One stop shop VI: taxonomic update with molecular phylogeny for important phytopathogenic genera: 126–150

Table of Contents and contributors

126. Coleosporium - AK Gautam, S Avasthi, RK Verma, CS Bhunjun

127. Dichotomophthora - SC Karunarathna, DJ Soares, CS Bhunjun

128. Didymella - D Gomdola, CS Bhunjun

129. Leptosphaerulina - D Gomdola, CS Bhunjun, DJ Soares

130. Macrophomina - DJ Soares, CS Bhunjun, A Alghamdi

131. Melampsora - AK Gautam, S Avasthi, RK Verma

132. Myrothecium - CS Bhunjun, C Phukhamsakda

133. Neopseudocercosporella - N Gunasinghe, CS Bhunjun

134. Neopestalotiopsis - C Phukhamsakda, CS Bhunjun

135. Neosetophoma - CS Bhunjun, C Phukhamsakda, DJ Soares

136. Nigrospora - S Tibpromma, CS Bhunjun

137. Oculimacula - X Tang, RS Jayawardena

138. Phaeosphaeriopsis - SC Karunarathna, C Phukhamsakda, CS Bhunjun

139. Phakopsora - AK Gautam, S Avasthi, RK Verma, CS Bhunjun

140. Pleiocarpon - CS Bhunjun, C Phukhamsakda, DJ Soares

141. Pseudopestalotiopsis - C Phukhamsakda, CS Bhunjun

142. Pseudogymnoascus - C Phukhamsakda, CS Bhunjun

143. Pyrenophora - YR Sun, CS Bhunjun

144. Ramichloridium - YR Sun, RS Jayawardena, CS Bhunjun

145. Ravenelia - AK Gautam, S Avasthi, RK Verma, CS Bhunjun

146. Seiridium - C Phukhamsakda, CS Bhunjun, IF Halawani

147. Stenocarpella - X Tang, RS Jayawardena

148. Tubakia - C Phukhamsakda, CS Bhunjun

149. Zasmidium - DS Tennakoon, CS Bhunjun

150. Zymoseptoria - S Tibpromma, CS Bhunjun

Introduction

Fungi are a diverse and heterogeneous group of organisms with an estimated 1.5 to 12 million species (Hawksworth 1991, Wu et al. 2019, Hyde et al. 2024b). There are over 150,000 described species that have been reported with different lifestyles including endophytic, saprotrophic and parasitism (Hyde et al. 2020, Bhunjun et al. 2022, Phukhamsakda et al. 2022, 2025). As pathogens, fungi have profound implications for human health, agriculture, and ecosystems (Hyde et al. 2018, 2019, Fisher et al. 2020). Pathogenic fungi contribute significantly to infectious diseases in humans, ranging from superficial skin infections to life-threatening systemic mycoses (Köhler et al. 2017). In agriculture, pathogens cause devastating diseases in crops, leading to substantial yield losses and threatening food security on a global scale (Fones et al. 2020). The study of pathogens provides a better understanding of the complex interplay between fungi and their hosts, shedding light on mechanisms of infection and virulence factors, thus providing potential avenues for disease management (Zeilinger et al. 2013). There has been a surge in the number of fungal diseases which may have resulted in die-offs and extinction of various wild species (Fisher et al. 2012, Fisher & Denning 2023). A number of these diseases were caused by recently described pathogens, but there has also been a rise in the risks posed by emerging pathogens as a result of climate change (Blehert et al. 2009, Casadevall 2019, Seidel et al. 2024).

This is the sixth paper in the “One Stop Shop series”. Previous contributions have been vital in providing a stable platform for the taxonomy of phytopathogenic fungi and fungus-like organisms (Hyde et al. 2014, Jayawardena et al. 2019a, b, 2020, 2025, Table 1). The series has provided clarification on the taxonomy and classification of 125 genera and three families. In this study, we focus on 25 phytopathogenic genera and discuss the background information, distribution, hosts and disease symptoms for each genus.

Materials and methods

Isolates and Molecular analyses

The sequences of ex-type, ex-epitype or reference/voucher strains for all species were retrieved from GenBank. Polymerase chain reaction was used to amplify the partial gene regions of Dichotomophthora conlutea with primer pairs as described in Phukhamsakda et al. (2016) and sequenced by Shanghai Sangon Biological Engineering Technology & Services Co. (Shanghai, China). The large subunit ribosomal RNA (LSU) gene, internal transcribed spacer (ITS) region, the small subunit ribosomal RNA (SSU) gene, the translation elongation factor alpha (tef) gene, beta-tubulin (tub2), calmodulin (cal), actin (act), the glyceraldehyde-3-phosphate dehydrogenase (gapdh) gene, the cytochrome c oxidase (cox), histone H3 (his3), Minichromosome Maintenance Complex Component 7 gene (mcm7), the RNA polymerase II largest (rpb1) and second largest subunit (rpb2) genes were used in the phylogeny as per related bibliography. The sequences were aligned with MAFFT v. 7 (Katoh et al. 2019), with minimal adjustment using AliView v. 1.26 (Larsson 2014) and BioEdit v. 7.2.5 (Hall 1999). The concatenation of the multi-marker datasets was created by using Sequence Matrix v. 1.8 (Vaidya et al. 2011) and BioEdit v. 7.2.5. Phylogenetic analyses were performed using maximum likelihood, maximum parsimony and Bayesian inference posterior probabilities. Maximum likelihood analyses (ML) were performed on the CIPRES science gateway platform (Miller et al. 2017) using RAxML v. 8.2.12 (Stamatakis 2014). The general time reversible (GTR) model with a discrete gamma distribution plus invariant site (GTR + I + G) was used for nucleotide substitution. The best model for the different gene partitions was determined in JModelTest v. 2.1.10 or MrModeltest v. 2.3 for the Bayesian analysis (Nylander 2004, Darriba et al. 2012). Bayesian inference was conducted using MrBayes v. 3.2.7a on XSEDE (Ronquist and Huelsenbeck 2003).

Case studies

The genera are listed in alphabetical order and the classification follows Hyde et al. (2024b). For all the genera treated herein, we included species published until 30 May 2025 in the phylogenetic analyses (Index Fungorum 2025).

126. Coleosporium Lév., Annls Sci. Nat., Bot., sér. 3 8: 373 (1847)

Type species: Coleosporium campanulae (Pers.) Tul.

Classification: Pucciniomycetes, Pucciniales, Coleosporiaceae

Background: Coleosporium rusts are a group of plant pathogenic fungi belonging to Coleosporiaceae (Wijayawardene et al. 2022, Hyde et al. 2024a). Species of Coleosporium are obligate plant pathogens, similar to all rust fungi. The rust disease caused by these fungi is commonly known as Coleosporium rust, i.e., commonly referred to either by the genus or the host plant name, followed by the word “rust”. These fungi have been studied extensively as they cause diseases on ornamentals grown in gardens (McTaggart & Aime 2018).

Coleosporium was established by Léveille (1847) and comprises about 125 species worldwide (Kirk et al. 2008, Wijayawardene et al. 2020, 2022, Hyde et al. 2024a). These fungi have been documented from a significant number of telial hosts, but only one genus of aecial host, i.e., Pinus (Hedgcock 1928). About 12 species of Coleosporium infect Pinus densiflora as an aecial host (Hedgcock 1928). However, Coleosporium on pine trees have rarely been identified to the species level because of their morphological similarities (Suzuki et al. 2018). Scanning electron microscopy has been

used to describe these taxa based on surface sculptures of aeciospores and urediniospores along with different morphologies (Hiratsuka and Kaneko 1975). Two types of basidial development in Coleosporium were recognised by Kaneko (1981). In type 1, the entire cell becomes a basidium without change in length while in type two, the cell matures and a septum develops which separates the upper basidial part from the basal stalk-like part. Cummins (1978) and Kaneko (1981) described variations in the shape of the basidiospores. The genus has a wide geographic range, including almost every continent. The fungi primarily damage their telial hosts, which are often ornamentals, including aster, morning glories, and Plumeria. Plumeria rust caused by Coleosporium plumeriae is perhaps the most important example of a disease caused by the genus. DNA sequence data from the LSU and ITS regions available in GenBank for the species of Coleosporium were analysed by Gautam et al. (2021) who provided a taxonomic outline of Indian Pucciniales.

Distribution: Coleosporium spp. have been observed on almost every continent: North and South America, Asia, islands in the Pacific, Africa and Europe (McMillan 1984, Kolmer et al. 2009). These rust fungi have been reported from many countries including Argentina, Brazil, China, Cuba, Mexico, Georgia, France, India, Japan, Israel, Russia, Malta, Mongolia, Nepal, Philippines, São Tomé and Príncipe, Sierra Leone, Slovenia, Sri Lanka and the USA (Farr & Rossman 2025).

Disease symptoms: Coleosporium rust disease symptoms include chlorosis on leaves, dieback, leaf spotting and defoliation. The yellow-orange uredinia of these rust fungi often appear on the lower leaf surface with corresponding chlorotic spots on the upper surface (Fig. 1). Spermogonia are formed on the aecial host while aecia are formed on the lower surface. These rusts primarily damage their telial hosts, with infected leaves turning yellow to brown with premature leaf fall (Cummins & Hiratsuka 2003, Nelson 2009).

Hosts: species of Adenocaulon, Adenophora, Anemone, Aposeris, Aristolochia, Arnica, Arundina, Aster, Begonia, Brickellia, Cacalia, Calendula, Campanula, Campanumoea, Carpesium, Carphochaete, Choerospondias, Cirsium, Clematis, Clerodendrum, Codonopsis, Coleosanthus, Coleus, Convolvulus, Dahlia, Datisca, Erigeron, Erythrina, Eucommia, Euodia, Eupatorium, Exacum, Fauria, Heterotheca, Ipomoea, Knoxia, Labiatae, Lacinaria, Spigelia, Liabum, Ligularia, Lonicera, Lycopus, Madia, Microrhamnus, Myriactis, Myripnois, Orsina, Paederia, Pedicularis, Perezia, Perilla, Pertya, Phlomis, Phyllanthus, Pinus, Plumeria, Polymnia, Rubia, Salvia, Satyrium, Saussurea, Senecio, Serratula, Sicyos, Sida, Smilax, Stevia, Synurus, Thunbergia, Tylophora, Verbesina, Vernonia, Viburnum, Viola and Zanthoxylum (Farr & Rossman 2025).

Fig. 1. Coleosporium plumeriae on Plumeria sp. a-b Host plant with infected and healthy leaves. c-d Sori at various stages of infection. e-f Urediniospores. g Teliospores. Scale bars: 10 µm. This picture is copyright of Ajay Kumar Gautam, Shubhi Avasthi and Rajnish Kumar Verma.

Pathogen biology, disease cycle and epidemiology: Coleosporium comprises several species of rust fungi that are obligate biotrophic pathogens. These rust fungi possess a complex life cycle that typically alternates between pines and angiosperms, i.e., two hosts (heteroecious) and consisting of five spore types (macrocyclic), with spermogonia and aecia forming on needles of Pinus and uredinia and telia on both monocot and dicot plants (especially Asteraceae, but also on Campanulaceae, Orobanchaceae, Ranunculaceae) (Kirk et al. 2008). Coleosporium species can infect the same host repeatedly, or more than one host, in a single growing season. A single summer may allow up to 15 disease cycles. The disease cycle begins when basidiospores, produced from germinating teliospores, are wind-dispersed to infect the host plant (Harrington & Wingfield 2000). Upon landing on susceptible tissue, the basidiospores germinate and penetrate the plant's epidermis, initiating infection. The pathogen then forms uredinia, which produce urediniospores that facilitate the spread of the disease. The production of urediniospores continues throughout the summer showing the polycyclic nature of some species of Coleosporium (Chapell & Rausher 2016). In the late season, telia develop, producing teliospores that overwinter and serve as the primary inoculum for the next growing season. The epidemiology of Coleosporium is influenced by environmental factors such as humidity, temperature, and the presence of susceptible host plants (Chapell & Rausher 2016). Management strategies include removing infected plant material, applying fungicides, and planting resistant cultivars to reduce disease incidence and spread (McTaggart & Aime 2018).

Morphological-based identification: Coleosporium is characterised by 1-celled teliospores that are produced in a 1-layered crust under host epidermal cells (Cummins & Hiratsuka 2003). These taxa produce branched, septate mycelium and haustoria for absorption (Kolmer et al. 2009). They produce subepidermal, Group I (Type 2) spermogonia (Zhao et al. 2023). The colour of the spermogonia, the arrangement, size, and shape of the aecia, and the aecial host plant are three important morphological characteristics used to distinguish species of Coleosporium (Hedgcock 1928). Aecia are peridermium-type, subepidermal, erumpent with noticeable peridermium and found on the underside of leaves of the aecial host (abaxial). Aeciospores are catenulate, oblong to round in shape with verrucose spore walls. The aeciospores usually appear a month or two later than the spermogonia. These fungi produce subepidermal, erumpent, caeoma-type uredinia as bright orange pustules initially, fading to whitish later in the disease cycle. The uredinia develop catenuate, echinulate, reticulate, or verrucose urediniospores (with scattered or obscure pores) under the epidermis of the telial host plant. Telia produced by these fungi are often flat and dense, yellow or orange, wax-like, subepidermal, erumpent as low or columnar cushions and gelatinous when wet. Telia are composed of sessile, 1-celled spores, in 1-layered crusts, or pseudocatenuate by the intrusion of young spores among older spores, thick-walled, gelatinizing above teliospores. The teliospores germinate without dormancy and give rise to a phragmobasidium (a 4-celled internal basidium). Each cell of internal basidium produces a sterigma and one ellipsoidal to globoid basidiospore which infects the aecial host (Weir 1925, Hedgcock 1928, Arthur 1934, Cummins & Hiratsuka 2003).

