricatum is recovered in a supported clade with Filosporella fistucella, Varicosporium delicatum, and Hydrocina chaetocladia outside Gelatinodiscaceae (Fig. S1). ITS-only phylogenies group these species in Gelatinodiscaceae p.p. (Johnston et al. 2019, Johnston & Park 2025), but multilocus sequence data from these same studies, from Ekanayaka et al. (2019), and our analysis confidently show that none of these species belong in Gelatinodiscaceae. Ekanayaka et al. (2019) placed all these taxa, including Xerombrophila, in Hydrocinaceae, but our results, along with Johnston et al. (2019) and Johnston & Park (2025) show Xerombrophila belongs in Gelatinodiscaceae.

Our analysis recovered four potentially unnamed genera based on combinations of phylogenetic position (Figs 1, S2), life histories, and/or morphological characteristics. The first is represented by ZML09-3, from a wetland in China (Wu et al. 2013), and 203 OA-2013, from peatland bog sediment in Russia (Grum-Grzhimaylo et al. 2016). These isolates are sister to an isolate identified as Ombrophila and are recovered in a broader clade with Dimorphospora and Helicodendron, genera with aquatic life histories. Neobulgaria species have also been found in bog sediment (Grum-Grzhimaylo et al. 2016) and on submerged detritus in peatlands (Raudabaugh et al. 2021). We note that sequences identified as Ombrophila in GenBank generally lack multilocus data (including those identified as the type, O. janthina) and often do not share high pairwise identity. Therefore, we are uncertain if our single isolate represents the true Ombrophila, which some have suggested is a synonym of Neobulgaria (Baral 1999). Similarly, sequences identified as Helicodendron are found scattered throughout Helotiales. We selected sequences from the type species in the genus, which if correctly identified, places Helicodendron in Gelatinodiscaceae (see Johnston & Park 2025 for further discussion).

The second potentially new genus is isolate CBS 301.50, identified as Helicodendron triglitziense from a submerged leaf in England, which is recovered as sister to Xerombrophila. Xerombrophila is only known from dead wood, so this isolate indicates either a new life history strategy in Xerombrophila, or an additional aquatic genus. The third clade is represented by a culture originating from a mossy log in New York, USA (AFTOL-ID1259). It is identified as Neobulgaria pura, but is supported as sister to Ascotremella, separate from other Neobulgaria species. Neobulgaria pura is the type species of the genus, but there is not enough information to determine if this sequence from America represents the true N. pura, which was described from Europe. Finally, a collection on dead wood from Guyana (NY3416357) is recovered in an unsupported position near the Dimorphospora/Helicodendron clade. Morphologically, the collection is apothecial and has large, 3-septate, thick-walled, hyaline ascospores (42.1–)44.2–50.3(–52.6) × (10.5–)11.2–13.1(–14.0) µm with a wrinkled appearance. Globose conidia are 1.6–4.0 µm in diam. and abundant, although their interaction with the ascospores, if any, is not apparent. Calcium oxalate crystals are present in the excipular tissues and the ectal excipulum has an amyloid reaction. Despite its unique ascospores, we chose not to describe a new genus in the absence of additional collections, vital taxonomic data, and culture characteristics.

The species A. turficola is recovered in an unsupported position between all other species of Ascocoryne and the genus Chloroscypha (Fig. 1). Ascocoryne turficola is divergent compared with other Ascocoryne species due to its long-stipitate, green ascomata, ascospores that may have a gelatinous sheath (Filippova et al. 2013), and growth on Sphagnum species, although in some cases it may be growing from herbaceous material under moss (Filippova & Bulyonkova 2013). Although its morphology and substrate are unique, its phylogenetic position is unsupported and does not conflict with its current generic placement. Therefore, we do not propose a new genus for this taxon (Vellinga et al. 2015).

We have examined the largest number of isolates in a phylogeny of Gelatinodiscaceae to date, but our sampling is not without bias. Only 33 isolates of the nine genera and four unnamed potentially genus-level clades or single isolates come from outside the genus Ascocoryne, and 12 of these are in the genus Neobulgaria. Therefore, incorporating more DNA sequence data outside of the genera Ascocoryne and Neobulgaria will be key in resolving deeper nodes and better understanding unnamed clades in the future.

Intrageneric clades and morphological characters in Ascocoryne

All but five Ascocoryne isolates are recovered within four clades that are generally supported and identifiable by morphological features (Figs 2, S3). But, depending on the clade and method of analysis (2-minimum tree, all-taxa tree, ML or Bayesian tree), there may be variability in support and morphological information is unknown for many isolates.

Sexual morphs of taxa in clade 1 have ascospores with large LBs, conidia that form in chains from mature ascospores, highly gelatinized medullary excipula, and do not contain calcium oxalate crystals. Paraphyses may be consistently inflated or not. This clade, and the subclade with inflated paraphyses are only supported in Bayesian analyses. Sexual morphs of taxa in clade 2, which is supported in all multilocus analyses, are similar to those in clade 1. Differences include that crystals are present, although this is variable in A. wilsoniae, and that the medullary excipulum is generally less gelatinized. Clade 3 is not supported in any analyses and contains variable taxa between the 2-minimum and all-taxa trees. In the 2-minimum tree, species have amyloid ectal excipula and crystals are variable. Ascocoryne vinosa is recovered in this group in the all-taxa tree. Clade 4 contains the divergent A. fibrosa sp. nov. with an amyloid and gelatinized ectal excipulum and the ‘cylichnium group’ of several species that all have sexual morphs with abundant crystals, inamyloid ectal excipula, and ascospores with small LBs and conidia that form on the ascospore wall, not in chains.

Summarizing, a synapomorphic morphological characteristic of the larger clades 3 and 4 (supported in both 2-minimum trees and the Bayesian all-taxa tree) are the ascospores with small LBs only and conidia that form on the ascospore wall, not in chains. Clades 1 and 2 form ascospores with at least 1 large LB and the conidia form in chains. Apically inflated paraphyses may be a synapomorphic characteristic within clade 1, but this is only supported in the multilocus Bayesian analyses. Other characteristics, including the presence of crystals and an amyloid ectal excipulum are useful in determining clade, subclade, or species, but are not unique to monophyletic groups. We note that care should be taken in determining whether the ectal excipulum is amyloid or only the hyphae connecting the ectal excipulum to the substrate (if present). In many cases, these connecting hyphae are amyloid, but the ectal excipulum is not. For example, a collection of A. albida (K-M1434233, as A. solitaria) has blue connecting hyphae in IKI, which does not appear to have been reported previously, but an inamyloid ectal excipulum.

Species richness of Ascocoryne with notes on individual taxa

Ascocoryne contains 13 described species (including those described here) and our phylogenetic analyses suggest there may be as many as 50 species (Fig. 2). Although we tried to improve sampling from tropical locations, there is still a bias towards temperate America and Europe in our dataset, as is true for most fungi (Haelewaters et al. 2024, Stallman et al. 2024). Of the 143 Ascocoryne isolates, 97 are from temperate Europe or North America. While we have 19 isolates from tropical America, we have no representatives from Africa, only two from tropical Asia, and isolates from tropical localities suggest high species richness at these latitudes. For example, the three isolates we examined from the Hawaiian Islands (CUP063507, CUP063506, ARIZ-M-AN00673) and three from Guadeloupe (FH00374802, FH00374803, FH374851) all represent unique species, indicating potentially high species richness over small geographic areas.

Not enough data are available to draw in-depth conclusions about biogeography or host preference in Ascocoryne species except to say that both may be broad. For example, A. albida is found in Europe, North America, and China on both angiosperm and gymnosperm substrates. Ascocoryne sarcoides is also found in Europe, North America, and Asia, but also in South America and Australasia on a variety of hosts including gymnosperms and angiosperms. Ascocoryne aff. cylichnium 1 grows on trees in Bignoniaceae, Fagaceae, Pinaceae, Platanaceae, and Sapindaceae and occurs in India in addition to the USA. Some species may be localized — for example A. fibrosa in the midwestern USA — but this understanding may be due to a lack of sampling or a skewed specialist effect (Haelewaters et al. 2024) in the few localities where those studying Ascocoryne have lived or traveled for research. While most isolates included in our phylogenetic analysis are known from macroscopic sexual or asexual sporocarps, others were isolated from unique substrates. Isolates from soil (e.g., UDB01334927), orchid roots (Herrera et al. 2010), mines (Held et al. 2020), and caves (Burow et al. 2019) are all found in clade 2 of our analysis (Fig. 2), indicating that life histories beyond lignicolous saprotrophism or endophytism in Ascocoryne are possible.

Ascocoryne javanica

Coryne javanica was described from Indonesia (Penzig & Saccardo 1902) and later transferred to Ascocoryne (Thind & Singh 1972). The type was not available for sampling due to lack of material, but notes written by Dr. Richard P. Korf from microscopic examination of the syntype (CUP-K-4733) state, ‘ectal excipulum blue–green in iodine!’ (T. Iturriaga pers. comm.).

The only Ascocoryne specimen we obtained DNA sequence data for from Indonesia (NY3416361) had an amyloid ectal excipulum, but lacked crystals and had smaller ascospores compared to A. javanica (14.4–16.5 vs 19–21 µm). A collection reported by Thind & Singh (1972) as A. javanica (number 3785) from India was not located at K upon a loan request. The authors stated that this collection resembled the concept of A. javanica in all respects, but the ascospores are reportedly much shorter (12–17 µm) than the original description of A. javanica and the paraphyses are stated to be abruptly enlarged into pyriform heads, while the original description states the paraphyses are filiform (Penzig & Saccardo 1902). Coryne sarcoides var. javanica, described from Indonesia (Hennings 1900), has ascospores closer in size (16–18 µm) to NY3416361, but the asci are (erroneously?) reported as 50–65 µm, much smaller than NY3416361 (X̄ = 110–123 µm). More collections from Indonesia and southeastern Asia will improve our understanding of the species richness of Ascocoryne in this region and may lead to modern collections of A. javanica that can provide additional morphological details and DNA sequence data for this taxon.

Ascocoryne cylichnium

Based on our phylogenetic trees, there are eight different lineages within the A. cylichnium group (Figs 2, S3), four of which are found in Europe, from where the species was described (Tulasne 1853). It is unclear which of these isolates, if any, constitutes the ‘true’ Ascocoryne cylichnium. The holotype of A. cylichnium could not be found at P or PC upon request (B. Buyck, pers. comm.) and we are not aware of modern collections with complete descriptions and associated DNA sequence data from Europe. A small portion of an apothecium of the type of Bulgaria urnalis (H6036559), collected in Finland in 1860 and later synonymized with A. cylichnium, was permitted for DNA sequencing and morphological analysis. Although not enough material was available for a comprehensive description, the ascospores measured †(17.5–)18.8–23.0(–24.7) × (4.2–)4.6–5.4(–5.8) µm, the asci lacked croziers, and the ITS sequence shows > 99.4 % percent similarity with isolates from the USA, Canada, and Korea. Croziers are present in all other species recovered in the cylichnium group in our analyses: A. aff. cylichnium 1 and A. aff. cylichnium 2 from the midwestern USA, PULF29797 from the pacific northwestern USA, and at least one or more taxa from Europe and the Canary Islands (Baral 2000; Quijada et al. 2017). Until the holotype of A. cylichnium is found and yields useful information, or other material is epitypified, it would be premature to describe species in the A. cylichnium group that lack unique features.

Ascocoryne striata

The description of A. striata states that it contains calcium oxalate crystals, conidia in chains, and many small LBs (Kučera & Lizon 2005), meaning it does not morphologically fit into any of the four clades in our phylogenetic analysis (Fig. 2). In no other Ascocoryne species described, or specimen examined in this study, do conidia occur in chains from ascospores without also containing at least one large LB. Photos of the holotype (NY 7596) on Mycoportal.org show a small piece of wood, 1.5 × 10 cm, containing one potentially mature apothecium. Annotations show the collection has been examined at least four times (by J.B. Ellis & B.M. Everhart, F. Petrak, D.E. Wilson, and V. Kučera & P. Lizon), likely depleting apothecia available for additional destructive sampling. Petrak annotated the specimen as conspecific with Coryne sarcoides, while Wilson’s annotation states that she considered the two species distinct, noting that Persoon’s specimen of A. sarcoides at L had larger ascospores (15–26 × 3.8–6.0 μm vs. 13–21 × 3.2–4.6 μm) and more septa [5(–6) vs. 3(–5)], and that differences also occur in the cultural morph. The DNA sequence data show at least two collections identified as A. striata by Wilson in her thesis (Wilson 1962) are actually A. sarcoides (see below), but the only published, full description of the holotype of A. striata (Kučera & Lizon 2005) does not report features consistent with A. sarcoides. Kučera & Lizon (2005) stated that they were unable to examine collections from Wilson (1962) and they only found one other collection of A. striata, W6596, after ‘all major world herbaria were searched.’ The specimen at W, now stored under herbarium number W0383792, was collected in Austria in 1898 and contains four small pieces of wood. The largest piece, approximately 2 × 4cm, appears to host apothecia of approximately 1mm in diam, but both the number of apothecia and whether they are mature is uncertain based on the photos available.

Ascocoryne (Coryne) species reported in Wilson’s thesis and the utility of cultural data

Doreen E. Wilson (née Wells), who passed away in 2021, studied the genus Coryne (Ascocoryne) for her PhD dissertation at the University of Toronto (Wilson 1962), examining ~106 Gelatinodiscaceae collections during at least a six-year period from 1956 to 1962. She reported eight species, including four new species that were never formally published. We were able to loan specimens, view permanent slides, and in some cases destructively sample material for morphological examination and DNA extraction, generating sequence data for at least one isolate identified as each of her eight reported species.

The DNA sequence data for Coryne ‘minima’ (TRTC32202) shows it is a Neobulgaria sp. (Fig. 1), and it will not be further discussed. Collections identified as C. striata (TRTC32215, DAOM57653) and C. constricta (DAOM35421) are recovered in a clade with A. sarcoides (Fig. 2). Wilson proposed TRTC32215 as the neotype of A. sarcoides five years following the publication of her thesis (Groves & Wilson 1967), indicating her species concept, at least in this case, changed after finishing her dissertation. Wilson considered C. ‘constricta’ to be similar to C. striata with minor differences in the shapes of the ascospores and different sizes of conidia and conidiophores in culture. Collections identified as C. ‘angustata’ (TRTC32198, TRTC32257 and TRTC33484), C. urnalis (DAOM40674, DAOM40678), and C. cylichnium (DAOM43039) all are recovered in a clade with the holotype of Bulgaria urnalis (Fig. 2; labelled as A. cf. cylichnium). Wilson’s descriptions of these three species are similar, primarily differing in culture characteristics: C. ‘angustata’ was considered unique for rarely having ovoid conidia, and having narrower, more curved allantoid/rod-shaped conidia. Coryne cylichnium was considered unique for having a slimy/metallic appearance of the culture colony. Measurements of ascospores and asci overlap. A collection of C. ‘inflata’ (TRTC32178) is recovered with other collections from northeastern North America with inflated paraphyses (Fig. 2). Finally, C. sarcoides (DAOM33818) is recovered with collections of A. aff. cylichnium 1 (Fig. 2). Collections identified as C. sarcoides were not available for destructive sampling beyond cultures, but the original description is consistent with a species in the cylichnium group with conidia not occurring in chains from ascospores.

In summary, the eight Coryne species in Wilson (1962) likely represent four Ascocoryne and one Neobulgaria species: A. aff. cylichnium 1, A. cf. cylichnium, A. ‘inflata,’ A. sarcoides, and Neobulgaria sp. (Table S4). These results suggest that features of Ascocoryne cultures are polymorphic and this led Wilson to separate taxa that otherwise had very similar apothecial morphology. In addition to frequently producing two types of conidia, both sizes and shapes of conidia, sizes of conidiophores, and the macroscopic appearance of the colony can vary within the culture of a single species. We note that permission to destructively sample Wilson’s collections was limited, and some collections consisted only of cultures. Therefore, we could not examine several replicates of each of Wilson’s eight putative species. Doing so may have uncovered more species than the five we report here.

Names described in Coryne

The genus Coryne accommodated a wide assortment of taxa described from both asexual and sexual collections since its inception by Nees (1816). Later, Groves & Wilson (1967) introduced the genus Ascocoryne to accommodate Coryne species known from the ‘perfect’ (sexual) morph. In reviewing the 86 Coryne taxa listed on Index Fungorum (2025), we found 31 with a description of a sexual morph that had not been transferred out of Coryne or Ascocoryne (Table S5). Most of these taxa can quickly be excluded as synonyms for the new Ascocoryne species described here based on colour, substrate, ascospore sizes, or other characteristics. The type localities of most of these taxa are from Europe (23) and Brazil (4).

Evaluating the ITS and CTP loci as DNA barcodes of Ascocoryne species

Both ITS and CTP loci were adequate in identifying all six Ascocoryne species tested. The CTP locus performed marginally better, successfully barcoding the target species in 15/18 attempts compared with 14/18 for ITS. Additionally, the CTP locus identified more isolates with the tree-based method (five to three), had more diagnostic characters on average, and had a larger average p-distance of the barcode gap. Higher variability in distances led to fewer identifications at the CTP locus than the ITS locus using the barcode gap methodology, despite the higher average. While both the CTP and ITS loci were successful, we were only able to include data for eight other putative species in our analysis. Because the genus Ascocoryne is species rich, the ability of these single loci to effectively barcode all taxa in the genus when more species are included is unclear. We note that two of the species that failed to be identified in at least two methods were both the best-sampled: Ascocoryne sarcoides (seven sequences) and A. aff. cylichnium (nine sequences). This indicates that in addition to the inclusion of other species, adding more sequences from the same species may decrease identification likelihood if intraspecific phylogenetic diversity is high.

CONCLUSIONS

Gelatinodiscaceae and Ascocoryne are more diverse than previously understood. Clathrosporium and Phaeangellina are excluded from Gelatinodiscaceae, but Skyathea, Xerombrophila, and multiple other unnamed, genus-level clades are included. We describe four new Ascocoryne species and our phylogenies suggest the genus contains more than three times the 13 currently described species. The CTP and ITS loci provide similar discriminatory power as DNA barcodes in the six Ascocoryne species we analysed. Species with ascospores that form conidia in chains and have at least one large LB are found in two distinct clades, while all species with only small LBs and conidia that do not occur in chains form a separate group. Further investigation in Gelatinodiscaceae will benefit from more collections and multilocus sequence data outside the genera Ascocoryne and Neobulgaria. Further investigation in Ascocoryne will benefit from more collections from tropical localities and adding data for species lacking DNA sequence data (e.g., A. striata) or modern descriptions (e.g., A. javanica), only known from a single morph (e.g., A. vinosa), or where it is unclear which species name applies in the modern context (e.g., A. cylichnium).

Acknowledgments

We thank Julia Peterson, Kathryn Maley, Jingyu (Lucy) Liu, and Patrick Verdier for sharing collections of Ascocoryne. We thank Hans-Otto Baral for sharing data on Ascocoryne collections and providing feedback on a draft of the manuscript. We thank Britt Bunyard and Zheng Wang for sharing DNA sequence data of Ascocoryne turficola. We thank Qian Gao for sharing information on specimen FPYF3016. We thank all herbarium staff that helped answer inquiries, loan, or accession Ascocoryne collections: Bart Buyck at PC, Betsy Arnold and Joseph Myers at ARIZ, Christian Lange at C, D. Rabern Simmons at PUL, Kathie Hodge and Teresa Iturriaga at CUP, Laura Briscoe at NY, Lee Davies at K, Michaela Schmull, Genevieve Tocci, and Hannah Merchant at FH, Jennifer Wilkinson at TRTC, Monique Slipher at WSP, Otto Miettinen at H, Simona Margaritescu at TRTC, Tanja M. Schuster at W, and staff of the Guyana Centre for Study of Biological Diversity for BRG. Additional thanks to Dillon Husbands for assistance in accessioning collections at BRG. We thank Tara Rintoul and Emily Ranahan at DAOMC for permitting and accessioning cultures. Funding for this project was provided by the following organizations: the Garden Club of America (Fellowship in Tropical Botany to JKS), the Indiana Academy of Sciences (Winona Welch Award to JKS), the Linnean Society of London (LinnéSys: Systematics Research Fund to JKS), the Mycological Society of America (Clark T. Rogerson Student Research Award to JKS, Forest Fungal Ecology Postdoctoral Research Award to DH), Purdue University (D. Woods Thomas Memorial Fund to JKS), Ramon y Cajal Programme (contract RYC2022-036618-I funded by MCIN/AEI/10.13039/501100011033 and FSE+ to LQ), Society of Systematic Biologists (Graduate Student Research Award to JKS), the U.S. Department of Agriculture Hatch Project 1010662 (MCA), the U.S. National Science Foundation (DEB-2018215 to CAQ, DEB-2018098 to DH, and DEB-1556412 and DEB-2127290 to MCA) and the Czech Academy of Sciences (Lumina Quaeruntur Fellowship, LQ200962501 to DH). Collecting and research permits in Guyana were granted by the Guyana Environmental Protection Agency. Permits in Honduras were granted by the Instituto de Conservación Forestal (ICF) with the help of Phoenix Davies and Eduardo Batista. We thank the Aime lab group and Christian Cruz at Purdue University for feedback on earlier versions of this paper, and Anusha Ekanayaka and an anonymous reviewer for feedback on the submitted manuscript.

Footnotes

DATA AVAILABILITY The sequences generated in this study are deposited with NCBI GenBank (Table S1). The alignments are provided as supplementary files (Files S3–S9) as are additional files, figures, and data.

Declaration on conflict of interest: The authors declare that there is no conflict of interest.