Molecular-based identification: Maier et al. (2003) investigated LSU sequence data for 52 rust fungi of nine families including Coleosporium and showed that Coleosporium is monophyletic. Molecular data employing LSU sequence was used to characterize Coleosporium plectranthi on Perilla frutescens var. japonica (Yun et al. 2007). Holcomb & Aime (2010) explored Plumeria rust from Louisiana and Malaysia and identified the causal organism as C. plumeriae based on morpho-taxonomy. By using ITS and LSU sequences, phylogenetic analyses of Coleosporium on Solidago revealed the first molecular evidence of the North American rust C. solidaginis in Europe (Beenken et al. 2017). McTaggart & Aime (2018) carried out a study (ca. 60 collections) of Coleosporium infecting species of Asteraceae from North America. Based on phylogenetic analyses (ITS and LSU) and morphology of teliospores and basidia, McTaggart & Aime (2018) found that at least three species of Coleosporium occur on Solidago in North America rather than only one. Based on PCR-RFLP analyses, four species, C. asterum, C. clematidis-apiifoliae, C. lycopodis, and C. phellodendri were found to be associated with Pinus densiflora (Suzuki et al. 2018). Coleosporium zanthoxyli was identified on Zanthoxylum planispinum based on the host type, morphological characteristics, and phylogenetic analyses using ITS and LSU sequences (Shin et al. 2019). Both ITS and LSU are suitable loci for the identification of Coleosporium species. Aime & McTaggart (2020) resolved the deeper nodes of the rust fungus tree of life and provided an updated higher-rank classification for Pucciniales using LSU, SSU, and cox sequences from fresh and herbarium material of various species of rust fungi, including species of Coleosporium. This study provides an updated phylogeny of Coleosporium based on combined LSU and ITS sequence data (Fig. 2, Table 2).

Recommended genetic marker (genus level): ITS, LSU

Recommended genetic marker (species level): ITS, LSU

Accepted number of species: 125

References: Hiratsuka & Kaneko (1975), Cummins (1978), Kaneko (1981), Cummins & Hiratsuka (2003), Maier et al. (2003), Yun et al. (2007), Holcomb & Aime (2010), Beenken et al. (2017), Aime et al. (2018), McTaggart & Aime (2018), Suzuki et al. (2018), Shin et al. (2019), Aime & McTaggart (2020), Sun et al. (2024) (morphology and phylogeny)

127. Dichotomophthora Mehrl. & Fitzp. ex M.B. Ellis, Dematiaceous Hyphomycetes (Kew): 388 (1971)

Type species: Dichotomophthora portulacae Mehrl. & Fitzp. ex M.B. Ellis

Classification: Dothideomycetes, Pleosporales, Pleosporaceae

Background: Dichotomophthora was introduced by Mehrlich & Fitzpatrick (1935) to accommodate the hyphomycetous species D. portulacae, causing leaf spots on common purslane (Portulaca oleracea) in Hawaii. However, it was not validly published, and Rao (1966) attempted to validate it. Due to misapplications, this attempt was also unsuccessful, and the name was considered invalid until Ellis (1971) validly published it based on the holotype specimen of D. portulacae (Marin-Felix et al. 2019b). A second species D. lutea was introduced by De Hoog & van Oorschot (1983). The classification of Dichotomophthora in Pleosporaceae, Pleosporales was confirmed by Marin-Felix et al. (2019b) based on multi-gene phylogenetic analysis. Two further species D. basellae and D. brunnea were introduced, making a total of four accepted species in this genus (Marin-Felix et al. 2019b). Dichotomophthora species have been commonly reported as plant pathogens on various hosts (mostly P. oleracea), as saprobes and soil-inhabiting fungi and as a causative organism of human keratitis (Mehrlich & Fitzpatrick 1935, Ellis 1971, De Hoog & Oorschot 1983, Klisiewicz et al. 1983, Alfieri et al. 1984, Baudoin 1986, Pfeiffer et al. 1989, De Hoog et al. 2000, Eken 2003, Soares & Nechet 2017, Heidari et al. 2018, Marin-Felix et al. 2019b, Farr & Rossman 2025).

Distribution: worldwide (Farr & Rossman 2025)

Disease symptoms: Disease symptoms caused by Dichotomophthora typically manifest as dark, necrotic lesions (Fig. 3). The soil-inhabiting Dichotomophthora portulacae is known for causing rot, blight and dieback of the stems and leaf spots in Portulaca oleracea (Merhlich & Fitzpatrick 1935, Ellis 1971, Klisiewicz et al. 1983, Baudoin 1986, Mitchell 1986). The disease symptoms are characterised by necrotic, oval to irregular, pale brown spots with darker borders (Kliseiwicz 1985, Heidari et al. 2018, Abdulla 2021). Klisiewicz et al. (1983) observed that the stems of purslane plants, especially at the soil line, showed dark discolouration and constriction. The development of lesions was more frequently observed in parts of the stem that were in contact with the soil and the roots usually were infected by the same pathogen infecting the stems (Klisiewicz et al. 1983). Eken (2003) reported D. portulacae to cause crown and root rot of purslane. During experimental studies, symptoms of seed rot and post-emergence damping-off were observed within eight days, when all the seedlings died. In pathogenicity studies, sunken, necrotic lesions becameapparent on leaves and stems within 48 hours of inoculation. Later, the leaves turned yellow and dropped, while dark lesions developed on the stems causing dieback (Eken 2003). Dichotomophthora portulacae is more likely to cause the death of its host plants when infected at a young age (Kliseiwicz et al. 1983). Dichotomophthora lutea was reported producing brown to black lesions. The infection resulted in a

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A.

greyish dry rot and the gradual collapse of the tissue (Pfeiffer et al. 1989).

Hosts: Dichotomophthora portulacae was reported from Hawaii, USA on Portulaca oleracea (Mehrlich & Fitzpatrick 1935). Other hosts include Basella rubra, Nicotiana glauca, Portulaca grandiflora and P. lutea (Ellis 1971, Raabe et al. 1981, De Hoog & Oorschot 1983, Richardson 1990, Jing et al. 2008, Camino-Vilaro et al. 2019, Marin-Felix et al. 2019b, Farr & Rossman 2025). Dichotomophthora lutea has been isolated from Beta vulgaris (Minter et al. 2001), from soil, on seeds, stems, leaves and roots of Anredera cordifolia, Portulaca oleracea, Gymnocalycium mihanovichii var. friedrichii and Myrtillocactus geometrizans (De Hoog & Oorschot 1983, Miller 1990, Soares & Nechet 2017, Heidari et al. 2018, Camino-Vilaro et al. 2019, Marin-Felix et al. 2019b) in various regions, and from the seedbed of Pinus radiata in Italy (Heidari et al. 2018, Marin-Felix et al. 2019b). Dichotomophthora basellae was isolated from the leaves of Basella alba in Thailand (Marin-Felix et al. 2019b). The location and substrate of D. brunnea are unknown (Marin-Felix et al. 2019b).

Pathogen biology, disease cycle and epidemiology: Dichotomophthora portulacae is often reported as a major pathogen on purslane (Farr & Rossman 2025). It has been listed as an introduced plant pathogen in the United Kingdom (Jones & Baker 2007). According to Klisiewicz (1985), the fungus produces sclerotia and conidia on both artificial media and dead plant tissues. The sclerotia help the fungus to survive in the soil and act as an inoculum for basal stem infection (Klisiewicz 1985). The plant may become more prone to damage by insects or larvae following the stem infection by D. portulacae. Dichotomophthora portulacae prefers a temperature range of 27–33 °C and moist, humid conditions for growth and reproduction (in culture media). Thus, the fungus is less likely to infect plants where there is a lack of rain or moisture (Klisiewicz 1985). Once infected, the disease can develop regardless of the environmental conditions, leading to rapid death, especially in seedlings (Klisiewicz 1985, Mitchell 1986). Dichotomophthora portulacae can also infect carpetweed (Mollugo verticillata), but experimental results showed only mild symptoms (Mitchell 1986). In China, it was reported as an endophyte in Malabar spinach (Basella rubra) by Jing et al. (2008). The possibility of using D. portulacae as a biological control agent to control weeds, especially in areas with heavy rain and high humidity has been mentioned and investigated to an extent (Rader 1948, Klisiewicz et al. 1983, Klisiewicz 1985, Baudoin 1986, Vink & Elsas 2017). Alves et al. (2020) isolated Dichotomophthora as an endophyte from Jatropha curcas and two strains showed promising antioxidant activities. Further studies focusing on possible bioactive compounds of Dichotomophthora will lead to a better understanding of this genus.

Morphological-based identification: Dichotomophthora species are characterised by their macronematous, mononematous, dichotomously branched, irregularly orunbranched conidiophores, and simple, ellipsoidal to

Fig. 2. Maximum likelihood tree of Coleosporium species based on the concatenated LSU and ITS sequence data. The best scoring RAxML tree had a final likelihood value of -5967.127022. The tree was rooted to Chrysomyxa zhuoniensis (BJFC R00521). Estimated base frequencies were as follows: A = 0.308609, C = 0.152661, G =0.226954, and T = 0.311777; substitution rates AC = 1.138546, AG = 3.032190, AT = 3.398998, CG = 0.509751, CT = 6.693790, and GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

cylindrical, subhyaline, pale brown to dark brown conidia with 1–5 septa. The sexual morph is not known (Ellis 1971, Marin-Felix 2019b). The colonies of both D. basellae and D. lutea produce luteous, orange, diffusible pigments in culture (Marin-Felix et al. 2019b). Several reports of Dichotomophthora have remained uncertain due to fungal or host misidentifications, such as D. portulacae being confused with D. lutea in previous studies (De Hoog & van Oorschot 1983, Soares & Nechet 2017, Vink & Elsas 2017, Marin-Felix et al. 2019b). Therefore, more collections are needed to investigate their diversity.

Molecular-based identification: Heidari et al. (2018) used a combination of ITS and rpb2 sequence data to identify pathogenic strains of D. lutea from Iran. Marin-Felix et al. (2019b) included various strains of Dichotomophthora isolated from different hosts and regions using ITS, LSU, gapdh and rpb2 molecular data. As a result, four Dichotomophthora species (including two new species) were confirmed. In our phylogenetic analysis (Fig. 4), we included two isolates of Dichotomophthora (CMAA 1612 and CMAA 1613) from Anredera cordifolia (Soares & Nechet 2017). Their identification could not be confirmed in Soares and Nechet (2017) as the isolates (CMAA 1612 and CMAA 1613) only had ITS region. In this study, we provide additional sequence data for the isolate CMAA 1612 and we provide an updated phylogeny of Dichotomophthora based on combined ITS, LSU, gapdh and rpb2 sequence data (Fig. 4, Table 3). We also propose the new species Dichotomophthora conlutea (Fig. 3 and 4). Dichotomophthora cactacearum is excluded from our analysis as there are no description or molecular data for the species.

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A. Species confirmed with pathogenicity studies are marked with #.

Dichotomophthora conlutea D.J. Soares & Bhunjun, sp. nov.

Index Fungorum number: IF905057; Facesoffungi number: FoF 19465; Fig. 3

Etymology: Named based on its close affinity to Dichotomophthora lutea.

Conidiophores erect, simple or irregularly branched, sometimes verruculose near the apex, forming a stipe and head; stipe up to 900 μm long, 9–12 μm wide, pale brown to brown, smooth; head usually divided 3 to 4 times, frequently proliferating to produce additional head. Conidiogenous cells polytretic, integrated and terminal. Conidia 24–94 × 10–17 μm, solitary, smooth to finely verruculose, 1–6-septate, ellipsoidal to cylindrical, sometimes slightly curved, with rounded ends, initially subhyaline, becoming brown to reddish-brown. Sclerotia resembling immature perithecia, immersed in agar, abundant on potato dextrose agar and oatmeal agar, sporadic and scattered on malt extract agar. Sexual morph: unknown.

Culture characteristics: Colonies on PDA, MEA and OA reaching 50–60 mm after 1 week at 25 °C under ultraviolet light (12 h light, 12 h dark), white to grey (PDA and MEA) or dark grey to olivaceous (OA), cottony, velvety, somewhat fluffy, margin regular, effuse; reverse brown or dark-grey to black, periphery white to hazel. Diffusible pigment not observed under the culture conditions and media tested.

Material examined: Brazil, Jaguariúna, São Paulo, on symptomatic leaves of Anredera cordifolia, 16 January 2016, DJ Soares, DJS_771 (UB24608, holotype); ex-type CMAA 1612 = CMAA 1613.