SUPPLEMENTARY MATERIAL

Fig. S1

Leotiomycetes-wide phylogenetic tree to place Clathrosporium intricatum, Hydrocina chaetocladia, Phaeangellina empetri, Skyathea hederae, and Xerombrophila crystallifera. The Leotiomycetes v. 1 reference tree was used in T-BAS v. 2.4, selecting “Place Unknowns” and the “de novo” option for the IQ-TREE placement under partitioned models with a GTRGAMMA as substitution model for each partition. Neurospora crassa and Xylaria hypoxylon were used as outgroups and 1000 SH-aLRT and ultrafast bootstrap (UFBoot2) supports were used. Clades including the taxa of interest are highlighted in pink.

per-2026-56-07-SF1.pdf^{(59.7KB, pdf)}

Fig. S2

Gelatinodiscaceae 2-minimum phylogenetic tree. The maximum-likelihood topology is displayed with bootstrap support values ≥ 70 and Bayesian posterior probability values ≥ .90 when nodes are shared between trees. Within Ascocoryne, clades or individual isolates with morphological data are highlighted and ascospore lipid bodies (LBs) and conidia type, whether paraphyses apices are apically inflated, whether the ectal excipulum is amyloid, and whether calcium oxalate crystals are present is shown. The scale bar indicates the number of expected substitutions per site.

per-2026-56-07-SF2.pdf^{(179.6KB, pdf)}

Fig. S3

Ascocoryne 2-minimum phylogenetic tree. The maximum-likelihood topology is displayed with bootstrap support values ≥ 70 and Bayesian posterior probability values ≥ .90 when nodes are shared between trees. Within Ascocoryne, clades or individual isolates with morphological data are highlighted and ascospore lipid bodies (LBs) and conidia type, whether paraphyses apices are apically inflated, whether the ectal excipulum is amyloid, and whether calcium oxalate crystals are present is shown. The scale bar indicates the number of expected substitutions per site.

per-2026-56-07-SF3.pdf^{(173.6KB, pdf)}

Fig. S4

Ascocoryne ITS-only phylogenetic tree. The maximum-likelihood topology is displayed with bootstrap support values ≥ 70 and Bayesian posterior probability values ≥ .90 when nodes are shared between trees. Within Ascocoryne, clades or individual isolates with morphological data are highlighted and ascospore lipid bodies (LBs) and conidia type, whether paraphyses apices are apically inflated, whether the ectal excipulum is amyloid, and whether calcium oxalate crystals are present is shown. The scale bar indicates the number of expected substitutions per site.

per-2026-56-07-SF4.pdf^{(182.6KB, pdf)}

File S1

Reads from Illumina sequence data after processing with DADA2.

per-2026-56-07-SFF1.xlsx^{(401.4KB, xlsx)}

File S2

Collection information for any specimens discussed in the text but without full metadata provided in the Taxonomy section.

per-2026-56-07-SFF2.docx^{(32KB, docx)}

File S3

Gelatinodiscaceae all-taxa phylogenetic tree alignment.

per-2026-56-07-SFF3.txt^{(1,001.3KB, txt)}

File S4

Gelatinodiscaceae 2-minimum phylogenetic tree alignment.

per-2026-56-07-SFF4.txt^{(619.7KB, txt)}

File S5

Ascocoryne all-taxa phylogenetic tree alignment.

per-2026-56-07-SFF5.txt^{(782.3KB, txt)}

File S6

Ascocoryne 2-minimum phylogenetic tree alignment.

per-2026-56-07-SFF6.txt^{(447.4KB, txt)}

File S7

Ascocoryne ITS-only phylogenetic tree alignment.

per-2026-56-07-SFF7.txt^{(75KB, txt)}

File S8

CTP alignment for barcode analysis.

per-2026-56-07-SFF8.txt^{(28.7KB, txt)}

File S9

ITS alignment for barcode analysis.

per-2026-56-07-SFF9.txt^{(21.4KB, txt)}

File S10

Gelatinodiscaceae 2-minimum Bayesian phylogenetic tree.

per-2026-56-07-SFF10.txt^{(13.7KB, txt)}

File S11

Gelatinodiscaceae all-taxa Bayesian phylogenetic tree.

per-2026-56-07-SFF11.txt^{(22.6KB, txt)}

File S12

Gelatinodiscaceae 2-minimum maximum likelihood phylogenetic tree.

per-2026-56-07-SFF12.txt^{(22.6KB, txt)}

File S13

Gelatinodiscaceae all-taxa maximum likelihood phylogenetic tree.

per-2026-56-07-SFF13.txt^{(12.4KB, txt)}

File S14

Ascocoryne 2-minimum Bayesian phylogenetic tree.

per-2026-56-07-SFF14.txt^{(9.8KB, txt)}

File S15

Ascocoryne all-taxa Bayesian phylogenetic tree.

per-2026-56-07-SFF15.txt^{(167.2KB, txt)}

File S16

Ascocoryne 2-minimum maximum likelihood phylogenetic tree.

per-2026-56-07-SFF16.txt^{(5.4KB, txt)}

File S17

Ascocoryne all-taxa maximum likelihood phylogenetic tree.

per-2026-56-07-SFF17.txt^{(9.6KB, txt)}

File S18

Ascocoryne ITS-only Bayesian phylogenetic tree.

per-2026-56-07-SFF18.txt^{(158.3KB, txt)}

File S19

Ascocoryne ITS-only maximum likelihood phylogenetic tree.

per-2026-56-07-SFF19.txt^{(9.2KB, txt)}

File S20

CTP barcode analysis distance matrix.

per-2026-56-07-SFF20.xlsx^{(26.1KB, xlsx)}

File S21

ITS barcode analysis distance matrix.

per-2026-56-07-SFF21.xlsx^{(23.6KB, xlsx)}

File S22

CTP barcode phylogenetic tree.

per-2026-56-07-SFF22.txt^{(2.6KB, txt)}

File S23

ITS barcode phylogenetic tree.

per-2026-56-07-SFF23.txt^{(2.4KB, txt)}

Table S1

Herbarium and GenBank accession numbers for collections with associated metadata.

per-2026-56-07-ST1.xlsx^{(40.7KB, xlsx)}

Table S2

Information on alignments and models selected for phylogenetic analyses.

per-2026-56-07-ST2.xlsx^{(18.1KB, xlsx)}

Table S3

Summary of Ascocoryne species supported in phylogenetic analyses.

per-2026-56-07-ST3.xlsx^{(17.9KB, xlsx)}

Table S4

Identifications of Gelatinodiscaceae collections from Wilson (1962) and this study.

per-2026-56-07-ST4.xlsx^{(16.3KB, xlsx)}

Table S5

Notes on Coryne taxa with teleomorphs that have not been transferred outside of Coryne or Ascocoryne.

per-2026-56-07-ST5.xlsx^{(20.9KB, xlsx)}

REFERENCES

Abarenkov K, Zirk A, Piirmann T, et al. (2024). UNITE general FASTA release for Fungi version 04.04.2024. UNITE Community. 10.15156/BIO/2959332 [DOI]
Albu S, Toome M, Aime MC. (2015). Violaceomyces palustris gen. et sp. nov. and a new monotypic lineage, Violaceomycetales ord. nov. in Ustilaginomycetes. Mycologia 107: 1193–1204. 10.3852/14-260 [DOI] [PubMed] [Google Scholar]
Altschul SF, Gish W, Miller W, et al. (1990). Basic local alignment search tool. Journal of Molecular Biology 215: 403–410. 10.1016/s0022-2836(05)80360-2 [DOI] [PubMed] [Google Scholar]
Arhipova N, Jansons A, Zaluma A, et al. (2015). Bark stripping of Pinus contorta caused by moose and deer: wounding patterns, discoloration of wood, and associated fungi. Canadian Journal of Forest Research 45: 1434–1438. [Google Scholar]
Baral HO. (1992). Vital versus herbarium taxonomy: morphological differences between living and dead cells of Ascomycetes, and their taxonomic implications. Mycotaxon 44: 333–390. [Google Scholar]
Baral HO. (1999). Ombrophila hemiamyloidea, an aquatic discomycete. Mycologia Bavarica 3: 50–63. [Google Scholar]
Baral HO. (2000). Key to Ascocoryne. www.ascofrance.com/uploads/forum_file/5598.doc. Accessed on 4 May 2025.
Baral HO, Galan R, Platas G, et al. (2013a). Phaeohelotium undulatum comb. nov. and Phaeoh. succineoguttulatum sp. nov., two segregates of the Discinella terrestris aggregate found under Eucalyptus in Spain: taxonomy, molecular biology, ecology and distribution. Mycosystema 32: 386–428. [Google Scholar]
Baral HO, Haelewaters D, Pärtel K. (2015). A new attempt to classify the families of the Helotiales. Second International Workshop on Ascomycete Systematics: 26. 22–24 April 2015, Amsterdam. [Google Scholar]
Baral HO, Marson G, Bogale M, et al. (2013b). Xerombrophila crystallifera, a new genus and species in the Helotiales. Mycological Progress 12: 475–488. 10.1007/s11557-012-0854-6 [DOI] [Google Scholar]
Berkeley MJ. (1868, ‘1869’) On a collection of fungi from Cuba. Part. II., including those belonging to the families Gasteromycetes, Coniomycetes, Hyphomycetes, Physomycetes, and Ascomycetes. Botanical Journal of the Linnean Society 10: 341–392 [Google Scholar]
Berkeley MJ, Curtis MA. (1868, ‘1869’). Fungi Cubenses. (Hymenomycetes). Botanical Journal of the Linnean Society 10: 280–341. [Google Scholar]
Beug MW, Bessette AE, Bessette AR. (2014). Ascomycete Fungi of North America: A Mushroom Reference Guide. University of Texas Press, USA. [Google Scholar]
Bouckaert R, Vaughan TG, Barido-Sottani J, et al. (2019). BEAST 2.5: An advanced software platform for Bayesian evolutionary analysis. PLOS Computational Biology 15: e1006650. 10.1371/journal.pcbi.1006650 [DOI] [PMC free article] [PubMed] [Google Scholar]
Bouckaert RR, Drummond AJ. (2017). bModelTest: Bayesian phylogenetic site model averaging and model comparison. BMC Evolutionary Biology 17: 42. 10.1186/s12862-017-0890-6 [DOI] [PMC free article] [PubMed] [Google Scholar]
Bunyard B, Wang Z, Malloch D, et al. (2008). New North American records for Ascocoryne turficola (Ascomycota: Helotiales). Fungi Magazine 1: 23–31. [Google Scholar]
Burow K, Grawunder A, Harpke M, et al. (2019). Microbiomes in an acidic rock–water cave system. FEMS Microbiology Letters 366: fnz167. 10.1093/femsle/fnz167 [DOI] [PubMed] [Google Scholar]
Callahan BJ, McMurdie PJ, Rosen MJ, et al. (2016). DADA2: High-resolution sample inference from Illumina amplicon data. Nature Methods 13: 581–583. 10.1038/nmeth.3869 [DOI] [PMC free article] [PubMed] [Google Scholar]
Capella-Gutiérrez S, Silla-Martínez JM, Gabaldón T. (2009) trimAl: a tool for automated alignment trimming in large-scale phylogenetic analyses. Bioinformatics 25: 1972–1973. 10.1093/bioinformatics/btp348 [DOI] [PMC free article] [PubMed] [Google Scholar]
Carbone I, White JB, Miadlikowska J, et al. (2019). T-BAS Version 2.1: Tree-Based Alignment Selector Toolkit for evolutionary placement of DNA sequences and viewing alignments and specimen metadata on curated and custom trees. Microbiology Resource Announcements 8: 10.1128/mra.00328-19. 10.1128/mra.00328-19 [DOI] [PMC free article] [PubMed] [Google Scholar]
Carpenter SE. (1976). Taxonomy, morphology and ontogeny of Gelatinodiscus flavidus. Mycotaxon 3: 209–232. 10.5962/p.413977 [DOI] [Google Scholar]
Chernomor O, Minh BQ, von Haeseler A. (2015). Consequences of common topological rearrangements for partition trees in phylogenomic inference. Journal of Computational Biology 22: 1129–1142. 10.1089/cmb.2015.0146 [DOI] [PMC free article] [PubMed] [Google Scholar]
Crous PW, Jurjević Ž, Balashov S, et al. (2024). Fungal Planet description sheets: 1614–1696. Fungal Systematics and Evolution 13: 183–440. 10.3114/fuse.2024.13.11 [DOI] [PMC free article] [PubMed] [Google Scholar]
Dennis RWG. (1954). Some inoperculate discomycetes of tropical America. Kew Bulletin 9: 289–348. 10.2307/4114399 [DOI] [Google Scholar]
Dennis RWG. (1955) Two proposed new genera of Helotiales. Kew Bulletin 10: 359–362. 10.2307/4109223 [DOI] [Google Scholar]
Dennis RWG. (1956) A revision of the British Helotiaceae in the herbarium of the Royal Botanic Gardens, Kew, with notes on related European species. Mycological Papers 62: 1–216. [Google Scholar]
Dennis RWG. (1958) Critical notes on some Australian Helotiales and Ostropales. Kew Bulletin 13: 321–358. 10.2307/4109542 [DOI] [Google Scholar]
Dennis RWG. (1971). New or interesting British microfungi. Kew Bulletin 25: 335–374. 10.2307/4103238 [DOI] [Google Scholar]
Douglas J, Zhang R, Bouckaert R. (2021). Adaptive dating and fast proposals: Revisiting the phylogenetic relaxed clock model. PLOS Computational Biology 17: e1008322. 10.1371/journal.pcbi.1008322 [DOI] [PMC free article] [PubMed] [Google Scholar]
Ekanayaka A, Hyde K, Gentekaki E, et al. (2019). Preliminary classification of Leotiomycetes. Mycosphere 10: 310–489. 10.5943/mycosphere/10/1/7 [DOI] [Google Scholar]
Filippova N, Bulyonkova TM. (2013). Notes on the ecology of Ascocoryne turficola (Ascomycota: Helotiales) in West Siberia. Environmental Dynamics and Global Climate Change 4: 1–6. 10.17816/edgcc421-6 [DOI] [Google Scholar]
Filippova N, Zvyagina E, Bulyonkova T. (2013). Ascocoryne turficola (Boud.) Korf records from West Siberia. Fungi Magazine 6: 26–30. [Google Scholar]
Forin N, Nigris S, Voyron S, et al. (2018). Next generation sequencing of ancient fungal specimens: The case of the Saccardo Mycological Herbarium. Frontiers in Ecology and Evolution 6: 129. 10.3389/fevo.2018.00129 [DOI] [Google Scholar]
Gao Q, Yan D, Wang K, et al. (2017). Metabolites diversity and hydrocarbons analysis on volatile organic compounds produced by Ascocoryne strains. Forest Research 30: 376–383. [Google Scholar]
Gardes M, Bruns TD. (1993). ITS primers with enhanced specificity for basidiomycetes - application to the identification of mycorrhizae and rusts. Molecular Ecology 2: 113–118. 10.1111/j.1365-294X.1993.tb00005.x [DOI] [PubMed] [Google Scholar]
Griffin MA, Spakowicz DJ, Gianoulis TA, et al. (2010). Volatile organic compound production by organisms in the genus Ascocoryne and a re-evaluation of myco-diesel production by NRRL 50072. Microbiology 156: 3814–3829. [DOI] [PMC free article] [PubMed] [Google Scholar]
Groves JW, Wilson DE. (1967). The nomenclatural status of Coryne. Taxon 16: 35–41. 10.2307/1217104 [DOI] [Google Scholar]
Grum-Grzhimaylo OA, Debets AJM, Bilanenko EN. (2016). The diversity of microfungi in peatlands originated from the White Sea. Mycologia 108: 233–254. 10.3852/14-346 [DOI] [PubMed] [Google Scholar]
Guindon S, Dufayard J-F, Lefort V, et al. (2010). New algorithms and methods to estimate maximum-likelihood phylogenies: Assessing the performance of PhyML 3.0. Systematic Biology 59: 307–21. 10.1093/sysbio/syq010. [DOI] [PubMed] [Google Scholar]
Haelewaters D, Park D, Johnston PR. (2021). Multilocus phylogenetic analysis reveals that Cyttariales is a synonym of Helotiales. Mycological Progress 20: 1323–1330. 10.1007/s11557-021-01736-2 [DOI] [Google Scholar]
Haelewaters D, Matthews TJ, Wayman JP, et al. (2024). Biological knowledge shortfalls impede conservation efforts in poorly studied taxa — A case study of Laboulbeniomycetes. Journal of Biogeography 51: 29–39. 10.1111/jbi.14725 [DOI] [Google Scholar]
Hawksworth DL. (2001). The magnitude of fungal diversity: the 1.5 million species estimate revisited. Mycological Research 105: 1422–1432. 10.1017/S0953756201004725 [DOI] [Google Scholar]
Hebert PDN, Stoeckle MY, Zemlak TS, et al. (2004). Identification of birds through DNA barcodes. PLoS Biology 2: e312. 10.1371/journal.pbio.0020312 [DOI] [PMC free article] [PubMed] [Google Scholar]
Held BW, Salomon CE, Blanchette RA. (2020). Diverse subterranean fungi of an underground iron ore mine. PLoS ONE 15: e0234208. 10.1371/journal.pone.0234208 [DOI] [PMC free article] [PubMed] [Google Scholar]
Hennings P. (1900). Fungi. In: Monsunia: Beiträge zur kenntniss der vegetation des süd- und ostasiatischen monsungebietes (Warburg O, ed). W. Engelmann, Germany: 30–31. [Google Scholar]
Herrera P, Suárez JP, Kottke I. (2010). Orchids keep the ascomycetes outside: a highly diverse group of ascomycetes colonizing the velamen of epiphytic orchids from a tropical mountain rainforest in Southern Ecuador. Mycology 1: 262–268. 10.1080/21501203.2010.526645 [DOI] [Google Scholar]
Hoang DT, Chernomor O, von Haeseler A, et al. (2018). UFBoot2: Improving the ultrafast bootstrap approximation. Molecular Biology and Evolution 35: 518–522. 10.1093/molbev/msx281 [DOI] [PMC free article] [PubMed] [Google Scholar]
Hofstetter V, Miadlikowska J, Kauff F, et al. (2007). Phylogenetic comparison of protein-coding versus ribosomal RNA-coding sequence data: A case study of the Lecanoromycetes (Ascomycota). Molecular Phylogenetics and Evolution 44: 412–426. 10.1016/j.ympev.2006.10.016 [DOI] [PubMed] [Google Scholar]
Hyde KD, Noorabadi MT, Thiyagaraja V, et al. (2024). The 2024 Outline of Fungi and fungus-like taxa. Mycosphere 15: 5146–6239. 10.5943/mycosphere/15/1/25 [DOI] [Google Scholar]
Index Fungorum (2025). Search Index Fungorum. https://www.indexfungorum.org/Names/Names.asp
Jaén-Molina R, Marrero-Rodríguez Á, Reyes-Betancort JA, et al. (2015). Molecular taxonomic identification in the absence of a ‘barcoding gap’: a test with the endemic flora of the Canarian oceanic hotspot. Molecular Ecology Resources 15: 42–56. 10.1111/1755-0998.12292 [DOI] [PubMed] [Google Scholar]
Jaklitsch W, Baral HO, Lücking R, et al. (2016). Syllabus of plant families: A. Engler’s syllabus der Pfanzenfamilien part 1/2 (Frey W. ed.). Borntraeger Gebrueder, Germany. [Google Scholar]
Johnston PR, Baschien C. (2020). Tricladiaceae fam. nov. (Helotiales, Leotiomycetes). Fungal Systematics and Evolution 6: 233–242. 10.3114/fuse.2020.06.10 [DOI] [PMC free article] [PubMed] [Google Scholar]
Johnston PR, Park D. (2025). An updated multigene phylogeny of Leotiomycetes. Persoonia 55: 527–566. 10.3114/persoonia.2025.55.13 [DOI] [PMC free article] [PubMed] [Google Scholar]
Johnston PR, Quijada L, Smith CA, et al. (2019). A multigene phylogeny toward a new phylogenetic classification of Leotiomycetes. IMA Fungus 10: 1. 10.1186/s43008-019-0002-x [DOI] [PMC free article] [PubMed] [Google Scholar]
Johnston PR, Seifert KA, Stone JK, et al. (2014). Recommendations on generic names competing for use in Leotiomycetes (Ascomycota). IMA Fungus 5: 91–120. 10.5598/imafungus.2014.05.01.11 [DOI] [PMC free article] [PubMed] [Google Scholar]
Kalyaanamoorthy S, Minh BQ, Wong TKF, et al. (2017). ModelFinder: fast model selection for accurate phylogenetic estimates. Nature Methods 14: 587–589. 10.1038/nmeth.4285 [DOI] [PMC free article] [PubMed] [Google Scholar]