Notes: The isolates (CMAA 1612 and CMAA1613) were described based on morphology and ITS sequences (Soares & Nechet 2017). The isolates were not assigned to a specific epithet as they lacked multi-locus data. In the analyses of combined ITS, LSU, gapdh and rpb2 sequence data, Dichotomophthora conlutea (CMAA 1612 and CMAA 1613) formed a sister clade to D. lutea isolates (Fig. 4). Dichotomophthora conlutea morphologically resemble D. lutea, however, D. conlutea differs by having conidiogenous head that are usually divided 3 to 4 times and frequently proliferating to produce additional heads (De Hoog et al. 1983, Soares & Nechet 2017). The conidia of D. conlutea are 1–6-septate while D. lutea have 0–4-septate conidia (De Hoog et al. 1983, Marin-Felix et al. 2019b). The two species also differ in the size of their conidia (24–94 × 10–17 μm vs 14–65.5 × 7.5–13 μm in D. lutea) (De Hoog et al. 1983, Soares & Nechet 2017, Marin-Felix et al. 2019b). Therefore, we introduce Dichotomophthora conlutea as a new species.

Recommended genetic markers (genus level): ITS

Recommended genetic markers (species level): ITS, LSU, gapdh and rpb2

Accepted number of species: five

References: Mehrlich & Fitzpatrick (1935), Routien (1957), Rao (1966), Ellis (1971), De Hoog & van Oorschot (1983) (morphology); Marin-Felix et al. (2019b) (phylogeny); Klisiewicz et al. (1983), Klisiewicz (1985), Baudoin (1986),

Fig. 3. Symptoms of Dichotomophthora conlutea on Anredera cordifolia (UB24608, holotype). a Initial symptoms. b Coalescent lesions with leaf yellowing. c Earlier senesced leaves under highly humid conditions showing profuse fungal sporulation. d-e Conidiophores apices showing conidiogenous heads. f-k Conidia. Scale bars: d = 50 μm, e, f, g, h, i, j, k = 10 μm. This picture is copyright of Dartanha Jose Soares.

Fig. 4. Maximum likelihood tree of Dichotomophthora species based on the concatenated ITS, LSU, gapdh and rpb2 sequence data. The best scoring RAxML tree had a final likelihood value of -6277.820. The tree was rooted to Curvularia buchloes (CBS 246.49) and C. subpapendorfii (CBS 656.74). Estimated base frequencies were as follows: A = A: 0.250, C = A: 0.250, G = A: 0.250, T = A: 0.250; substitution rates AC = 1.00000, AG = 1.90836, AT = 1.00000, CG = 1.00000, CT = 4.98741, GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Type strains are in bold. The new species is in blue. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

Mitchell (1986), Pfeiffer et al. (1989), Eken (2003), Soares & Nechet (2017) (pathogenicity)

128. Didymella Sacc., Michelia 2(6): 57 (1880)

Type species: Didymella exigua (Niessl) Sacc.

Classification: Dothideomycetes, Pleosporales, Didymellaceae

Background: Didymella was initially accommodated in Mycosphaerellaceae and later placed in Pleosporaceae, Phaeosphaeriaceae and Venturiaceae. Eventually, Didymellaceae was introduced to accommodate Didymella (De Gruyter et al. 2009). Didymella exigua has been accepted as the type or lectotype species of the genus by several authors (Höhnel 1918, Holm 1975, Corlett 1981). Didymella is a species-rich genus with a myriad of phytopathogens (Chen et al. 2015, 2017). As Didymella appears to be polyphyletic, its taxonomic revision and species delineation might be challenging as several species remain phylogenetically unresolved (Aveskamp et al. 2010, Chen et al. 2015).

Distribution: worldwide (Farr & Rossman 2025)

Disease symptoms: Didymella generally causes foliar diseases such as necrotic and tan leaf spots, and leaf blights, bracken blight disease, gummy stem blights, and black rot of fruits (Fig. 5). Didymella species cause dark sunken necrotic spots of table beets in New York (Vaghefi et al. 2016). Furthermore, a serious foliar disease was reported from baby lima bean fields in New York, initially exhibiting small necrotic spots that later coalesced and covered the entire leaf area (Gorny et al. 2016). Didymella glomerata was found to cause Didymella leaf blight on maize in China (Ma et al. 2022). The infected leaves displayed oval to oblong, grey sunken lesions, scattered over leaf surfaces. However, the stalks and other parts of the plant remained unaffected (Ma et al. 2022).

In addition to foliar diseases, Didymella taxa cause gummy stem blight of economically important plants including certain cucurbit crops. Didymella blight initially presents as small, irregularly circular, water-soaked spots. These early lesions may appear as a fading of the fruit color before darkening to shades of gray, brown, or eventually black. As the disease progresses, these spots rapidly enlarge, becoming sunken, often acquiring a firm, leathery texture (Kehinde et al. 2013). Gummy stem blight caused by Di. bryoniae has been reported in many countries including China, Tanzania and the United States (Mao et al. 2020). Furthermore, Di. lycopersici is an opportunistic pathogen that causes cankerous symptoms on tomato stems (Blancard 2012).

Hosts: Didymella taxa have been reported from a wide range of hosts including Acacia in Australia, Araucaria in Italy, Berberis in Netherlands, Camellia in China, Castanea in New Jersey, Citrus in Paraguay, Cucurbita in Italy, Eucalyptus in Australia, Fucus and Mangifera in Brazil, Olea in Iran, Pandanus in Philippines, Prunus and Rosa in Pakistan, Rubus in Australia, Canada and Pakistan, Vitis in Germany, and Zea in China, among many other hosts (Farr & Rossman 2025).

Pathogen biology, disease cycle and epidemiology: The disease cycle typically starts with the survival of Didymella in infected plant debris, seeds, or soil, from which it produces spores that are dispersed by wind or rain splash (Keinath 2008). Upon landing on a susceptible host, the spores germinate and infect through wounds or natural openings, leading to disease symptoms (Babadoost & Zitter 2009, Deb et al. 2020). Under favourable conditions, Didymella rapidly develops and sporulates, leading to secondary infections. Several species release their ascospores by an eruptive mechanism. This increases the chance for spores to spread and propagate over long distances. The bitunicate ascus comprises a sensitive inner and rigid outer wall resulting in an explosion of the latter under wet conditions, which causes the release of the ascospores (Ingold 1971, Sadyś & West 2017).

Morphological-based identification: Didymella taxa have ostiolate or poroid conidiomata that are pycnidial and sub-globose. The conidiogenous cells are generally phialidic, hyaline and ampulliform. The conidia usually display diverse shapes being ellipsoidal to subglobose, oblong to ovoid, or cylindrical. Generally, the conidia are hyaline and aseptate, and become pigmented when mature. The ascomata are mostly pseudothecial, globose to sub-globose and ostiolate. The bitunicate asci are cylindrical, clavate or saccate, usually containing 8 spores. The ascospores are hyaline or brown and can be ellipsoidal to cymbiform, and uni- or multi-septate (Gruyter et al. 2009, Aveskamp et al. 2010, Zhang et al. 2012b, Chen et al. 2015).

Fig. 5. Didymella sp. a Disease symptom of D. gei on Geum sp. b Disease symptom of D. qilianensis on Rheum officinale. c Ascus. d-e Ascospores. f Chlamydospores. Scale bars: c-e = 10 µm, f = 5 µm. This picture is copyright of Qian Chen (Chen et al. 2015, 2022).

Molecular-based identification: Studies focusing on the revision of Didymellaceae were based on multi-gene phylogenetic analyses of the concatenated ITS, LSU, tub2, and rpb2 sequences (Gruyter et al. 2009, Aveskamp et al. 2010, Chen et al. 2015, Jayasiri et al. 2017, Phukhamsakda et al. 2020). We constructed our phylogenetic tree in accordance with previous studies. Phylogenetic analysis based on the ITS, LSU and tub2 sequences does not completely resolve the taxonomic placement of certain taxa in Didymellaceae. This explains the rationale for including rpb2 in subsequent phylogenetic analyses (Aveskamp et al. 2010, Chen et al. 2015). This study provides an updated phylogeny of Didymella based on combined ITS, LSU, rpb2 and tub2 sequence data (Fig. 6, Table 4).

Recommended genetic markers (genus level): LSU

Recommended genetic markers (species level): ITS, LSU, rpb2 and tub2

Accepted number of species: 99

References: Widmer et al. (2002), Ozkilinc et al. (2011), Woudenberg et al. (2012), Khan et al. (2013), Gorny et al. (2016), Pearce et al. (2016), Havenga et al. (2019), Lee et al. (2019), Owati et al. (2019), Ren et al. (2019), Armstrong-Cho et al. (2020), Guo et al. (2020), Scarpari et al. (2020), Jiang et al. (2021), Šišić et al. (2021), Wang et al. (2021), Indermaur et al. (2022), Ma et al. (2022) (pathogenicity data)

129. Leptosphaerulina McAlpine, Fungus Diseases of stone-fruit trees in Australia: 103 (1902)

Type species: Leptosphaerulina australis McAlpine

Classification: Dothideomycetes, Pleosporales, Didymellaceae

Background: Leptosphaerulina was established by McAlpine (1902) with L. australis as the type species, which was isolated from the leaves of Prunus armeniaca (apricot). Initially, Leptosphaerulina was accommodated in Pseudosphaeriaceae based on morphological features (Luttrell 1955, Graham & Luttrell 1961, Barr 1982). Later, it was placed in Pleosporaceae (Eriksson 1993, 2006, Kirk et al. 2001). Eventually, its placement was confirmed in Didymellaceae based on molecular data (Aveskamp et al. 2010, Zhang et al. 2012b, Hyde et al. 2013, Hongsanan et al. 2020a, b). Didymellaceae is a species-rich family in Pleosporales that includes taxa from a wide range of hosts from different ecosystems (Chen et al. 2017, Thambugala et al. 2018).

Distribution: Leptosphaerulina taxa have cosmopolitan distributions and have been recorded from both temperate and tropical countries, including Canada, China, Colombia, Georgia, India, Indonesia, Japan, Kenya, the Netherlands, 2013, Chen et al. 2017, Farr & Rossman 2025).

Fig. 6. Maximum likelihood tree of Didymella species based on the concatenated ITS, LSU, rpb2 and tub2 sequence data. The best scoring RAxML tree had a final likelihood value of -16957.247. The tree was rooted with Epicoccum camelliae (CGMCC 3.18343) and E. latusicollum (CGMCC 3.18346). Estimated base frequencies were as follows: A = 0.250, C = 0.250, G = 0.250, T = 0.250; substitution rates AC = 1.45908, AG = 6.33861, AT = 1.45908, CG = 1.00000, CT = 11.74000, GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Type strains are in bold. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A. Species confirmed with pathogenicity studies are marked with #.

Disease symptoms: Major diseases caused by Leptosphaerulina species are leaf spots and blights, as well as scorch spots (Fig. 7). Leaf blight caused by Leptosphaerulina spp. is common in humid and damp regions, primarily on grasses such as Agrostis stolonifera (creeping bentgrass), Poa pratensis (Kentucky bluegrass), and Lolium perenne (perennial ryegrass) (Shurtleff et al. 1987, Smiley et al. 1992). The symptoms displayed during the course of leaf blight disease formation are tip dieback often with yellow to brown lesions extending down to the leaf sheaths. Sometimes, the entire leaf or sheath may develop water-soaked spots and quickly become chlorotic (Couch 1962, Smiley et al. 1992).

Hosts: Leptosphaerulina taxa have been reported worldwide from a wide range of hosts. These include Acacia mearnsii in South Africa, Arachis hypogaea in China, Boerhaavia difusa in India, Brassica spp. in China, Cannabis sativa in India, Crotalaria striata in India, Euphorbia geniculata in India, Ficus retusa in India, Medicago laciniata in Kenya, Morus sp. in Japan, Oryza sativa in India, Terminalia bellerica in India, Trifolium pretense in Georgia, and Zea mays in Malaysia, among several others (Farr & Rossman 2025).

Pathogen biology, disease cycle and epidemiology: Leptosphaerulina species cause leaf spots and scorch diseases of leguminous plants, including alfalfa, peanut, red and white clover, and soybean (Graham & Luttrell 1961, Anahosur & Fazalnoor 1972). The ascospores of Leptosphaerulina are easily dispersed via wind (Graham & Luttrell 1961). The mucilaginous sheath allows them to adhere to leaf surfaces and adsorb water (Tehon & Stout 1928, Furtado & Olive 1971). Following adherence, germ tubes are formed, after which a penetration peg enters the cuticle and epidermal cell walls (Hopskins 1923, Miles 1925, Sundheim & Wilcoxson 1965, Wu & Hanlin 1992). The germ tubes can also enter plant tissues via stomata; however, this process is rather infrequent (Wu & Hanlin 1992).

Morphological-based identification: Leptosphaerulina species have ostiolate, papillate and immersed to erumpent ascomata. The asci are bitunicate and saccate while

Fig. 7. Leptosphaerulina crassiasca on Arachis hypogaea. a Adaxial and abaxial b views of the wedge-shaped scorch symptom. c A typical leaf scorch. d Transverse section of perithecium with asci. e Ostiolum. f Immature ascus. g Mature 8-spored ascus and ascospore with longitudinal and transverse septa (indicated with arrow). Scale bars = 25 µm. This picture is copyright of Dartanha Jose Soares.

ascospores are oblong to cylindrical, usually muriform and hyaline, becoming brown when mature (Graham & Luttrell 1961, Abler et al. 2003, Zhang et al. 2012b, Phookamsak et al. 2013). Some phylogenetically distinct Leptosphaerulina taxa are cryptic (Phookamsak et al. 2013).

Molecular-based identification: Multi-locus phylogenetic analyses of ITS, LSU, tub2 and rpb2 sequences are recommended for species delineation in Leptosphaerulina (Chen et al. 2015, Valenzuela-Lopez et al. 2018, Hou et al. 2020). The ITS and LSU markers alone do not provide sufficient phylogenetic resolution (Liang et al. 2021). Phylogenetic analyses of rpb2 sequences only can yield a phylogenetic tree with a similar topology as that of the concatenated dataset (ITS, LSU, tub2, rpb2 and tef) (Liang et al. 2021). Therefore, rpb2 is the best DNA marker for species delimitation in Leptosphaerulina (Liang et al. 2021), however, several species lack rpb2 sequences. Leptosphaerulina crassiasca (XWS02F460) was excluded from the molecular analyses as only the ITS region is available for this taxon. This study provides an updated phylogeny of Leptosphaerulina based on combined ITS, LSU, rpb2, tub2 and tef sequence data (Fig. 8, Table 5).

Recommended genetic markers (genus level): LSU

Recommended genetic markers (species level): ITS, LSU, rpb2, tub2, and tef

Accepted number of species: 18

References: Liu et al. (2019), Victoria et al. (2020), Zhang & Li (2021), Sheng et al. (2022) (species with pathogenicity data)

130. Macrophomina Petr., Annls Mycol. 21(3/4): 314 (1923)

Type species: Macrophomina phaseolina (Tassi) Goid.

Classification: Dothideomycetes, Botryosphaeriales, Botryosphaeriaceae

Background: Macrophomina was introduced by Petrak (1923) with M. philippinensis as the type species. However, the correct name for the type species was followed by a lot of confusion, especially with M. phaseoli and M. phaseolina being used interchangeably during the first half of the 20th century. It was only in the late 1970s that M. phaseolina was widely recognized as the correct name and M. philippinensis was synonymised with M. phaseolina (Ashby 1927, Holliday & Punithalingam 1970, Babu et al. 2010). Macrophomina was widely accepted as a monotypic genus, despite several attempts to propose distinct sub-specific ranks based mainly on host-specificity (Phillips et al. 2013, Sarr et al. 2014).

Fig. 8. Maximum likelihood tree of Leptosphaerulina species based on the concatenated ITS, LSU, rpb2, tub2 and tef sequence data. The best scoring RAxML tree had a final likelihood value of -9591.838. The tree was rooted with Epicoccum plurivorum (CBS 558.81). Estimated base frequencies were as follows: A = A: 0.250, C = A: 0.250, G = A: 0.250, T = A: 0.250; substitution rates AC = 1.00000, AG = 3.47926, AT = 1.00000, CG = 1.00000, CT = 9.30598, GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Type strains are in bold. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A. Species confirmed with pathogenicity studies are marked with #.

Macrophomina is a diverse, soil- or seed-borne plant pathogen occurring worldwide. Macrophomina species commonly cause diseases such as stem and root rot, charcoal rot, seedling blight and stem blight (Agarwal 2010, Ndiaye et al. 2015, Marquez et al. 2021). Macrophomina has been reported on approximately 600 cultivated and wild plant species, belonging to over 100 families in more than 80 countries (Khan 2007, Agarwal 2010, Ndiaye et al. 2015, Zhao et al. 2019, Farr & Rossman 2025).

Distribution: worldwide, but mostly in tropical, subtropical and warm-temperate areas (Cohen et al. 2022a, Farr & Rossman 2025)

Disease symptoms: Macrophomina species generally cause charcoal-rot or ashy stem blight in various host species (Fig. 9). Macrophomina symptoms may vary depending on host and weather conditions. The main disease symptoms are dry or wet dark rot of the root and lower stem (Agarwal 2010, Poudel et al. 2021). Roots and soft-tissue plants, such as melon and strawberry, exhibit early tissue rot due to cell-wall-degrading enzymes, whereas plants with lignified stems, such as cotton, sesame, soybean, and castor bean, usually start exhibiting yellowing and wilt symptoms without visible external stem rot, due to the disruption of water and nutrient flow to aerial parts. Lower stem and taproot cortical tissue become slough. Under heat or drought stress, the symptoms are exacerbated usually inducing premature death of branches or of the whole plant. Symptoms occasionally remain mild until plants start flowering or fruiting, when, under favourable conditions, the plant can quickly die (Dhingra & Sinclair 1978, Cohen et al. 2022b). Macrophomina exhibits a strong capacity for rapid colonization of host tissues, eventually leading to their death in a short timeframe. During colonization, it forms microsclerotia and pycnidia on hosts (Dhingra & Sinclair 1978, Khan 2007, Su et al. 2021). On seedling stage, wet root rot followed by damping-off predominates. Aboveground symptoms on adult plants usually appear as foliar yellowing and wilt of either some branches or the entire plant. Eventually symptoms progress to light-brown, grey, or black lesions, due to fungal colonization of plant tissues and production of numerous microsclerotia, characterizing the classical charcoal rot/ashy stem symptoms. At this stage, pycnidia can sometimes, be found on aboveground colonized tissues. As the fungus colonization spreads up the stem during the seasons, it may eventually reach the pods, capsules or grains (Gupta et al. 2012). Underground organs, such as tubers and rhizomes, can also be partially or completely colonized (Mello et al. 2021).

Hosts: Macrophomina affects a wide range of hosts, including field crops, vegetables, ornamentals, shrubs and trees (Poudel et al. 2021, Cohen et al. 2022a, Bhunjun et al. 2024, Farr & Rossman 2025). Macrophomina species have been recorded worldwide on a broad range of economically important hosts including soybean (Glycine max), maize (Zea mays), common bean (Phaseolus vulgaris), black gram (Vigna mungo), chickpea (Cicer arietinum), cotton (Gossypium spp.), pigeon pea (Cajanus cajan), sorghum (Sorghum bicolor), sunflower (Helianthus annuus), and sesame (Sesamum indicum) (Dhingra & Sinclair 1978, Root 1986, Reyes-Franco et al. 2006, Khan 2007, Agarwal 2010, Ndiaye et al. 2015, Santos et al. 2020, Poudel et al. 2021, Cohen et al. 2022b). Macrophomina phaseolina is the most common and widespread species causing stem and root rot, charcoal rot and seedling blight in hundreds of plant species (Islam et al. 2012, Farr & Rossman 2024). Macrophomina pseudophaseolina has been recorded on cowpea (Vigna unguiculata), peanut (Arachis hypogaea), sorrel (Hibiscus sabdarifa), okra (Abelmoschus esculentus), cotton (G. hirsutum), castor bean (Ricinus communis), cassava (Manihot esculenta), sweet potato (Ipomoea batatas), lentil (Lens culinaris), common bean, and few other weeds (Sarr et al. 2014, Ndiaye et al. 2015, Brito et al. 2019, Machado et al. 2019, Mello et al. 2021, Viejobueno et al. 2022, Kouadri et al. 2023). Macrophomina euphorbiicola was originally reported on castor bean and bellyache bush (Jatropha gossypifolia), and later on Stevia rebaudiana, Amaranthus retroflexus, and Convolvulus arvensis (Machado et al. 2019, Mello et al. 2021, Sanabria-Velazquez et al. 2022, Moslemi et al. 2023). Macrophomina tecta was first reported on sorghum and mung bean (Vigna radiata), and later on corn (Poudel et al. 2021, Viejobueno et al. 2022). On the other hand, M. vaccinii is known only through its original reports on blueberries (Vaccinium spp.) (Zhao et al. 2019).

Pathogen biology, disease cycle and epidemiology: Microsclerotia are the primary inoculum source of Macrophomina species (Fig. 10). Pycnidia and conidia production has been occasionally observed in different host plants, however, their role in disease development and epidemiology is yet to be understood (Dhingra & Sinclair 1978, Romero Luna et al. 2017). Early infection stages demonstrate that Macrophomina should be regarded as hemibiotrophic, instead of a truly necrotrophic fungus (Shirai & Eulgem 2023). Although Macrophomina is considered a poor soil competitor, its microsclerotia in infected roots and plant residues play an

Fig. 9. Macrophomina phaseolina on various hosts. a Castor bean plant showing stem blight and branch death. b-c Roots of castor bean with extensive black lesions. d Longitudinal section of castor bean stem showing profuse microsclerotia formation. e Castor bean stem with presence of pycnidia and conidia cirrus. f Wilted and dead sesame plants. g-h Profuse formation of microsclerotia externally and internally (h) on sesame stem. i Longitudinal section of soybean stem with microsclerotia. j-k Corn stem with microsclerotia. l Microsclerotia formed on sesame root tissue. m Microsclerotium. n Transversal section of a pycnidium with conidia. o-q Immature M. pseudophaseolina conidium with apical mucous appendage and (p-q) mature pigmented conidia. Scale bars: l = 100 µm, m-q = 10 µm. This picture is copyright of Dartanha Jose Soares (except i by HK da Silva, j-k by R Veras and o-q by AR Machado).

Fig. 10. Charcoal rot lifecycle of Macrophomina phaseolina. Redrawn from Smith et al. (2014).

important role in its survival (Dhingra & Sinclair 1978, Gupta et al. 2012, Romero Luna et al. 2017). Depending on the environmental conditions and on the constitution of the host tissues, the microsclerotia can survive for several years in the soil (Gupta et al. 2012). Temperatures between 25 and 35 °C are favourable to microsclerotia germination and disease development. The microsclerotia can infect seedlings and roots of both young and mature plants. During colonization, Macrophomina produces enzymes and toxins that degrade root tissue, and with the onset of infection, the root and stem tissue is colonized within 2-3 weeks, with abundant microsclerotia being produced in the colonized tissues (Romero Luna et a. 2017). Macrophomina species can survive not only in soil and crop debris for long periods (mostly through microsclerotia), but also in asymptomatic seeds as mycelium and in symptomatic seeds as microsclerotia (Dhingra & Sinclair, 1978, Machado et al. 2019).

Morphological-based identification: Macrophomina is readily recognized based on the presence of microsclerotia (Petrak 1923). Conidiomata are pycnidial, globose, thick-walled, variable in size, dark brown, mostly immersed to erumpent. Ostiole is central, unilocular and papillate. Conidiophores are absent. Conidiogenous cells are hyaline, lageniform to cylindrical, proliferating percurrently near the apex and covered with a mucous layer when young. Conidia are hyaline, ellipsoid to obovoid, truncate at base, rounded at apex and covered with a thin outer mucous sheath when young (type C according to Nag Raj (1993)). Mature conidia become septate, medium to dark-brown, without mucoid appendages. Sclerotia are composed of dark-brown to black thick-walled cells that are irregular in shape and variable in size (Sutton, 1980, Saar et al. 2014, Zhao et al. 2019, Machado et al. 2019, Poudel et al. 2021, Moslemi et al. 2023).

Molecular-based identification: The identification of Macrophomina species based on morphology is difficult due to the largely overlapping of morphological features between the species (Machado et al. 2019). However, only the ITS region should not be used due to its low resolution to distinguish Macrophomina species (Machado et al. 2019, Poudel et al. 2021). The concatenation of multiple loci has been widely used for recent taxonomic studies of Macrophomina species. Most studies were based on act, cal, ITS, tef, and tub2 loci (Sarr et al. 2014, Machado et al. 2019, Zhao et al. 2019, Poudel et al. 2021). This study provides an updated phylogeny of Macrophomina based on combined ITS, tef, tub2, act and cal sequence data (Fig. 11, Table 6).

Recommended genetic markers (genus level): ITS

Recommended genetic markers (species level): ITS, tef, tub2, act and cal

Accepted number of species: five

References: Huda-Shakirah et al. (2019), Zhao et al. (2019), Machado et al. (2019), Nouri et al. (2020), Poudel et al. (2021) (morphology, phylogeny and accepted species numbers).

Fig. 11. Maximum likelihood tree of Macrophomina species based on the concatenated ITS, tef, tub2, act and cal sequence data. The best scoring RAxML tree had a final likelihood value of -4916.432. The tree was rooted with Melanops tulasnei (CBS 116805 and CBS 116806). Estimated base frequencies were as follows: A = 0.202, C = 0.317, G = 0.255, T = 0.225; substitution rates AC = 1.00000, AG = 3.15050, AT = 1.00000, CG = 1.00000, CT = 5.38755, GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Type strains are in bold. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A. Species confirmed with pathogenicity studies are marked with #.

131. Melampsora Castagne, Observ. Uréd. 2: 18 (1843)

Type species: Melampsora euphorbiae (Ficinus & C. Schub.) Castagne

Classification: Pucciniomycetes, Pucciniales, Melampsoraceae

Background: Melampsora is a genus of plant pathogenic rust fungi belonging to Basidiomycota. The genus was established by Castagne (1843) with Me. euphorbiae as the type species causing rust on Euphorbia sp. Melampsora is characterised by the formation of sessile and unicellular teliospores which are united laterally and form crusts or columns on the host. The genus is an obligate biotrophic parasite that contains about 100 species with most of them being heteroecious. Members of this genus infect a variety of host plants including agricultural and non-agricultural crops. Poplar leaf rusts caused by various species of Melampsora including Me. larcici-populina, Me. medusa and Me. occidentalis are important tree diseases. Linin flax rust, Melampsora lini (Ehrenb.) Lév., is the most important pathogen of cultivated Linum usitatissimum.