Karakehian J, Quijada L, Pfister D, et al. (2021). Methods for observing, culturing, and studying living ascospores. Asian Journal of Mycology 4: 1–18. [Google Scholar]
Katoh K, Misawa K, Kuma K, et al. (2002). MAFFT: a novel method for rapid multiple sequence alignment based on fast Fourier transform. Nucleic Acids Research 30: 3059–3066. 10.1093/nar/gkf436 [DOI] [PMC free article] [PubMed] [Google Scholar]
Katoh K, Standley DM. (2013). MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Molecular Biology and Evolution 30: 772–780. 10.1093/molbev/mst010 [DOI] [PMC free article] [PubMed] [Google Scholar]
Kosonen T, Huhtinen S, Hansen K. (2021). Taxonomy and systematics of Hyaloscyphaceae and Arachnopezizaceae. Persoonia 46: 26–62. 10.3767/persoonia.2021.46.02 [DOI] [PMC free article] [PubMed] [Google Scholar]
Kučera V, Lizon P. (2005). Ascocoryne striata, comb. nov. Mycotaxon 93: 163–165. [Google Scholar]
Larena I, Salazar O, González V, et al. (1999). Design of a primer for ribosomal DNA internal transcribed spacer with enhanced specificity for ascomycetes. Journal of Biotechnology 75: 187–194. 10.1016/s0168-1656(99)00154-6 [DOI] [PubMed] [Google Scholar]
Liu YJ, Whelen S, Hall BD. (1999). Phylogenetic relationships among ascomycetes: evidence from an RNA polymerse II subunit. Molecular Biology and Evolution 16: 1799–1808. 10.1093/oxfordjournals.molbev.a026092 [DOI] [PubMed] [Google Scholar]
Matheny PB, Liu YJ, Ammirati JF, et al. (2002). Using RPB1 sequences to improve phylogenetic inference among mushrooms (Inocybe, Agaricales). American Journal of Botany 89: 688–698. 10.3732/ajb.89.4.688 [DOI] [PubMed] [Google Scholar]
Melie T. (2023). Genomic investigations in Ascomycota: Leveraging machine learning, phylogenetics, and comparative genomics methods to explore evolution and ecological specialization. Ph.D. dissertation. Department of Ecology and Evolutionary Biology, University of Colorado at Boulder, USA. https://scholar.colorado.edu/concern/graduate_thesis_or_dissertations/8049g6598 [Google Scholar]
Meyer CP, Paulay G. (2005). DNA barcoding: Error rates based on comprehensive sampling. PLoS Biology 3: e422. 10.1371/journal.pbio.0030422 [DOI] [PMC free article] [PubMed] [Google Scholar]
Meyn R, Langer GJ, Gross A, et al. (2019). Fungal colonization patterns in necrotic rootstocks and stem bases of dieback-affected Fraxinus excelsior L. Forest Pathology 49: e12520. 10.1111/efp.12520 [DOI] [Google Scholar]
Miadlikowska J, Lutzoni F. (2000). Phylogenetic revision of the genus Peltigera (lichen‐forming Ascomycota) based on morphological, chemical, and large subunit nuclear ribosomal DNA Data. International Journal of Plant Sciences 161: 925–958. 10.1086/317568 [DOI] [Google Scholar]
Miller AN, Karakehian J, Raudabaugh DB. (2022). Next-generation sequencing of ancient and recent fungarium specimens. Journal of Fungi 8: 932. 10.3390/jof8090932 [DOI] [PMC free article] [PubMed] [Google Scholar]
Molina L, Rajchenberg M, de Errasti A, et al. (2020). Sapwood-inhabiting mycobiota and Nothofagus tree mortality in Patagonia: Diversity patterns according to tree species, plant compartment and health condition. Forest Ecology and Management 462: 117997. 10.1016/j.foreco.2020.117997 [DOI] [Google Scholar]
Moyne G. (2012). Deux ascomycètes méconnus: Ascocoryne inflata et Rutstroemia alni. Bulletin Annuel de la Fédération Mycologique de l’Est 10: 21–24. [Google Scholar]
Nees von Esenbeck CG. (1816). Das System der Plze und Schwämme. Würzburg, Germany. [Google Scholar]
Nguyen L-T, Schmidt HA, von Haeseler A, et al. (2015). IQ-TREE: A fast and effective stochastic algorithm for estimating maximum-likelihood phylogenies. Molecular Biology and Evolution 32: 268–274. 10.1093/molbev/msu300 [DOI] [PMC free article] [PubMed] [Google Scholar]
Olds CG, Berta-Thompson JW, Loucks JJ, et al. (2023), Applying a modified metabarcoding approach for the sequencing of macrofungal specimens from fungarium collections. Applications in Plant Sciences 11: e11508. 10.1002/aps3.11508 [DOI] [PMC free article] [PubMed] [Google Scholar]
Penzig O, Saccardo PA. (1902). Diagnoses fungorum novorum in insula Java collectorum. Series Tertia. Discomycetes. Malpighia: rassegna mensuale di botanica 15: 201–260. [Google Scholar]
Quandt A, Haelewaters D. (2021). Phylogenetic advances in Leotiomycetes, an understudied clade of taxonomically and ecologically diverse Fungi. In: Encyclopedia of mycology Vol. 1 (Zaragoza Ó, Casadevall A. eds). Elsevier, United Kingdom. 10.1016/b978-0-12-819990-9.00052-4 [DOI] [Google Scholar]
Quijada L, Baral HO, Johnston PR, et al. (2022). A review of Hyphodiscaceae. Studies in Mycology 103: 59–85. 10.3114/sim.2022.103.03 [DOI] [PMC free article] [PubMed] [Google Scholar]
Quijada L, Ribes M, Negrín R, et al. (2017). Lignicolous species of Helotiales associated with major vegetation types in the Canary Islands. Willdenowia 47: 271–291. 10.3372/wi.47.47310 [DOI] [Google Scholar]
R Foundation for Statistical Computing (2023). R Core Team. R: A Language and Environment for Statistical Computing. Austria. [Google Scholar]
Raudabaugh DB, Bach EM, Allen JM, et al. (2021). Fungal communities of submerged fine detritus from temperate peatland and stream habitats. Aquatic Microbial Ecology 86: 191–207. 10.3354/ame01969 [DOI] [Google Scholar]
Rehner SA, Buckley E. (2005). A Beauveria phylogeny inferred from nuclear ITS and EF1-α sequences: evidence for cryptic diversification and links to Cordyceps teleomorphs. Mycologia 97: 84–98. 10.3852/mycologia.97.1.84 [DOI] [PubMed] [Google Scholar]
Roll-Hansen F, Roll-Hansen H. (1979a). Microflora of sound-looking wood in Picea abies stems. European Journal of Forest Pathology 9: 308–316. 10.1111/j.1439-0329.1979.tb00693.x [DOI] [Google Scholar]
Roll-Hansen F, Roll-Hansen H. (1979b). Ascocoryne sarcoides and Ascocoryne cylichnium. Descriptions and comparison. Norwegian Journal of Botany 26: 193–206. [Google Scholar]
Saccardo PA, Traverso GB, Trotter A. (1889). Sylloge fungorum omnium hucusque cognitorum Vol. VIII. Patavii, Italy. [Google Scholar]
Seaver J. (1951). The North American cup-fungi (Inoperculates). Lancaster Press, USA. [Google Scholar]
Seifert KA. (1989). Coryne trichophora, comb. nov., and the implications of its conidiomatal anatomy. Studies in Mycology 31: 157–164. [Google Scholar]
Smith AL. (1901). On some Fungi from the West Indies. Botanical Journal of the Linnean Society 35: 1–19. 10.1111/j.1095-8339.1901.tb02317.x [DOI] [Google Scholar]
Sri-indrasutdhi V, Tsui CKM, Chuaseeharonnachai C, et al. (2015). Helicocentralis hyalina gen. et sp. nov., an aero-aquatic helicosporous fungus (Leotiomycetes, Ascomycota) in Thailand. Mycological Progress 14: 81. 10.1007/s11557-015-1103-6 [DOI] [Google Scholar]
Stallman JK, Johnston PR, Lickey EB, et al. (2024). Recent fieldwork and fungarium studies double known diversity of Chlorosplenium and improve understanding of species distributions. Mycologia 116: 993–1018. 10.1080/00275514.2024.2364567 [DOI] [PubMed] [Google Scholar]
Stallman JK, Haelewaters D, Koch Bach RA, et al. (2024). The contribution of tropical long-term studies to mycology. IMA Fungus 15: 35. 10.1186/s43008-024-00166-5 [DOI] [PMC free article] [PubMed] [Google Scholar]
Stielow JB, Lévesque CA, Seifert KA, et al. (2015). One fungus, which genes? Development and assessment of universal primers for potential secondary fungal DNA barcodes. Persoonia 35: 242–263. 10.3767/003158515x689135 [DOI] [PMC free article] [PubMed] [Google Scholar]
Stiller JW, Hall BD. (1997). The origin of red algae: Implications for plastid evolution. Proceedings of the National Academy of Sciences USA 94: 4520–4525. 10.1073/pnas.94.9.4520 [DOI] [PMC free article] [PubMed] [Google Scholar]
Tamura K, Stecher G, Kumar S. (2021). MEGA11: Molecular Evolutionary Genetics Analysis Version 11. Molecular Biology and Evolution 38: 3022–3027. 10.1093/molbev/msab120 [DOI] [PMC free article] [PubMed] [Google Scholar]
Tanney JB, Seifert KA. (2020). Mollisiaceae: An overlooked lineage of diverse endophytes. Studies in Mycology 95: 293–380. 10.1016/j.simyco.2020.02.005 [DOI] [PMC free article] [PubMed] [Google Scholar]
Taylor JW, Jacobson DJ, Kroken S, et al. (2000). Phylogenetic species recognition and species concepts in Fungi. Fungal Genetics and Biology 31: 21–32. 10.1006/fgbi.2000.1228 [DOI] [PubMed] [Google Scholar]
Thind KS, Singh H. (1972). The Helotiales of India—XVI. Journal of the Indian Botanical Society 51: 44–51. [Google Scholar]
Untereiner WA, Yue Q, Chen L, et al. (2019). Phialophora section Catenulatae disassembled: New genera, species, and combinations and a new family encompassing taxa with cleistothecial ascomata and phialidic asexual states. Mycologia 111: 998–1027. 10.1080/00275514.2019.1663106 [DOI] [PubMed] [Google Scholar]
Vaidya G, Lohman DJ, Meier R. (2011). SequenceMatrix: concatenation software for the fast assembly of multi-gene datasets with character set and codon information. Cladistics 27: 171–180. 10.1111/j.1096-0031.2010.00329.x [DOI] [PubMed] [Google Scholar]
Valenzuela R, Raymundo T, Reyes P, et al. (2021). Ascomycetes from the relic forest of Oreomunnea mexicana, Oaxaca, Mexico. Phytotaxa 528: 19–44. 10.11646/phytotaxa.528.1.3 [DOI] [Google Scholar]
Vandegrift R, Newman DS, Dentinger BTM, et al. (2023). Richer than gold: the fungal biodiversity of Reserva Los Cedros, a threatened Andean cloud forest. Botanical Studies 64: 17. 10.1186/s40529-023-00390-z [DOI] [PMC free article] [PubMed] [Google Scholar]
Vasiliauskas R, Stenlid J. (1998). Fungi inhabiting stems of Picea abies in a managed stand in Lithuania. Forest Ecology and Management 109: 119–126. 10.1016/S0378-1127(98)00226-6 [DOI] [Google Scholar]
Vellinga EC, Kuyper TW, Ammirati J, et al. (2015). Six simple guidelines for introducing new genera of fungi. IMA Fungus 6: A65–A68. 10.1007/BF03449356 [DOI] [Google Scholar]
Vilgalys R, Hester M. (1990). Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species. Journal of Bacteriology 172: 4238–4246. 10.1128/jb.172.8.4238-4246.1990 [DOI] [PMC free article] [PubMed] [Google Scholar]
Voglmayr H. (2004). Spirosphaera cupreorufescens sp. nov., a rare aeroaquatic fungus. Studies in Mycology 50: 221–228. [Google Scholar]
Vu D, Groenewald M, de Vries M, et al. (2019). Large-scale generation and analysis of filamentous fungal DNA barcodes boosts coverage for kingdom fungi and reveals thresholds for fungal species and higher taxon delimitation. Studies in Mycology 92: 135–154. 10.1016/j.simyco.2018.05.001 [DOI] [PMC free article] [PubMed] [Google Scholar]
Wang Z, Binder M, Hibbett DS. (2005). Life history and systematics of the aquatic discomycete Mitrula (Helotiales, Ascomycota) based on cultural, morphological, and molecular studies. American Journal of Botany 92: 1565–1574. 10.3732/ajb.92.9.1565 [DOI] [PubMed] [Google Scholar]
Wang J, Ji-li E, Liu J. (2017). Endophytic fungi diversity of Platycladus orientalis in different age. Journal of Southwest Forestry University 37: 113–117. 10.11929/j.issn.2095-1914.2017.03.018 [DOI] [Google Scholar]
White T, Bruns T, Lee S, et al. (1990). Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: PCR Protocols: A Guide to Methods and Applications (Innis MA, Gelfand DH, Sninsky JJ, et al. eds.). Academic Press, USA: 315–322. [Google Scholar]
Wijayawardene NN, Hyde KD, Lumbsch HT, et al. (2018). Outline of Ascomycota: 2017. Fungal Diversity 88: 167–263. 10.1007/s13225-018-0394-8 [DOI] [Google Scholar]
Wilson D. (1962). A study of eight North American species of Coryne. Ph.D. dissertation. School of Graduate Studies, University of Toronto, Canada. [Google Scholar]
Wu B, Tian J, Bai C, et al. (2013). The biogeography of fungal communities in wetland sediments along the Changjiang River and other sites in China. The ISME Journal 7: 1299–1309. 10.1038/ismej.2013.29 [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Fig. S1

per-2026-56-07-SF1.pdf^{(59.7KB, pdf)}

Fig. S2

per-2026-56-07-SF2.pdf^{(179.6KB, pdf)}

Fig. S3

per-2026-56-07-SF3.pdf^{(173.6KB, pdf)}

Fig. S4

per-2026-56-07-SF4.pdf^{(182.6KB, pdf)}

File S1

Reads from Illumina sequence data after processing with DADA2.

per-2026-56-07-SFF1.xlsx^{(401.4KB, xlsx)}

File S2

Collection information for any specimens discussed in the text but without full metadata provided in the Taxonomy section.

per-2026-56-07-SFF2.docx^{(32KB, docx)}

File S3

Gelatinodiscaceae all-taxa phylogenetic tree alignment.

per-2026-56-07-SFF3.txt^{(1,001.3KB, txt)}

File S4

Gelatinodiscaceae 2-minimum phylogenetic tree alignment.

per-2026-56-07-SFF4.txt^{(619.7KB, txt)}

File S5

Ascocoryne all-taxa phylogenetic tree alignment.

per-2026-56-07-SFF5.txt^{(782.3KB, txt)}

File S6

Ascocoryne 2-minimum phylogenetic tree alignment.

per-2026-56-07-SFF6.txt^{(447.4KB, txt)}

File S7

Ascocoryne ITS-only phylogenetic tree alignment.

per-2026-56-07-SFF7.txt^{(75KB, txt)}

File S8

CTP alignment for barcode analysis.

per-2026-56-07-SFF8.txt^{(28.7KB, txt)}

File S9

ITS alignment for barcode analysis.

per-2026-56-07-SFF9.txt^{(21.4KB, txt)}

File S10

Gelatinodiscaceae 2-minimum Bayesian phylogenetic tree.

per-2026-56-07-SFF10.txt^{(13.7KB, txt)}

File S11

Gelatinodiscaceae all-taxa Bayesian phylogenetic tree.

per-2026-56-07-SFF11.txt^{(22.6KB, txt)}

File S12

Gelatinodiscaceae 2-minimum maximum likelihood phylogenetic tree.

per-2026-56-07-SFF12.txt^{(22.6KB, txt)}

File S13

Gelatinodiscaceae all-taxa maximum likelihood phylogenetic tree.

per-2026-56-07-SFF13.txt^{(12.4KB, txt)}

File S14

Ascocoryne 2-minimum Bayesian phylogenetic tree.

per-2026-56-07-SFF14.txt^{(9.8KB, txt)}

File S15

Ascocoryne all-taxa Bayesian phylogenetic tree.

per-2026-56-07-SFF15.txt^{(167.2KB, txt)}

File S16

Ascocoryne 2-minimum maximum likelihood phylogenetic tree.

per-2026-56-07-SFF16.txt^{(5.4KB, txt)}

File S17

Ascocoryne all-taxa maximum likelihood phylogenetic tree.

per-2026-56-07-SFF17.txt^{(9.6KB, txt)}

File S18

Ascocoryne ITS-only Bayesian phylogenetic tree.

per-2026-56-07-SFF18.txt^{(158.3KB, txt)}

File S19

Ascocoryne ITS-only maximum likelihood phylogenetic tree.

per-2026-56-07-SFF19.txt^{(9.2KB, txt)}

File S20

CTP barcode analysis distance matrix.

per-2026-56-07-SFF20.xlsx^{(26.1KB, xlsx)}

File S21

ITS barcode analysis distance matrix.

per-2026-56-07-SFF21.xlsx^{(23.6KB, xlsx)}

File S22

CTP barcode phylogenetic tree.

per-2026-56-07-SFF22.txt^{(2.6KB, txt)}

File S23

ITS barcode phylogenetic tree.

per-2026-56-07-SFF23.txt^{(2.4KB, txt)}

Table S1

Herbarium and GenBank accession numbers for collections with associated metadata.

per-2026-56-07-ST1.xlsx^{(40.7KB, xlsx)}

Table S2

Information on alignments and models selected for phylogenetic analyses.

per-2026-56-07-ST2.xlsx^{(18.1KB, xlsx)}

Table S3

Summary of Ascocoryne species supported in phylogenetic analyses.

per-2026-56-07-ST3.xlsx^{(17.9KB, xlsx)}

Table S4

Identifications of Gelatinodiscaceae collections from Wilson (1962) and this study.

per-2026-56-07-ST4.xlsx^{(16.3KB, xlsx)}

Table S5

Notes on Coryne taxa with teleomorphs that have not been transferred outside of Coryne or Ascocoryne.

per-2026-56-07-ST5.xlsx^{(20.9KB, xlsx)}

[R1] Abarenkov K, Zirk A, Piirmann T, et al. (2024). UNITE general FASTA release for Fungi version 04.04.2024. UNITE Community. 10.15156/BIO/2959332 [DOI]

[R2] Albu S, Toome M, Aime MC. (2015). Violaceomyces palustris gen. et sp. nov. and a new monotypic lineage, Violaceomycetales ord. nov. in Ustilaginomycetes. Mycologia 107: 1193–1204. 10.3852/14-260 [DOI] [PubMed] [Google Scholar]

[R3] Altschul SF, Gish W, Miller W, et al. (1990). Basic local alignment search tool. Journal of Molecular Biology 215: 403–410. 10.1016/s0022-2836(05)80360-2 [DOI] [PubMed] [Google Scholar]

[R4] Arhipova N, Jansons A, Zaluma A, et al. (2015). Bark stripping of Pinus contorta caused by moose and deer: wounding patterns, discoloration of wood, and associated fungi. Canadian Journal of Forest Research 45: 1434–1438. [Google Scholar]

[R5] Baral HO. (1992). Vital versus herbarium taxonomy: morphological differences between living and dead cells of Ascomycetes, and their taxonomic implications. Mycotaxon 44: 333–390. [Google Scholar]

[R6] Baral HO. (1999). Ombrophila hemiamyloidea, an aquatic discomycete. Mycologia Bavarica 3: 50–63. [Google Scholar]

[R7] Baral HO. (2000). Key to Ascocoryne. www.ascofrance.com/uploads/forum_file/5598.doc. Accessed on 4 May 2025.

[R8] Baral HO, Galan R, Platas G, et al. (2013a). Phaeohelotium undulatum comb. nov. and Phaeoh. succineoguttulatum sp. nov., two segregates of the Discinella terrestris aggregate found under Eucalyptus in Spain: taxonomy, molecular biology, ecology and distribution. Mycosystema 32: 386–428. [Google Scholar]

[R9] Baral HO, Haelewaters D, Pärtel K. (2015). A new attempt to classify the families of the Helotiales. Second International Workshop on Ascomycete Systematics: 26. 22–24 April 2015, Amsterdam. [Google Scholar]

[R10] Baral HO, Marson G, Bogale M, et al. (2013b). Xerombrophila crystallifera, a new genus and species in the Helotiales. Mycological Progress 12: 475–488. 10.1007/s11557-012-0854-6 [DOI] [Google Scholar]

[R11] Berkeley MJ. (1868, ‘1869’) On a collection of fungi from Cuba. Part. II., including those belonging to the families Gasteromycetes, Coniomycetes, Hyphomycetes, Physomycetes, and Ascomycetes. Botanical Journal of the Linnean Society 10: 341–392 [Google Scholar]

[R12] Berkeley MJ, Curtis MA. (1868, ‘1869’). Fungi Cubenses. (Hymenomycetes). Botanical Journal of the Linnean Society 10: 280–341. [Google Scholar]

[R13] Beug MW, Bessette AE, Bessette AR. (2014). Ascomycete Fungi of North America: A Mushroom Reference Guide. University of Texas Press, USA. [Google Scholar]

[R14] Bouckaert R, Vaughan TG, Barido-Sottani J, et al. (2019). BEAST 2.5: An advanced software platform for Bayesian evolutionary analysis. PLOS Computational Biology 15: e1006650. 10.1371/journal.pcbi.1006650 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R15] Bouckaert RR, Drummond AJ. (2017). bModelTest: Bayesian phylogenetic site model averaging and model comparison. BMC Evolutionary Biology 17: 42. 10.1186/s12862-017-0890-6 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R16] Bunyard B, Wang Z, Malloch D, et al. (2008). New North American records for Ascocoryne turficola (Ascomycota: Helotiales). Fungi Magazine 1: 23–31. [Google Scholar]

[R17] Burow K, Grawunder A, Harpke M, et al. (2019). Microbiomes in an acidic rock–water cave system. FEMS Microbiology Letters 366: fnz167. 10.1093/femsle/fnz167 [DOI] [PubMed] [Google Scholar]

[R18] Callahan BJ, McMurdie PJ, Rosen MJ, et al. (2016). DADA2: High-resolution sample inference from Illumina amplicon data. Nature Methods 13: 581–583. 10.1038/nmeth.3869 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R19] Capella-Gutiérrez S, Silla-Martínez JM, Gabaldón T. (2009) trimAl: a tool for automated alignment trimming in large-scale phylogenetic analyses. Bioinformatics 25: 1972–1973. 10.1093/bioinformatics/btp348 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R20] Carbone I, White JB, Miadlikowska J, et al. (2019). T-BAS Version 2.1: Tree-Based Alignment Selector Toolkit for evolutionary placement of DNA sequences and viewing alignments and specimen metadata on curated and custom trees. Microbiology Resource Announcements 8: 10.1128/mra.00328-19. 10.1128/mra.00328-19 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R21] Carpenter SE. (1976). Taxonomy, morphology and ontogeny of Gelatinodiscus flavidus. Mycotaxon 3: 209–232. 10.5962/p.413977 [DOI] [Google Scholar]

[R22] Chernomor O, Minh BQ, von Haeseler A. (2015). Consequences of common topological rearrangements for partition trees in phylogenomic inference. Journal of Computational Biology 22: 1129–1142. 10.1089/cmb.2015.0146 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R23] Crous PW, Jurjević Ž, Balashov S, et al. (2024). Fungal Planet description sheets: 1614–1696. Fungal Systematics and Evolution 13: 183–440. 10.3114/fuse.2024.13.11 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R24] Dennis RWG. (1954). Some inoperculate discomycetes of tropical America. Kew Bulletin 9: 289–348. 10.2307/4114399 [DOI] [Google Scholar]

[R25] Dennis RWG. (1955) Two proposed new genera of Helotiales. Kew Bulletin 10: 359–362. 10.2307/4109223 [DOI] [Google Scholar]

[R26] Dennis RWG. (1956) A revision of the British Helotiaceae in the herbarium of the Royal Botanic Gardens, Kew, with notes on related European species. Mycological Papers 62: 1–216. [Google Scholar]

[R27] Dennis RWG. (1958) Critical notes on some Australian Helotiales and Ostropales. Kew Bulletin 13: 321–358. 10.2307/4109542 [DOI] [Google Scholar]

[R28] Dennis RWG. (1971). New or interesting British microfungi. Kew Bulletin 25: 335–374. 10.2307/4103238 [DOI] [Google Scholar]