Distribution: Algeria, Argentina, Canada, China, Colorado, France, Germany, Greenland, India, Iran, Japan, Mongolia, Morocco, Mozambique, Nepal, New Zealand, Pakistan, Russia, Shanxi, Siberia, Spain, Sri Lanka, Tibet, USA, Uzbekistan and Venezuela (Farr & Rossman 2025).

Hosts: Abies, Acalypha, Aleurites, Chelidonium, Chosenia, Corydalis, Euphorbia, Hypericum, Hypericum, Larix, Lobelia, Mercurialis, Pinus, Piscaria, Polygala, Populus, Ribes, Salix, Saxifraga, Serissa, Stellera, Stereospermum, Vernonia and Wikstroemia (Farr & Rossman 2025).

Disease symptoms: Melampsora species produce yellow leaf spots, which eventually become necrotic (Fig. 12). The common rust symptoms appear as orange spore-producing structures (uredinia) on the underside of host leaves. On alternate hosts (conifers), after sporulation Melampsora species cause shrinkage and finally death of infected needles. Yellow aecia of these rust fungi are sometimes visible on cones of the alternate hosts (Cummins1959, Cummins & Hiratsuka 2003).

Fig. 12. Melampsora on Euphorbia sp. a Infection of rust on leaves. b Uredinia. c Telia (tiny black dots). This picture is copyright of Ajay Kumar Gautam, Shubhi Avasthi and Rajnish Kumar Verma.

Pathogen biology, disease cycle and epidemiology: Melampsora species enter the host surface either through stomatal openings or by direct penetration through epidermal cells. After entry, the pathogen forms primary and secondary mycelium intercellularly for the establishment and giving rise to haustoria. Melampsora spp. often form a crust-like brown membrane compacted into flat, dark-coloured groups of single-celled sessile teliospores below the cuticle or epidermis of the host plant. Melampsora species are often heteroecious requiring two unrelated host plants or complete their life cycle on the one host (autoecious). They may produce five different spore types (macrocyclic) which reflects the complexity of these fungi. In several Melampsora species, however, only uredinial and telial stages have been described. Over half of the described species of Melampsora are known on Salix (willow) and Populus (poplar). None of the species of Salix in the world are entirely free of rust infection. For heteroecious species, the alternate hosts include conifers (Tsuga, Abies, Larix and Ribes spp.) (Sinclair 1987, Pei et al. 1993).

Morphological-based identification: The identification of Melampsora rust fungi is mainly based on their morphology, alternate hosts, and telial host range. The identification is also based on the position of uredinia and telia on leaves of host plants, the thickness of spore walls along with morphology of uredinial paraphyses. Melampsora produces intercellular, septate, branched, dikaryotic, and subepidermal mycelium within the parenchymatous tissues of the host. Spermogonia are of Group I (type 3 subcuticular; or type 2, subepidermal), flask-shaped, pale yellow, containing paraphyses and minute, oval to globose spermatia. Aecia are orange-yellow, scattered on the undersurface of the leaves or sometimes on the stem, subepidermal, erumpent, and of Caeoma–type (some species have peridial cells adherent to the host epidermis). Aeciospores are polygonal in shape, echinulate, or verrucose with rod-like columns or blocks. Uredinia are subepidermal, erumpent and Uredo-type occur on both surfaces of leaves. They are bright yellow or orange initially and fade to nearly colourless with the progress of the disease. Capitate paraphyses are found abundantly in uredinia. Urediniospores are orange in colour, echinulate, with colourless walls, borne singly on pedicels and intermixed with capitate paraphyses. The pores of these spores may be scattered or bizonate, and obscure. Telia are reddish brown, elongated, subepidermal or relatively subcuticular, and remain covered, forming adherent crusts of 1 spore deep on the stem. The 1-celled sessile teliospores have a single germ pore and a brown wall. Melampsora species form external basidia (Cummins & Hiratsuka 2003, Ebinghaus et al. 2020).

Molecular-based identification: Maier et al. (2003) analysed LSU sequence data of Melampsora and placed it in a basal position of the Pucciniales (Uredinales). This DNA sequence-based systematic placement of this genus also supported its proposal to be ancestral (Durrieu 1980). However, based on SSU sequence data, Wingfield et al. (2004) proposed Racospermyces (Endoraecium) in the basal-most position and Melampsora in a derived lineage. Pei et al. (2005) generated LSU and ITS sequences of 11 species of Melampsora from Salix sp. and Populus sp. and Aime et al. (2006) examined two nuclear rDNA genes (18S and 28S) from field or herbarium specimens and DNA sequences from GenBank. They included two Melampsora species (Me. epitea Thüm. and Me. euphorbiae Castagne) in their study along with other rust fungi. Similarly, based on morphological characteristics and the ITS sequence data, a new species of Melampsora (Me. iranica) was reported on Salix elbursensis in the northwest of Iran (Damadi et al. 2011). Using ITS-2 and large ribosomal subunit RNA gene (LSU) regions, Busby et al. (2012) identified Me. columbiana on two species of Populus from North America. The molecular analyses using rDNA sequence information of two heteroecious rust species, Me. chelidonii-pierotii and Me. yezoensis, which parasitize Salix spp. and members of Papaveraceae revealed their phylogenetic similarity, although they have different survival strategies (Shinyama and Yamaoka 2012). Fifteen Melampsora spp. were examined phylogenetically using six nuclear and mitochondrial loci by Vialle et al. (2013). They observed that only three lineages fitted exactly defined morphological species, whereas, 12 were concordant with host specialization. Zhao et al. (2015a, b) studied Melampsora species on willows in China, and two new species and one new record of Melampsora were identified. The internal transcribed spacer (ITS1 and ITS2) region and LSU sequences were used in molecular phylogeny along with morphological characteristics to identify these fungi. In a similar study of Melampsora species on willows, one new species (Melampsora salicis-reinii) and two new records (Me. ribesii-viminalis and Me. ribesii-purpureae) were identified from Japan based on morphological and molecular phylogenetic studies using ITS region (Zhao et al. 2015c). Ali et al. (2016) re-examined Melampsora euphorbiae, Me. euphorbiae-gerardianae and Me. helioscopiae reported previously on Euphorbia helioscopia in Pakistan and described a new species, Me. pakistanica. Yun et al. (2016) reported Me. yezoensis as a new rust species on Salix koreensis based on morphology and molecular data of the ITS region. Based on analysis of sequences of ITS region and LSU (D1/D2) along with morphological characters, Zheng et al. (2019) reported a North American poplar leaf rust species, Me. medusae, in China. Aime & McTaggart (2020) included two species of Melampsora (Me. euphorbiae and Me. laricis-populina) in their study to provide a higher-rank classification for Pucciniales. Based on a combined LSU and ITS dataset for 25 rust genera, including 12 species of Melampsora, a phylogenetic analysis of Indian Pucciniales was carried out by Gautam et al. (2021) which provided a better understanding of their phylogeny and evolution. This study provides an updated phylogeny of Melampsora based on combined LSU and ITS sequence data (Fig. 13, Table 7).

Recommended genetic marker (genus level): ITS, LSU

Recommended genetic marker (species level): ITS, LSU

Accepted number of species: 100

References: Cummins (1959), Sinclair (1987), Pei et al. (1993), Cummins & Hiratsuka (2003), Pei et al. (2005), Aime (2006), Aime et al. (2006), Feau et al. (2009), Damadi et al. (2011), Busby et al. (2012), Shinyama & Yamaoka (2012), Vialle et al. (2013), Padamsee & McKenzie (2014), Toome & Aime (2015), Ali et al. (2016), Giordano et al. (2019), Aime & McTaggart (2020) (morphology and phylogeny)

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A.

Fig. 13. Maximum likelihood tree of Melampsora species based on the concatenated LSU and ITS sequence data. The best scoring RAxML tree had a final likelihood value of -5783.292563. The tree was rooted to Naohidemyces vaccini. Estimated base frequencies were as follows: A =0.300219, C =0.176250, G =0.227689, and T =0.295842; substitution rates AC =1.261606, AG =1.929654, AT =0.708137, CG =0.305156, CT =2.230781, and GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Type strains are in bold. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

132. Myrothecium Tode, Fungi Mecklenburgenses Selecti 1: 25 (1790)

Type species: Myrothecium inundatum Tode.

Classification: Sordariomycetes, Hypocreales, Stachybotryaceae

Background: Myrothecium was introduced by Tode (1790). Myrothecium sensu lato encompasses approximately 18 genera, all grouped within Stachybotryaceae (Lombard et al. 2016, Liang et al. 2019). Many species are recognized for their significant biological activities (Brian et al 1948, Lee et al. 2008, Ruma et al. 2015). Although several Myrothecium species are considered weak or opportunistic plant pathogens, species such as Albifimbria verrucaria (synonym Myrothecium verrucaria) and Paramyrothecium roridum (synonym My. roridum) have been studied more in depth due to their aggressive pathogenicity on a wide range of plant hosts (Han et al. 2014, Ben et al. 2016, Weaver et al. 2021). Myrothecium have also been reported as saprobes and endophytes from different substrates and ecosystems, including soil, decaying plant matter, and living plant tissues (Ahrazem et al. 2000, Quezado Duval et al. 2010, Chen et al. 2016). Despite numerous studies, the taxonomy of many taxa within the genus remains unresolved, and further molecular and morphological studies are necessary to clarify their classification.

Distribution: worldwide (Farr & Rossman 2025)

Disease symptoms: Myrothecium species are opportunistic fungal pathogens that infect a wide range of plants, causing leaf spots, stem cankers, fruit rots, and seedling damping-off (Maji 2013, Chen et al. 2016, Fujinawa et al. 2016, Lombard et al. 2016). Infections usually appear as small, circular to irregular brown or tan necrotic spots with dark, well-defined margins on leaves. These lesions may expand, coalesce, and cause extensive blight (Lombard et al. 2016). Myrothecium can cause elongated, sunken, dark lesions or cankers on stems and petioles that may crack or become soft and water-soaked. Seedlings infected by Myrothecium spp. may suffer from pre- or post-emergence damping-off, showing basal stem rot and eventual seedling collapse (Maji 2013, Chen et al. 2016, Fujinawa et al. 2016).

Hosts: Chrysanthemum spp., cotton (Gossypium hirsutum), cucumber (Cucumis sativus), gerbera (Gerbera jamesonii), pepper (Capsicum spp.), poinsettia (Euphorbia pulcherrima), soybean (Glycine max), tomato (Solanum lycopersicum), and watermelon (Citrullus lanatus) (Farr & Rossman 2025).

Pathogen biology, disease cycle and epidemiology: Myrothecium thrives in warm, humid environments and has a broad host range, infecting vegetables, ornamentals, and field crops (Maji 2013, Chen et al. 2016, Fujinawa et al. 2016, Lombard et al. 2016). The disease cycle begins with the overwintering of Myrothecium in plant debris, soil, or on infected seeds (Nguyen et al. 1973, van Bruggen et al. 2016). Under favourable environmental conditions, particularly high humidity and warm temperatures, conidia germinate and infect host plants. Initial infection is often localized but can spread rapidly in moist environments. Myrothecium penetrates through wounds or weakened tissue, leading to leaf spots, stem lesions, and in severe cases, plant wilting and dieback. Secondary spread occurs via water splash, wind-driven rain, and mechanical transmission (Nguyen et al. 1973, van Bruggen et al. 2016).

Morphological-based identification: The generic concept of Myrothecium has undergone several emendations over time (Link 1809, von Höhnel 1905, Pidoplichko & Kirilenko 1971, Lombard et al. 2016). Myrothecium sensu stricto includes species with cup-shaped sporodochia, which are often covered with slimy masses of green to black conidia (Tode 1790, Chen et al. 2016, Lombard et al. 2016, Fig. 14). These sporodochia may be scattered or aggregated. The conidiophores are short and often branched, bearing phialidic conidiogenous cells that produce conidia in a slimy matrix. The conidia are usually cylindrical to ellipsoidal, smooth or roughened, and vary from hyaline to dark green or black. These morphological traits were previously used to distinguish Myrothecium from morphologically similar genera (Link 1809, von Höhnel 1905, Pidoplichko & Kirilenko 1971). However, due to overlapping morphology, molecular data are needed for species identification.

Molecular-based identification: There have been relatively few comprehensive phylogenetic studies focused on Myrothecium due to the lack of multi-loci data for a number of species (Ruiz et al. 2008, Alves et al. 2010, Jiang et al. 2014, 2017, Wu et al. 2014b). This underscores the need for extensive molecular studies incorporating broader taxon sampling, to clarify the evolutionary relationships and classification within this genus. Decock et al. (2008) used ITS sequence data to distinguish Septomyrothecium from Myrothecium, and their results indicated that Myrothecium is paraphyletic. Chen et al. (2015) re-evaluated the phylogeny of Myrothecium based on ITS and tef sequences, suggesting the polyphyly of Myrothecium within Stachybotryaceae. Lombard et al. (2016) resolved Myrothecium sensu lato to 18 genera and confirmed the placement of two species in Myrothecium and excluded 12 species previously identified as Myrothecium. This study provides an updated phylogeny of Myrothecium and related genera in Stachybotryaceae based on multi-loci sequence data (ITS, LSU, rpb2, tef, tub2 and cal) (Fig. 15, Table 8). We included Myrothecium species that have multi-locus data to determine their phylogenetic placement. We propose the new combinations Digitiseta chiangmaiensis and Myxospora thailandica (Fig. 15 and S1).