[R29] Douglas J, Zhang R, Bouckaert R. (2021). Adaptive dating and fast proposals: Revisiting the phylogenetic relaxed clock model. PLOS Computational Biology 17: e1008322. 10.1371/journal.pcbi.1008322 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R30] Ekanayaka A, Hyde K, Gentekaki E, et al. (2019). Preliminary classification of Leotiomycetes. Mycosphere 10: 310–489. 10.5943/mycosphere/10/1/7 [DOI] [Google Scholar]

[R31] Filippova N, Bulyonkova TM. (2013). Notes on the ecology of Ascocoryne turficola (Ascomycota: Helotiales) in West Siberia. Environmental Dynamics and Global Climate Change 4: 1–6. 10.17816/edgcc421-6 [DOI] [Google Scholar]

[R32] Filippova N, Zvyagina E, Bulyonkova T. (2013). Ascocoryne turficola (Boud.) Korf records from West Siberia. Fungi Magazine 6: 26–30. [Google Scholar]

[R33] Forin N, Nigris S, Voyron S, et al. (2018). Next generation sequencing of ancient fungal specimens: The case of the Saccardo Mycological Herbarium. Frontiers in Ecology and Evolution 6: 129. 10.3389/fevo.2018.00129 [DOI] [Google Scholar]

[R34] Gao Q, Yan D, Wang K, et al. (2017). Metabolites diversity and hydrocarbons analysis on volatile organic compounds produced by Ascocoryne strains. Forest Research 30: 376–383. [Google Scholar]

[R35] Gardes M, Bruns TD. (1993). ITS primers with enhanced specificity for basidiomycetes - application to the identification of mycorrhizae and rusts. Molecular Ecology 2: 113–118. 10.1111/j.1365-294X.1993.tb00005.x [DOI] [PubMed] [Google Scholar]

[R36] Griffin MA, Spakowicz DJ, Gianoulis TA, et al. (2010). Volatile organic compound production by organisms in the genus Ascocoryne and a re-evaluation of myco-diesel production by NRRL 50072. Microbiology 156: 3814–3829. [DOI] [PMC free article] [PubMed] [Google Scholar]

[R37] Groves JW, Wilson DE. (1967). The nomenclatural status of Coryne. Taxon 16: 35–41. 10.2307/1217104 [DOI] [Google Scholar]

[R38] Grum-Grzhimaylo OA, Debets AJM, Bilanenko EN. (2016). The diversity of microfungi in peatlands originated from the White Sea. Mycologia 108: 233–254. 10.3852/14-346 [DOI] [PubMed] [Google Scholar]

[R39] Guindon S, Dufayard J-F, Lefort V, et al. (2010). New algorithms and methods to estimate maximum-likelihood phylogenies: Assessing the performance of PhyML 3.0. Systematic Biology 59: 307–21. 10.1093/sysbio/syq010. [DOI] [PubMed] [Google Scholar]

[R40] Haelewaters D, Park D, Johnston PR. (2021). Multilocus phylogenetic analysis reveals that Cyttariales is a synonym of Helotiales. Mycological Progress 20: 1323–1330. 10.1007/s11557-021-01736-2 [DOI] [Google Scholar]

[R41] Haelewaters D, Matthews TJ, Wayman JP, et al. (2024). Biological knowledge shortfalls impede conservation efforts in poorly studied taxa — A case study of Laboulbeniomycetes. Journal of Biogeography 51: 29–39. 10.1111/jbi.14725 [DOI] [Google Scholar]

[R42] Hawksworth DL. (2001). The magnitude of fungal diversity: the 1.5 million species estimate revisited. Mycological Research 105: 1422–1432. 10.1017/S0953756201004725 [DOI] [Google Scholar]

[R43] Hebert PDN, Stoeckle MY, Zemlak TS, et al. (2004). Identification of birds through DNA barcodes. PLoS Biology 2: e312. 10.1371/journal.pbio.0020312 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R44] Held BW, Salomon CE, Blanchette RA. (2020). Diverse subterranean fungi of an underground iron ore mine. PLoS ONE 15: e0234208. 10.1371/journal.pone.0234208 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R45] Hennings P. (1900). Fungi. In: Monsunia: Beiträge zur kenntniss der vegetation des süd- und ostasiatischen monsungebietes (Warburg O, ed). W. Engelmann, Germany: 30–31. [Google Scholar]

[R46] Herrera P, Suárez JP, Kottke I. (2010). Orchids keep the ascomycetes outside: a highly diverse group of ascomycetes colonizing the velamen of epiphytic orchids from a tropical mountain rainforest in Southern Ecuador. Mycology 1: 262–268. 10.1080/21501203.2010.526645 [DOI] [Google Scholar]

[R47] Hoang DT, Chernomor O, von Haeseler A, et al. (2018). UFBoot2: Improving the ultrafast bootstrap approximation. Molecular Biology and Evolution 35: 518–522. 10.1093/molbev/msx281 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R48] Hofstetter V, Miadlikowska J, Kauff F, et al. (2007). Phylogenetic comparison of protein-coding versus ribosomal RNA-coding sequence data: A case study of the Lecanoromycetes (Ascomycota). Molecular Phylogenetics and Evolution 44: 412–426. 10.1016/j.ympev.2006.10.016 [DOI] [PubMed] [Google Scholar]

[R49] Hyde KD, Noorabadi MT, Thiyagaraja V, et al. (2024). The 2024 Outline of Fungi and fungus-like taxa. Mycosphere 15: 5146–6239. 10.5943/mycosphere/15/1/25 [DOI] [Google Scholar]

[R50] Index Fungorum (2025). Search Index Fungorum. https://www.indexfungorum.org/Names/Names.asp

[R51] Jaén-Molina R, Marrero-Rodríguez Á, Reyes-Betancort JA, et al. (2015). Molecular taxonomic identification in the absence of a ‘barcoding gap’: a test with the endemic flora of the Canarian oceanic hotspot. Molecular Ecology Resources 15: 42–56. 10.1111/1755-0998.12292 [DOI] [PubMed] [Google Scholar]

[R52] Jaklitsch W, Baral HO, Lücking R, et al. (2016). Syllabus of plant families: A. Engler’s syllabus der Pfanzenfamilien part 1/2 (Frey W. ed.). Borntraeger Gebrueder, Germany. [Google Scholar]

[R53] Johnston PR, Baschien C. (2020). Tricladiaceae fam. nov. (Helotiales, Leotiomycetes). Fungal Systematics and Evolution 6: 233–242. 10.3114/fuse.2020.06.10 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R54] Johnston PR, Park D. (2025). An updated multigene phylogeny of Leotiomycetes. Persoonia 55: 527–566. 10.3114/persoonia.2025.55.13 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R55] Johnston PR, Quijada L, Smith CA, et al. (2019). A multigene phylogeny toward a new phylogenetic classification of Leotiomycetes. IMA Fungus 10: 1. 10.1186/s43008-019-0002-x [DOI] [PMC free article] [PubMed] [Google Scholar]

[R56] Johnston PR, Seifert KA, Stone JK, et al. (2014). Recommendations on generic names competing for use in Leotiomycetes (Ascomycota). IMA Fungus 5: 91–120. 10.5598/imafungus.2014.05.01.11 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R57] Kalyaanamoorthy S, Minh BQ, Wong TKF, et al. (2017). ModelFinder: fast model selection for accurate phylogenetic estimates. Nature Methods 14: 587–589. 10.1038/nmeth.4285 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R58] Karakehian J, Quijada L, Pfister D, et al. (2021). Methods for observing, culturing, and studying living ascospores. Asian Journal of Mycology 4: 1–18. [Google Scholar]

[R59] Katoh K, Misawa K, Kuma K, et al. (2002). MAFFT: a novel method for rapid multiple sequence alignment based on fast Fourier transform. Nucleic Acids Research 30: 3059–3066. 10.1093/nar/gkf436 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R60] Katoh K, Standley DM. (2013). MAFFT multiple sequence alignment software version 7: improvements in performance and usability. Molecular Biology and Evolution 30: 772–780. 10.1093/molbev/mst010 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R61] Kosonen T, Huhtinen S, Hansen K. (2021). Taxonomy and systematics of Hyaloscyphaceae and Arachnopezizaceae. Persoonia 46: 26–62. 10.3767/persoonia.2021.46.02 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R62] Kučera V, Lizon P. (2005). Ascocoryne striata, comb. nov. Mycotaxon 93: 163–165. [Google Scholar]

[R63] Larena I, Salazar O, González V, et al. (1999). Design of a primer for ribosomal DNA internal transcribed spacer with enhanced specificity for ascomycetes. Journal of Biotechnology 75: 187–194. 10.1016/s0168-1656(99)00154-6 [DOI] [PubMed] [Google Scholar]

[R64] Liu YJ, Whelen S, Hall BD. (1999). Phylogenetic relationships among ascomycetes: evidence from an RNA polymerse II subunit. Molecular Biology and Evolution 16: 1799–1808. 10.1093/oxfordjournals.molbev.a026092 [DOI] [PubMed] [Google Scholar]

[R65] Matheny PB, Liu YJ, Ammirati JF, et al. (2002). Using RPB1 sequences to improve phylogenetic inference among mushrooms (Inocybe, Agaricales). American Journal of Botany 89: 688–698. 10.3732/ajb.89.4.688 [DOI] [PubMed] [Google Scholar]

[R66] Melie T. (2023). Genomic investigations in Ascomycota: Leveraging machine learning, phylogenetics, and comparative genomics methods to explore evolution and ecological specialization. Ph.D. dissertation. Department of Ecology and Evolutionary Biology, University of Colorado at Boulder, USA. https://scholar.colorado.edu/concern/graduate_thesis_or_dissertations/8049g6598 [Google Scholar]

[R67] Meyer CP, Paulay G. (2005). DNA barcoding: Error rates based on comprehensive sampling. PLoS Biology 3: e422. 10.1371/journal.pbio.0030422 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R68] Meyn R, Langer GJ, Gross A, et al. (2019). Fungal colonization patterns in necrotic rootstocks and stem bases of dieback-affected Fraxinus excelsior L. Forest Pathology 49: e12520. 10.1111/efp.12520 [DOI] [Google Scholar]

[R69] Miadlikowska J, Lutzoni F. (2000). Phylogenetic revision of the genus Peltigera (lichen‐forming Ascomycota) based on morphological, chemical, and large subunit nuclear ribosomal DNA Data. International Journal of Plant Sciences 161: 925–958. 10.1086/317568 [DOI] [Google Scholar]

[R70] Miller AN, Karakehian J, Raudabaugh DB. (2022). Next-generation sequencing of ancient and recent fungarium specimens. Journal of Fungi 8: 932. 10.3390/jof8090932 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R71] Molina L, Rajchenberg M, de Errasti A, et al. (2020). Sapwood-inhabiting mycobiota and Nothofagus tree mortality in Patagonia: Diversity patterns according to tree species, plant compartment and health condition. Forest Ecology and Management 462: 117997. 10.1016/j.foreco.2020.117997 [DOI] [Google Scholar]

[R72] Moyne G. (2012). Deux ascomycètes méconnus: Ascocoryne inflata et Rutstroemia alni. Bulletin Annuel de la Fédération Mycologique de l’Est 10: 21–24. [Google Scholar]

[R73] Nees von Esenbeck CG. (1816). Das System der Plze und Schwämme. Würzburg, Germany. [Google Scholar]

[R74] Nguyen L-T, Schmidt HA, von Haeseler A, et al. (2015). IQ-TREE: A fast and effective stochastic algorithm for estimating maximum-likelihood phylogenies. Molecular Biology and Evolution 32: 268–274. 10.1093/molbev/msu300 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R75] Olds CG, Berta-Thompson JW, Loucks JJ, et al. (2023), Applying a modified metabarcoding approach for the sequencing of macrofungal specimens from fungarium collections. Applications in Plant Sciences 11: e11508. 10.1002/aps3.11508 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R76] Penzig O, Saccardo PA. (1902). Diagnoses fungorum novorum in insula Java collectorum. Series Tertia. Discomycetes. Malpighia: rassegna mensuale di botanica 15: 201–260. [Google Scholar]

[R77] Quandt A, Haelewaters D. (2021). Phylogenetic advances in Leotiomycetes, an understudied clade of taxonomically and ecologically diverse Fungi. In: Encyclopedia of mycology Vol. 1 (Zaragoza Ó, Casadevall A. eds). Elsevier, United Kingdom. 10.1016/b978-0-12-819990-9.00052-4 [DOI] [Google Scholar]

[R78] Quijada L, Baral HO, Johnston PR, et al. (2022). A review of Hyphodiscaceae. Studies in Mycology 103: 59–85. 10.3114/sim.2022.103.03 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R79] Quijada L, Ribes M, Negrín R, et al. (2017). Lignicolous species of Helotiales associated with major vegetation types in the Canary Islands. Willdenowia 47: 271–291. 10.3372/wi.47.47310 [DOI] [Google Scholar]

[R80] R Foundation for Statistical Computing (2023). R Core Team. R: A Language and Environment for Statistical Computing. Austria. [Google Scholar]

[R81] Raudabaugh DB, Bach EM, Allen JM, et al. (2021). Fungal communities of submerged fine detritus from temperate peatland and stream habitats. Aquatic Microbial Ecology 86: 191–207. 10.3354/ame01969 [DOI] [Google Scholar]

[R82] Rehner SA, Buckley E. (2005). A Beauveria phylogeny inferred from nuclear ITS and EF1-α sequences: evidence for cryptic diversification and links to Cordyceps teleomorphs. Mycologia 97: 84–98. 10.3852/mycologia.97.1.84 [DOI] [PubMed] [Google Scholar]

[R83] Roll-Hansen F, Roll-Hansen H. (1979a). Microflora of sound-looking wood in Picea abies stems. European Journal of Forest Pathology 9: 308–316. 10.1111/j.1439-0329.1979.tb00693.x [DOI] [Google Scholar]

[R84] Roll-Hansen F, Roll-Hansen H. (1979b). Ascocoryne sarcoides and Ascocoryne cylichnium. Descriptions and comparison. Norwegian Journal of Botany 26: 193–206. [Google Scholar]

[R85] Saccardo PA, Traverso GB, Trotter A. (1889). Sylloge fungorum omnium hucusque cognitorum Vol. VIII. Patavii, Italy. [Google Scholar]

[R86] Seaver J. (1951). The North American cup-fungi (Inoperculates). Lancaster Press, USA. [Google Scholar]

[R87] Seifert KA. (1989). Coryne trichophora, comb. nov., and the implications of its conidiomatal anatomy. Studies in Mycology 31: 157–164. [Google Scholar]

[R88] Smith AL. (1901). On some Fungi from the West Indies. Botanical Journal of the Linnean Society 35: 1–19. 10.1111/j.1095-8339.1901.tb02317.x [DOI] [Google Scholar]

[R89] Sri-indrasutdhi V, Tsui CKM, Chuaseeharonnachai C, et al. (2015). Helicocentralis hyalina gen. et sp. nov., an aero-aquatic helicosporous fungus (Leotiomycetes, Ascomycota) in Thailand. Mycological Progress 14: 81. 10.1007/s11557-015-1103-6 [DOI] [Google Scholar]

[R90] Stallman JK, Johnston PR, Lickey EB, et al. (2024). Recent fieldwork and fungarium studies double known diversity of Chlorosplenium and improve understanding of species distributions. Mycologia 116: 993–1018. 10.1080/00275514.2024.2364567 [DOI] [PubMed] [Google Scholar]

[R91] Stallman JK, Haelewaters D, Koch Bach RA, et al. (2024). The contribution of tropical long-term studies to mycology. IMA Fungus 15: 35. 10.1186/s43008-024-00166-5 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R92] Stielow JB, Lévesque CA, Seifert KA, et al. (2015). One fungus, which genes? Development and assessment of universal primers for potential secondary fungal DNA barcodes. Persoonia 35: 242–263. 10.3767/003158515x689135 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R93] Stiller JW, Hall BD. (1997). The origin of red algae: Implications for plastid evolution. Proceedings of the National Academy of Sciences USA 94: 4520–4525. 10.1073/pnas.94.9.4520 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R94] Tamura K, Stecher G, Kumar S. (2021). MEGA11: Molecular Evolutionary Genetics Analysis Version 11. Molecular Biology and Evolution 38: 3022–3027. 10.1093/molbev/msab120 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R95] Tanney JB, Seifert KA. (2020). Mollisiaceae: An overlooked lineage of diverse endophytes. Studies in Mycology 95: 293–380. 10.1016/j.simyco.2020.02.005 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R96] Taylor JW, Jacobson DJ, Kroken S, et al. (2000). Phylogenetic species recognition and species concepts in Fungi. Fungal Genetics and Biology 31: 21–32. 10.1006/fgbi.2000.1228 [DOI] [PubMed] [Google Scholar]

[R97] Thind KS, Singh H. (1972). The Helotiales of India—XVI. Journal of the Indian Botanical Society 51: 44–51. [Google Scholar]

[R98] Untereiner WA, Yue Q, Chen L, et al. (2019). Phialophora section Catenulatae disassembled: New genera, species, and combinations and a new family encompassing taxa with cleistothecial ascomata and phialidic asexual states. Mycologia 111: 998–1027. 10.1080/00275514.2019.1663106 [DOI] [PubMed] [Google Scholar]

[R99] Vaidya G, Lohman DJ, Meier R. (2011). SequenceMatrix: concatenation software for the fast assembly of multi-gene datasets with character set and codon information. Cladistics 27: 171–180. 10.1111/j.1096-0031.2010.00329.x [DOI] [PubMed] [Google Scholar]

[R100] Valenzuela R, Raymundo T, Reyes P, et al. (2021). Ascomycetes from the relic forest of Oreomunnea mexicana, Oaxaca, Mexico. Phytotaxa 528: 19–44. 10.11646/phytotaxa.528.1.3 [DOI] [Google Scholar]

[R101] Vandegrift R, Newman DS, Dentinger BTM, et al. (2023). Richer than gold: the fungal biodiversity of Reserva Los Cedros, a threatened Andean cloud forest. Botanical Studies 64: 17. 10.1186/s40529-023-00390-z [DOI] [PMC free article] [PubMed] [Google Scholar]

[R102] Vasiliauskas R, Stenlid J. (1998). Fungi inhabiting stems of Picea abies in a managed stand in Lithuania. Forest Ecology and Management 109: 119–126. 10.1016/S0378-1127(98)00226-6 [DOI] [Google Scholar]

[R103] Vellinga EC, Kuyper TW, Ammirati J, et al. (2015). Six simple guidelines for introducing new genera of fungi. IMA Fungus 6: A65–A68. 10.1007/BF03449356 [DOI] [Google Scholar]

[R104] Vilgalys R, Hester M. (1990). Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species. Journal of Bacteriology 172: 4238–4246. 10.1128/jb.172.8.4238-4246.1990 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R105] Voglmayr H. (2004). Spirosphaera cupreorufescens sp. nov., a rare aeroaquatic fungus. Studies in Mycology 50: 221–228. [Google Scholar]

[R106] Vu D, Groenewald M, de Vries M, et al. (2019). Large-scale generation and analysis of filamentous fungal DNA barcodes boosts coverage for kingdom fungi and reveals thresholds for fungal species and higher taxon delimitation. Studies in Mycology 92: 135–154. 10.1016/j.simyco.2018.05.001 [DOI] [PMC free article] [PubMed] [Google Scholar]

[R107] Wang Z, Binder M, Hibbett DS. (2005). Life history and systematics of the aquatic discomycete Mitrula (Helotiales, Ascomycota) based on cultural, morphological, and molecular studies. American Journal of Botany 92: 1565–1574. 10.3732/ajb.92.9.1565 [DOI] [PubMed] [Google Scholar]

[R108] Wang J, Ji-li E, Liu J. (2017). Endophytic fungi diversity of Platycladus orientalis in different age. Journal of Southwest Forestry University 37: 113–117. 10.11929/j.issn.2095-1914.2017.03.018 [DOI] [Google Scholar]

[R109] White T, Bruns T, Lee S, et al. (1990). Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: PCR Protocols: A Guide to Methods and Applications (Innis MA, Gelfand DH, Sninsky JJ, et al. eds.). Academic Press, USA: 315–322. [Google Scholar]

[R110] Wijayawardene NN, Hyde KD, Lumbsch HT, et al. (2018). Outline of Ascomycota: 2017. Fungal Diversity 88: 167–263. 10.1007/s13225-018-0394-8 [DOI] [Google Scholar]

[R111] Wilson D. (1962). A study of eight North American species of Coryne. Ph.D. dissertation. School of Graduate Studies, University of Toronto, Canada. [Google Scholar]

[R112] Wu B, Tian J, Bai C, et al. (2013). The biogeography of fungal communities in wetland sediments along the Changjiang River and other sites in China. The ISME Journal 7: 1299–1309. 10.1038/ismej.2013.29 [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

Phylogeny of Gelatinodiscaceae (Leotiomycetes) with emphasis on the taxonomy of Ascocoryne in North America

JK Stallman

L Quijada

MC Aime

CA Quandt

DB Raudabaugh

T Melie

I Rimer

S McAdam

D Haelewaters

Abstract

INTRODUCTION

Gelatinodiscaceae

Ascocoryne

MATERIALS AND METHODS

Collections

Morphology

Sanger sequencing

Table 1.

High-throughput sequencing and genome mining

Construction of phylogenetic trees

Barcode analysis

RESULTS

High-throughput sequencing

Phylogenetic placement of Clathrosporium, Phaeangellina, Skyathea, and Xerombrophila

Gelatinodiscaceae phylogenies

Fig. 1.

Ascocoryne phylogenies — intrageneric clades and morphology

Fig. 2.

Ascocoryne — support for described species and species richness

ITS and CTP barcode analysis

Table 2.

TAXONOMY

Fig. 3.

Fig. 4.

Fig. 5.

Fig. 6.

Fig. 7.

Fig. 8.

Fig. 9.

Fig. 10.

Fig. 11.

DISCUSSION

Genera in Gelatinodiscaceae

Intrageneric clades and morphological characters in Ascocoryne

Species richness of Ascocoryne with notes on individual taxa

Ascocoryne javanica

Ascocoryne cylichnium

Ascocoryne striata

Ascocoryne (Coryne) species reported in Wilson’s thesis and the utility of cultural data

Names described in Coryne

Evaluating the ITS and CTP loci as DNA barcodes of Ascocoryne species

CONCLUSIONS

Acknowledgments

Footnotes

SUPPLEMENTARY MATERIAL

REFERENCES

Associated Data

Supplementary Materials

ACTIONS

PERMALINK

RESOURCES

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Links to NCBI Databases