Digitiseta chiangmaiensis (D.Q. Dai & K.D. Hyde) Bhunjun, Phukhams. & K.D. Hyde, comb. nov.

Index Fungorum number: IF905055; Facesoffungi number: FoF 19467

Synonym: Myrothecium chiangmaiense D.Q. Dai & K.D. Hyde, in Dai et al., Fungal Diversity 82: 49 (2016)

Description: See Dai et al. (2017)

Notes: Digitiseta chiangmaiensis was described as Myrothecium chiangmaiense from dead culms of bamboo in Thailand (Dai et al. 2017). In the analyses of combined ITS, LSU, rpb2, tef, tub2 and cal sequence data of Stachybotryaceae, the strain (MFLUCC 11–0506) clustered with Digitiseta multidigitata (MUCL 41187). Therefore, we transfer Myrothecium chiangmaiense to Digitiseta (Fig. 15 and S1).

Myxospora thailandica (D.Q. Dai & K.D. Hyde) Bhunjun, Phukhams. & K.D. Hyde, comb. nov.

Index Fungorum number: IF905056; Facesoffungi number: FoF 19466

Fig. 14. Myrothecium sp. a-b Conidiomata. c Sporodochia. d-e Conidiogenous cells. f Conidia. Scale bar: c = 100 µm, d-f = 10 µm. This picture is copyright of Pedro Crous (Lombard et al. 2016).

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A.

Synonym: Myrothecium thailandicum D.Q. Dai & K.D. Hyde, in Dai et al., Fungal Diversity 82: 49 (2016)

Description: See Dai et al. (2017)

Notes: Myxospora thailandica was described as Myrothecium thailandicum from dead culms of bamboo in Thailand (Dai et al. 2017). In the analyses of combined ITS, LSU, rpb2, tef, tub2 and cal sequence data of Stachybotryaceae, the strain (MFLUCC 11-0395) clustered with Myxospora crassiseta (CBS 731.83). Therefore, we transfer Myrothecium thailandicum to Myxospora (Fig. 15 and S1).

Recommended genetic markers (genus level): ITS

Recommended genetic markers (species level): ITS, LSU, rpb2, tef, tub2 and cal

Accepted number of species: two

References: Crous et al (2014), Fujinawa et al. (2016), Lombard et al. (2016) (morphology and phylogeny)

Fig. 15. Maximum likelihood tree of Myrothecium and related genera in Stachybotryaceae based on the concatenated ITS, LSU, rpb2, tef, tub2 and cal sequence data. The best scoring RAxML tree had a final likelihood value of -39144.227. The tree was rooted with Fusarium sambucinum (CBS 146.95 and CBS 136.24). Estimated base frequencies were as follows: A = 0.237, C = 0.272, G = 0.269, T = 0.222, with substitution rates AC = 1.24675, AG = 2.95299, AT = 1.24675, CG = 1.00000, CT = 5.51631, GT = 1.000000. ML support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Type strains are in bold. The new combinations are in blue. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

133. Neopseudocercosporella Videira & Crous, Stud. Mycol. 83:80 (2016)

Type species: Neopseudocercosporella capsellae (Ellis & Everh.) Videira & Crous

Classification: Dothideomycetes, Mycosphaerellales, Mycosphaerellaceae

Background: Neopseudocercosporella is an important pathogen that affects economic crops worldwide. The genus was established by Videira et al. (2016). Both species Neopseudocercosporella brassicae and N. capsellae cause white leaf spot disease in plants in the family Brassicaceae (Videira et al. 2016, Gunasinghe et al. 2020, 2021). Videira et al. (2016) suggested that the two species could be synonymous despite having distinct disease symptoms, ascospore morphology and culture characteristics.

Distribution: worldwide (Farr & Rossman 2025), however, the disease is more destructive in Subtropical, Mediterranean, or Temperate climate regions with cool and wet climates

Disease symptoms: Neopseudocercosporella species are generally foliar pathogens that produce leaf lesions with necrosis or chlorosis, but can also produce stem, and pod lesions under favourable weather conditions (Petrie & Vanterpool 1978, Fig. 16). Neopseudocercosporella capsellae initially causes numerous brown elongated spots on mature lower leaves that later coalesce to form large irregularly shaped, white or pale beige lesions on Brassica napus (rapeseed) (Gunasinghe et al. 2020). As the crop matures, P. capsellae spreads to stems and pods to develop grey to black lesions. These lesions can expand rapidly to cover large areas with ash-depressed centres (Carmody 2017) or purple to grey-speckled areas (Petrie & Vanterpool 1978). The leaf disease symptoms can vary among other brassica hosts depending on the degree of waxiness of the leaf surface and may not resemble the white leaf spot lesions on rapeseed (Crossan 1954). For example, the disease symptoms can be round, semi-transparent larger lesions with brownish-grey centres with well-defined brown or tan margins on turnip and wild mustard and ashy black centred, rectangular or rounded lesions with well-defined margins on cabbage (Miller & McWhorter 1948).

Hosts: Neopseudocercosporella capsellae infects a range of cultivated crucifers including Brassica napus and B. juncea (Boerema & Verhoeven 1977, Gudelj et al. 2004, Murtza et al. 2021, Farr & Rossman 2025) with Chinese cabbage (B. rapa sub sp. pekinensis) and cauliflower (B. oleracea) recorded as common hosts (Lancaster 2006). Neopseudocercosporella species have also been isolated from the leaf lesions of a number of different ‘wild’ or ‘weedy’ crucifers such as Capsella bursa-pastoris (shepherd’s purse), Malcolmia africana (African mustard) and Raphanus raphanistrum (white charlock) (Crossan 1954, Deighton 1973, Petrie & Vanterpool 1978, Föller & Paul 2002, Lancaster 2006).

Pathogen biology, disease cycle and epidemiology: Neopseudocercosporella is capable of completing its life cycle in an asexual or sexual stage (Petrie & Vanterpool 1978, Inman et al. 1991, Barbetti 2000, Fig. 17). In the sexual stage, the ascomata release ascospores which initiate the new disease cycle (Inman et al. 1991). During the asexual stage, the conidia initiate the new disease cycle and asexual resting structures (stromatic knots) are responsible for the survival of the pathogen between crops (Crossan 1954, Perron & Nourani 1990, Fitt et al. 2006).

Morphological-based identification: Neopseudocercosporella differs from the closely related genera Fusoidiella and Filiella which produce fusiform or acicular-filiform conidia, respectively, compared to the subcylindrical conidia of Neopseudocercosporella (Videira et al. 2016). Apseudocercosporella can be distinguished from Neopseudocercosporella by having slightly thickened and darkened conidial hila and conidiogenous cells (Braun 1995). The two Neopseudocercosporella species are distinguishable based on the morphology of ascospores and colony characteristics (Inman et al. 1991).

Molecular-based identification: Videira et al. (2016) established Neopseudocercosporella using LSU, ITS, act, tef, his3, rpb2, cal and tub2 sequences. Although the two species have morphologically distinct features, Videira et al. (2016) suggested that these two species could be synonymous, considering the high similarity in molecular data. However, they are currently considered two distinct species, pending recollection of fresh material. As the members share high sequence similarity, a polyphasic approach is recommended for species-level identification (Videira et al. 2016, 2017). This study provides an updated phylogeny of Neopseudocercosporella based on combined LSU, ITS, act, tef, rpb2, cal and tub2 sequence data (Fig. 18, Table 9).

Recommended genetic markers (genus level): LSU and ITS

Recommended genetic markers (species level): LSU, ITS, act, tef, rpb2, cal and tub2

Accepted number of species: two

References: Gunasinghe et al. (2016), Videira et al. (2016), (2017), Murtza et al. (2021) (morphology and phylogeny)

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A. Species confirmed with pathogenicity studies are marked with #.

Fig. 16. White leaf spot symptoms caused by Neopseudocercosporella capsellae on different hosts: a Leaf lesions on Brassica napus (rapeseed), initially brown, elongated spots that later become white with the production of conidia. b Irregular, semi-transparent larger lesions with brownish-grey centres with well-defined brown or tan margins on Raphanus sativus. c Initially, brown to elongated spots that later become white cantered on Brassica campestris var. chinensis (Chinese cabbage). d Rectangular or rounded with well-defined margins with ashy brown or black centre on Bressica oleracea var. capitata (cabbage). e Stem lesions on Brassica nigra appearing as large black lesions with ash depressed centres. This picture is copyright of Niroshini Gunasinghe.

Fig. 17. Disease cycle of Neopseudocercosporella species. Redrawn from Gunasinghe et al. (2020).

134. Neopestalotiopsis Maharachch., K.D. Hyde & Crous, in Maharachchikumbura et al., Stud. Mycol. 79:135 (2014)

Type species: Neopestalotiopsis protearum (Crous & L. Swart) Maharachch.

Classification: Sordariomycetes, Amphisphaeriales, Sporocadaceae

Background: Neopestalotiopsis was introduced by Maharachchikumbura et al. (2014) to accommodate pestalotiopsis-like taxa that have versicolorous median cells and indistinct conidiophores. Neopestalotiopsis are primarily known as plant pathogens but have also been reported as saprobes and endophytes. Neopestalotiopsis species can cause leaf spots, fruit rot, and other plant diseases, affecting economically important crops such as tea, strawberry, and avocado (Espinoza et al. 2008, González et al. 2012, Chamorro et al. 2016, Sun et al. 2023). The species have also been associated with a wide variety of host plants across tropical and subtropical regions (Rodríguez-Gálvez et al. 2020, Diogo et al. 2021, Zhang et al. 2024).

Distribution: worldwide (Farr & Rossman 2025)

Disease symptoms: Neopestalotiopsis species are known to cause various diseases including canker, blight, dieback, fruit rots and leaf spot (Fig. 19). Neopestalotiopsis taxa have been reported as an important pathogen causing dieback, trunk diseases, leaf blights, leaf spot diseases, fruit rot, postharvest disease and severe defoliation of grapevine

Fig. 18. Maximum likelihood tree of Neopseudocercosporella species based on the concatenated LSU, ITS, act, tef, rpb2, cal and tub2 sequence data. The best scoring RAxML tree had a final likelihood value of -6877.382. The tree was rooted with Neocercospora ammicola (CCTU 1186). Estimated base frequencies were as follows: A = 0.239, C = 0.255, G = 0.287, and T = 0.219; substitution rates AC = 1.65698, AG = 2.88680, AT = 1.65698, CG = 1.00000, CT = 7.29436, and GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

Fig. 19. Disease symptoms of: a Neopestalotiopsis sp. on Smilax sp. b Neopestalotiopsis sp. on Ginkgo biloba. c Neopestalotiopsis rhapidis on Podocarpus macrophyllus. d Neopestalotiopsis sp. on Smilax scobinicaulis. e Neopestalotiopsis sp. on Syzygium samarangense. f Neopestalotiopsis amomi on Amomum villosum. This picture is copyright of Yaru Sun.

cultivars (Jayawardena et al. 2015, 2016, Maharachchikumbura et al. 2017, Sun et al. 2023). As a result, they reduce the yield and quality of grapes, leading to the death of individual plants. Neopestalotiopsis have also been associated with stem blight, flower blight, twig dieback and fruit rots of a wide range of plants worldwide (Akinsanmi et al. 2016, Borrero et al. 2017, Mahapatra et al. 2018, Rodríguez-Gálvez et al. 2020, Zhang et al. 2024). Neopestalotiopsis species cause guava scab (Solarte et al. 2018), leaf spot on sweet potatoes (Maharachchikumbura et al. 2016), crown and root rot of strawberries, as well as canker and dieback on blueberries (Espinoza et al. 2008, González et al. 2012, Chamorro et al. 2016). Neopestalotiopsis also infects the leaves and fruits of strawberries, with the infection initially manifesting as circular, black, and slightly sunken spots that expand outwards on the surface (Ayoubi and Soleimani 2016). In fruit rots, the pathogen initially develops small, circular, black and slightly sunken spots on fruits that enlarge rapidly, becoming sunken and resulting in a soft decay of the fruit flesh (Maharachchikumbura et al. 2013a, b).

Hosts: Neopestalotiopsis have a wide host range including Acacia mollissima, Achras sapota, Acrostichum aureum, Alpinia malaccensis, Annona squamosa, Ardisia crenata, Artocarpus heterophyllus, Bulbophyllum, Calliandra haematocephala, Camellia, Carya illinoinensis, Cinchona, Cissus, Citrus, Cocos nucifera, Crotalaria juncea, Elaeis guineensis, Elettaria cardamomum, Eriobotrya japonica, Eucalyptus, Ficus macrocarpa, Fragaria × ananassa, Garcinia mangostana, Hevea brasiliensis, Ilex chinensis, Ipomoea, Kadsura coccinea, Leucospermum cuneiforme, Ligustrum lucidum, Litsea rotundifolia, Macadamia, Machilus thunbergia, Mangifera indica, Magnolia, Malus, Musa, Nopalea cochenillifera, Neodypsis decaryi, Paeonia suffruticosa, Pandanus, Paphiopedilum micranthum, Phoenix dactylifera, Picea, Pinus brutia, Platanus orientalis, Prunus dulcis, Psidium, Pteridium, Punica granatum, Rosa, Quercus rubra, Rhizophora, Sonneratia alba, Sporobolus elongatus, Syzygium, Taxus chinensis, Telopea, Thuja occidentalis, Vaccinium and Vitis (Farr & Rossman 2025).

Pathogen biology, disease cycle and epidemiology: Neopestalotiopsis species are opportunistic pathogens that primarily infect plants through wounds or natural openings, often under conditions of plant stress or environmental imbalance (Belisário et al. 2020, Diogo et al. 2021). The disease cycle typically begins with conidia landing on the surface of a susceptible host, where they germinate and penetrate plant tissues, leading to symptoms such as leaf spots, fruit rot, stem cankers, or dieback (Belisário et al. 2020). Under favourable conditions, Neopestalotiopsis rapidly colonizes plant tissue, producing new conidia on acervuli that emerge on infected surfaces. These spores are then dispersed by wind, rain splash or insects, facilitating secondary infections throughout the growing season (Diogo et al. 2021). The pathogen can persist in plant debris or soil, serving as a source of primary inoculum in subsequent seasons.

Morphological-based identification: Neopestalotiopsis species can be differentiated from other pestalotioid taxa by using morphology and molecular data (Maharachchikumbura et al. 2014b). Neopestalotiopsis species differ from Pestalotiopsis and Pseudopestalotiopsis in having versicolorous median cells whereas both Pestalotiopsis and Pseudopestalotiopsis have concolourous median cells (Maharachchikumbura et al. 2014b). However, using morphological characters to delimitate species is problematic as the morphological characters are plastic and vary between hosts and environments (Maharachchikumbura et al. 2011, 2016). Therefore, molecular data is important to identify different pestalotioid species (Maharachchikumbura et al. 2016, 2021).

Molecular-based identification: Neopestalotiopsis is closely related to Pestalotiopsis and Pseudopestalotiopsis. Neopestalotiopsis species can be roughly separated from Pestalotiopsis and Pseudopestalotiopsis based on base pair difference in the ITS region (Maharachchikumbura et al. 2014b). It is recommended to use ITS, tub2 and tef genes to provide a better resolution in phylogenetic analyses. This study provides an updated phylogeny of Neopestalotiopsis based on combined ITS, tub2 and tef sequence data (Fig. 20, Table 10).

Recommended genetic marker (genus level): LSU

Recommended genetic markers (species level): ITS, tub2 and tef

Accepted number of species: 112

References: Maharachchikumbura et al. (2012), (2014b), (2016), Jayawardena et al. (2015), (2016), Mahapatra et al. (2018), Rodríguez-Gálvez et al. (2020), Razaghi et al. (2024) (morphology, phylogeny and pathogenicity)

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A. Species confirmed with pathogenicity studies are marked with #.

135. Neosetophoma Gruyter, Aveskamp & Verkley, in Gruyter et al., Mycologia 102(5): 1075 (2010)

Type species: Neosetophoma samararum (Desm.) Gruyter, Aveskamp & Verkley

Classification: Dothideomycetes, Pleosporales, Phaeosphaeriaceae

Background: Neosetophoma was described as an asexual morph with Ne. samararum as the type species (Gruyter et al. 2010). Members have been reported as pathogens, endophytes, or saprobes in a wide range of habitats (Phookamsak et al. 2014, Hernandez-Restrepo et al. 2016, Karunarathna et al. 2017, Wanasinghe et al. 2018). Neosetophoma is characterized by globose to irregular conidiomata, with papillate ostioles, and yellowish conidia that are attenuated at one end (De Gruyter et al. 2010, Liu et al. 2015).

Distribution: worldwide (Farr & Rossman 2025)

Disease symptoms: Neosetophoma species are known to cause a variety of plant disease symptoms, although their pathogenic roles are not as extensively documented. Neosetophoma typically infects the leaves, stems, or bark of host plants, leading to symptoms such as leaf spots, necrotic lesions, blight, and canker formation (Gruyter et al. 2010, Marin-Felix et al. 2019a). The leaf spots are usually small to medium in size, brown to dark brown, often with a yellow halo, and may coalesce, causing extensive tissue death (Marin-Felix et al. 2019a, Ebrahimi & Fotouhifar 2021). Neosetophoma has also been associated with stem dieback or branch decline in woody plants as well as decline symptoms in fruit trees and elms, respectively (Hashemi & Mohammadi 2016, Ebrahimi & Fotouhifar 2021).

Hosts: Neosetophoma species have been isolated from various host families, viz. Brassicaceae, Caprifoliaceae, Iridaceae, Malvaceae, Ranunculaceae and Salicaceae (Phookamsak et al. 2014, Karunarathna et al. 2017). They have been found on both monocotyledons and dicotyledons, including trees, shrubs, grasses, and herbaceous crops (Alidadi et al. 2019, Farr & Rossman 2025). They have also been reported from economically and ecologically important species such as apple (Malus domestica), elm (Ulmus spp.), ash (Fraxinus spp.), and various plants in Poaceae (Ebrahimi & Fotouhifar 2021). Neosetophoma species have been reported from legumes, maples, other hardwood trees and soil, often as saprobes or endophytes (Gruyter et al. 2010, Karunarathna et al. 2017, Marin-Felix et al. 2019a). This broad host range underlines the genus’s adaptability and potential impact on both natural ecosystems and agricultural systems.

Pathogen biology, disease cycle and epidemiology: The disease cycle of Neosetophoma species is not fully understood. Neosetophoma produce conidiomata on

Fig. 20. Maximum likelihood tree of Neopestalotiopsis species based on the concatenated ITS, tub2 and tef sequence data. The best scoring RAxML tree had a final likelihood value of -8335.116. The tree was rooted to Pseudopestalotiopsis avicenniae (MFLUCC 17-0434) and P. myanmarina (NBRC 112264). Estimated base frequencies were as follows: A = 0.223971, C = 0.279581, G = 0.195564, and T = 0.300884; substitution rates AC = 0.68043, AG = 2.40364, AT = 1.29389, CG = 0.68043, CT = 4.20028, and GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Type strains are in bold. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

infected plant tissues, which release conidia that serve as primary inoculum (Alidadi et al. 2019, Ebrahimi & Fotouhifar 2021). The conidia are disseminated by wind or water splashes, particularly during wet conditions, allowing them to infect new host tissues through natural openings or wounds (Magyar et al. 2016). Once established, the fungus may remain asymptomatic or cause symptoms such as leaf spots, necrosis, and stem lesions (Ebrahimi & Fotouhifar 2021). In some species, pseudothecia develop later in the disease cycle, contributing to long-term survival. The fungus may overwinter in infected plant debris or woody tissues, serving as a reservoir for infection in subsequent growing seasons. Neosetophoma can act as a primary pathogen, secondary invader or latent colonizer (Magyar et al. 2016). More studies are needed to clarify the infection strategies and disease cycles of Neosetophoma species.

Morphological-based identification: The identification of Neosetophoma was primarily based on a combination of macroscopic and microscopic characteristics. The colonies (PDA or MEA) range from olivaceous to dark gray or brown, often with a darkly pigmented reverse (Gruyter et al. 2010, Marin-Felix et al. 2019a). The conidiomata are pycnidial, globose to pyriform, brown to black, and sometimes possess setae around the ostioles, a distinctive feature that contributes to the genus name (Fig. 21). Conidiogenous cells are usually phialidic or annellidic, ampulliform to cylindrical, and line the inner walls of the pycnidia. The conidia are hyaline to pale brown, ellipsoidal to cylindrical, mostly 1-septate, and smooth-walled. The sexual morph is characterized by globose to subglobose, unilocular and ostiolate ascomata (Phookamsak et al. 2014, Mehrabi & Asgari 2024). The hamathecium comprises numerous, filiform, hyaline, pseudoparaphyses. Asci are bitunicate, fissitunicate, cylindrical and 8-spored. Ascospores are fusiform, smooth-walled, hyaline to brown and 1–3-septate, without a gelatinous sheath. Although these morphological traits are useful for preliminary identification, they often overlap with other genera such as Phoma and Didymella, making molecular analysis essential for accurate species-level classification.

Molecular-based identification: Molecular-based identification of Neosetophoma has become increasingly important due to the morphological overlap with other genera. Gruyter et al. (2010) introduced Neosetophoma and Setophoma in Phaeosphaeriaceae to accommodate Phoma species based on LSU and SSU sequences. The placement of the genus in Phaeosphaeriaceae was confirmed by several studies (De Gruyter et al. 2012, Zhang et al. 2012b, Hyde et al. 2013, Quaedvlieg et al. 2013, Phookamsak et al. 2014). Several novel species were further introduced based on morphology and multi-gene phylogeny (Hernandez-Restrepo et al. 2016, Karunarathna et al. 2017, Wanasinghe et al. 2018, Marin-Felix et al. 2019a). This study provides an updated phylogeny of Neosetophoma based on combined ITS, LSU, SSU, rpb2, tef and tub2 sequence data (Fig. 22, Table 11).

Recommended genetic markers (genus level): ITS and LSU

Fig. 21. Neosetophoma poaceicola. a Ascomata on host. b Vertical section through ascoma. c-d Asci. e-f Ascospores. Scale bar: b = 50 µm, c-d = 30 µm, e-f = 15 µm. This picture is copyright of Danushka S Tennakoon (Tennakoon et al. 2020).

Fig. 22. Maximum likelihood tree of Neosetophoma species based on the concatenated ITS, LSU, SSU, rpb2, tef and tub2 sequence data. The best scoring RAxML tree had a final likelihood value of -9494.891. The tree was rooted with Ampelomyces quisqualis (CBS 131.31 and CBS 133.32). Estimated base frequencies were as follows: A = 0.250, C = 0.250, G = 0.250, T = 0.250; substitution rates AC = 1.68999, AG = 5.54503, AT = 1.68999, CG = 1.00000, CT = 13.78932, GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Type strains are in bold. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A. Species confirmed with pathogenicity studies are marked with #.

Recommended genetic markers (species level): ITS, LSU, SSU, rpb2, tef and tub2

Accepted number of species: 28

References: Gruyter et al. (2010), Karunarathna et al. (2017), Alidadi et al. (2019), Marin-Felix et al. (2019a), Ebrahimi & Fotouhifar (2021) (morphology and phylogeny)

136. Nigrospora Zimm., Centbl. Bakt. ParasitKde, Abt. I 8: 220 (1902)

Type species: Nigrospora panici Zimm.

Classification: Sordariomycetes, Amphisphaeriales, Apiosporaceae

Background: Nigrospora is an important genus in ascomycetes with a cosmopolitan distribution and wide host range (Wang et al. 2017b). Nigrospora species are endophytes and saprobes of various hosts and have also been commonly recorded as plant pathogens with diverse host ranges including important economic crops, fruits and ornamentals (Sun et al. 2011, Wang et al. 2017b, Rashmi et al. 2019, Manawasinghe et al. 2025). In addition, this fungus can cause onychomycosis, corneal ulcers, hay fever, respiratory and allergic diseases in humans (De Hoog et al. 2000, Fan et al. 2009, Ananya et al. 2014).

Distribution: worldwide (Farr & Rossman 2025)

Disease symptoms: Nigrospora species can cause leaf spot as well as leaf, twigs, shoot and stem blight (Fig. 23). Nigrospora oryzae can cause minute black pustules on rice leaves, necrotic spots on rice grains and blight on glumes and culms, while on rice stems it produces grey to black lesions (Hajano et al. 2011). This species was reported to be highly pathogenic on cotton forming grey mycelium on bolls that later become discolored bolls (Palmateer et al. 2003, Sharma et al. 2013). This fungus also causes ear rot disease in maize with characteristic symptoms including superficial, dark grey to black lesions with a slight bluish cast on the lower inter-nodes (Sharma et al. 2013). Nigrospora oryzae also induces foliar symptoms in Kentucky bluegrass (Poa pratensis) in Ontario, Canada (Zheng et al. 2012). It was reported as seed-borne on barley, sorghum, and wheat and has been associated with postharvest diseases of citrus (Jagdish et al. 1985, Fakhrunnisa et al. 2006). Stem blight disease was observed on the lower portions of Brassica juncea stems during the cropping season, which was caused by Ni. oryzae in India (Sharma et al. 2013). Nigrospora sphaerica causes leaf blight symptoms on tea leaves in China and India (Dutta et al. 2015, Liu et al. 2016, 2020). Infected leaves have irregular-shaped yellow to brown leaf lesions, and later the entire leaf is infected and leaves turns dark brown and withered (Liu et al. 2020). Nigrospora sphaerica is an opportunistic pathogen causing onychomycosis in humans (De Hoog et al. 2000, Fan et al. 2009) and corneal ulcers (Ananya et al. 2014, Kindo et al. 2014, Motswaledi et al. 2018). Nigrospora musae is commonly known for causing squirter disease on banana plants (Jones and Stover 2000). The fungus resides on banana debris and sporulates after rainfall or after tree watering, and the disease develops during transportation and ripening of the fruit (Meredith 1961). The initial disease symptoms are a dark core or dark-red line of rubbery material along the center core or near the end of the banana fruit (Meredith 1961). This disease occurs in Australia, Jamaica, and Nigeria (Meredith 1961, https://agrobaseapp.com/).

Hosts: Nigrospora does not display evidence of host or geographical limitation (Palmateer et al. 2003, Wu et al. 2014a, Eken et al. 2016, Wang et al. 2017b, Orina et al. 2025). More than 900 records are found in the Fungal database (Farr & Rossman 2025). Hao et al. (2020) isolated five Nigrospora species from Shandong, China and found 13 novel host associations. Nigronapthaphenyl, a new compound was extracted from Ni. sphaerica (an endophyte) on Bruguiera gymnorrhyza (Ukwatta et al. 2019). Nigrospora sphaerica causes leaf spots, twigs and shoot blight of blueberry (Vaccinium corymbosum) (Wright et al. 2008). Foliar and cane rot of Arundo donax are caused by Ni. oryzae in Europe (Widmer et al. 2006). The same species caused lint rot or leaf spots of cotton in Alabama, China (Palmateer et al. 2003, Zhang et al. 2012a), stem blight on Brassica juncea in India (Sharma et al. 2013) and leaf spots in Aloe vera in China (Zhai et al. 2013). Nigrospora oryzae also causes leaf spots on Dendrobium candidum in China (Wu et al. 2014a) and on Kentucky bluegrass (Poa pratensis) in Ontario (Zheng et al. 2012). Nigrospora oryzae and Ni. sphaerica causes leaf spots on date palm leaves (Abass et al. 2014). Leaf blight on Stenotaphrum secundatum was caused by Nigrospora osmanthi in China (Mei et al. 2019). Nigrospora sphaerica causes leaf blight on Camellia sinensis and Cunninghamia lanceolata in China (Liu et al. 2016, Xu et al. 2017) and leaf spots of Kinnow mandarin in Pakistan (Alam et al. 2017, 2020). Reddish brown spot disease of dragon fruit (Hylocereus polyrhizus) was caused by Ni. lacticolonia and Ni. sphaerica in Malaysia (Kee et al. 2019).

Fig. 23. a-f Symptoms caused by Nigrospora sp. on sugarcane (Saccharum officinarum). This picture is copyright of Mubashar Raza (Raza et al. 2019).

Pathogen biology, disease cycle and epidemiology: Nigrospora spp., particularly Nigrospora oryzae, are primarily saprobes but can become weak pathogens under favourable conditions (Wang et al. 2017b). They produce dark, single-celled conidia in abundance, which are easily dispersed by wind, rain splash, and mechanical means (Magyar et al. 2016). The disease cycle typically begins when conidia land on senescing or wounded plant tissue and germinate under warm and humid conditions, hence colonizing the host (Doohan and Zhou 2017). Infection is often facilitated by plant stress or injury (Doohan and Zhou 2017). Disease development is favoured by warm temperatures and high humidity, especially in tropical and subtropical regions (Palmateer et al. 2003, Eken et al. 2016, Wang et al. 2017b, Orina et al. 2025). The pathogen survives in plant debris and infected plant parts, serving as the primary inoculum source for subsequent infections (Wang et al. 2017b).

Morphological-based identification: Species delimitation in Nigrospora was previously based on morphological characters, especially conidial dimensions (Mason 1927, 1933). The generic concept of Nigrospora has branched micronematous or semi-macronematous conidiophores, monoblastic conidiogenous cells and black, shiny, globose or subglobose, aseptate conidia while the sexual morphs comprise perithecial ascomata, short-stalked asci with biseriate ascospores (Webster 1952, Wang et al. 2017b, Raza et al. 2019). It was found that some key morphological characters (conidial dimensions) overlap, but there are phylogenetically distinct species of Nigrospora (Wang et al. 2017b). Thus, Wang et al. (2017) suggested using a combination of morphological and molecular data to distinguish Nigrospora species. The type of the genus, Ni. panici, was reported from Panicum amphibium from Java (Zimmerman 1902), but the holotype was lost (Wang et al. 2017b). Also, several species (Ni. aerophila, Ni. arundinacea, Ni. canescens, Ni. gallarum, Ni. gossypii, Ni. javanica, Ni. maydis, Ni. padwickii) are not available for molecular study, which to some extent has impeded the full resolution of species relationships (Wang et al. 2017b). Therefore, there is a need to find suitable specimens for epitypification of the type species and some other species.

Molecular-based identification: The phylogenetic investigations by Wang et al. (2017) significantly stabilized the taxonomy of the genus and affirmed the generic placement in Apiosporaceae (Xylariales) based on multi-locus molecular phylogeny of ITS, tef and tub2 gene regions. This study provides an updated phylogeny of Nigrospora based on combined ITS, tub2 and tef sequence data (Fig. 24, Table 12).

Recommended genetic markers (genus level): ITS and tub2

Recommended genetic markers (species level): ITS, tef and tub2

Accepted number of species: 46

References: Palmateer et al. (2003), Wu et al. (2014), Eken et al. (2016), Wang et al. (2017), Raza et al. (2019) (morphology and phylogeny)

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A.

Fig. 24. Maximum likelihood tree of Nigrospora species based on the concatenated ITS, tub2, and tef sequence data. The best scoring RAxML tree had a final likelihood value of -12521.122218. The tree was rooted with Apiospora species. Estimated base frequencies were as follows: A = 0.215197, C = 0.302061, G = 0.242950, T = 0.239792; substitution rates AC = 1.163224, AG = 3.223818, AT = 0.979868, CG = 1.107217, CT = 4.753930, GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Type strains are in bold. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

137. Oculimacula Crous & W. Gams, Eur. J. Pl. Path. 109: 845. (2003)

Type species: Oculimacula yallundae (Wallwork & Spooner) Crous & W. Gams

Classification: Leotiomycetes, Helotiales, Ploettnerulaceae

Background: To resolve the phylogenetic position and affinity of the genus Ramulispora and the sexual state of the eyespot fungi in the genus Tapesia, Crous et al. (2003) established Oculimacula to accommodate Tapesia and Helgardia, the asexual morph of Oculimacula. Following the one fungus, one name system, Johnston et al. (2014) treated the name Oculimacula as the eyespot disease complex instead of using Helgardia. They also placed H. aestiva (Nirenberg) Crous & W. Gams and H. anguioides (Nirenberg) Crous & W. Gams in Oculimacula as O. aestiva (Nirenberg) Crous and O. anguioides (Nirenberg) Crous, respectively (Johnston et al. 2014). Marin-Felix et al. (2019b) provided details of Oculimacula and provided the genetic information to identify the genus. Crous et al. (2021a) redefined Oculimacula by analysis of genetic sequence comparisons of ITS, tef, act, rpb1, rpb2 and LSU sequences. Subsequently, Oculimacula was reduced to two species, O. acuformis and O. yallundae, with O. aestiva placed in Cyphellophora and O. anguioides accommodated in a new genus, Helgardiomyces (Crous et al. 2021a, King et al. 2021). Oculimacula yallundae and O. acuformis induce the disease eyespot (Robertse et al. 1995, Crous et al. 2003, Ray et al. 2006, King et al. 2021). In temperate climates, eyespot is regarded as the most serious stem base disease of cereals (Crous et al. 2003, Ray et al. 2006). Field diagnosis of eyespot can be difficult since other diseases cause similar symptoms, complicating eyespot therapy techniques (King et al. 2021).

Distribution: Africa, Australasia, USA, UK, Western Europe, and other temperate areas of the world (Farr & Rossman 2025)

Disease symptoms: Oculimacula species are soil-borne, facultative fungi that cause eyespot disease of wheat (Fig. 25). It comprises a major component of the stem-base disease complex of wheat in temperate regions of the world with a cool and wet climate (Lucas et al. 2000, Crous et al. 2003, Wei et al. 2011). Eyespot disease of wheat generally occurs in autumn. The pathogen first infects the coleoptiles of wheat, then develops into the stem sheaths and stalks, producing typical eyespot lesions. In general, eyespot lesions form on leaf sheaths and culms that are close to the soil. The lesions cause yellowing of the straw, often with a black pupil-like spot in the center surrounded by a green-brown to dark brown ring. When the infection is severe, the wheat stems are weakened at the point of infection, making them prone to lodging, twisting, bending, and even breaking from the lesions (Lucas et al. 2000, Prescott et al. 2007, Wei et al. 2011, King et al. 2021). Eyespot lesions can cause moderate or severe damage to wheat, interfering with the movement of nutrients and water through the stem, resulting in a loss of wheat yield (Glynne & Salt 1958, Clarkson 1981, Lucas et al. 2000, Crous et al. 2003).

Hosts: Oculimacula has a wide range of hosts in cereals and wild grass species belonging to Poaceae, such as Bromus diandrus, Hordeum leporinum, Holcus lanatus, wheat, barley, rye, and oats with wheat being the most susceptible (Wallwork 1987, Murray et al. 1994, Lucas et al. 2000, Dyer & Bradshaw 2002, Chapman et al. 2008, Farr & Rossman 2025).

Pathogen biology, disease cycle and epidemiology: Eyespot is caused by the pathogens Oculimacula acuformis and O. yallundae (Marin-Felix et al. 2019b). Both Oculimacula species can enter the outer leaf sheath of the host directly through the epidermal cells or more commonly through the stomata. Normally, the fungi form a stroma on the host surface, from where infection occurs, and they then continue to grow, forming a pupil-like lesion and killing the tiller before the host head sprouts (Sprague & Fellows 1934). Oculimacula spp. develop in the host and produce many conidia in spring and autumn. The conidia are spread by rain splash. Oculimacula also reproduce sexually, producing ascospores that can lead to more infections. There is a second pathway of dispersal for spores in that a few spores can be transmitted and infect new hosts through the air, but Foëx and Rosella (1930) indicated that this conclusion needs to be confirmed by further research. Oculimacula spp. overwinter on diseased plants, and in spring produce spores that infect the newly seeded host (Sprague & Fellows 1934, Murray 1996, Douhan et al. 2002).

Morphological-based identification: Oculimacula produces sessile ascomata that are gregarious, circular to lobate and are formed on a subiculum of white to dark brown hyphae that are linked to the substrate by a superficial mat of light brown substratum. The apothecia are grey with a pale grey margin, emarginate and flattened to convex when they reach maturity. The receptacle is cup-shaped and pale brown to grey-brown. The medullary excipulum is composed of multiseptate, hyaline hyphae. The ectal excipulum of thin-walled, dark brown, angular cells, becomes more elongated towards the margin. Filiform paraphyses have obtuse ends and are as long as the asci. Asci are 8-spored, clavate to subcylindrical or fusoid, with a short stalk and an apical pore that can be stained blue by Melzer's reagent. Ascospores are mostly straight, bi- to multiseriate, hyaline, smooth, aseptate, fusoid to subcylindrical or clavate with rounded ends (Crous et al. 2003, Marin-Felix et al. 2019b).

Molecular-based identification: Crous et al. (2003) established this genus by using ITS and LSU sequences. The phylogenetic analysis indicates that there are four species in this genus (including the asexual morph of Oculimacula). Johnston et al. (2014) treated Oculimacula to include the asexual morph, Helgardia, based on the principle of “One fungus one name”. Marin-Felix et al. (2019b) provided identification of genes from species to genus and species levels. Crous et al. (2021a) redefined Oculimacula based on ITS, tef, act, rpb1, rpb2, and LSU sequences. When combined with morphological features, this genus is restricted to two species, both of which are important wheat pathogens (Crous et al. 2021a, King et al. 2021). This study provides an updated phylogeny of Oculimacula based on combined ITS, LSU, act, rpb2 and tef sequence data (Fig. 26, Table 13).

Recommended genetic markers (genus level): LSU

Recommended genetic markers (species level): ITS, LSU, act, rpb2 and tef

Accepted number of species: two

Reference: Crous et al. (2021a) (morphology and phylogeny)

138. Phaeosphaeriopsis M.P.S. Câmara, M.E. Palm & A.W. Ramaley, in Câmara et al., Mycol. Res. 10(5): 51 (2003)

Fig. 25. Symptoms caused by Oculimacula sp. on wheat. a Eyespot lodging. b-c Eyespots. d Whiteheads of wheat. This picture is copyright of Pedro Crous (Marin-Felix et al. 2019b).

Fig. 26. Maximum likelihood tree of Oculimacula species based on the concatenated ITS, LSU, act, rpb2 and tef sequence data. The best scoring RAxML tree had a final likelihood value of -6423.795453. The tree was rooted with Rhynchobrunnera agropyri (CBS 146762). Estimated base frequencies were as follows: A = 0.245708, C = 0.247574, G = 0.253386, T = 0.253333; substitution rates AC = 1.294731, AG = 2.248453, AT = 1.136711, CG = 0.484296, CT = 6.594746, GT = 1.000000. ML bootstrap support values over 70% and Bayesian posterior probabilities (BPP) greater than 0.95 are indicated. Scale bar indicates the number of substitutions per site. Hyphens (-) represent support values less than BS 70%/BPP 0.95.

Ex-type/ex-epitype/ex-neotype/ex-lectotype strains and voucher strains are in bold. Sequences that are not available are represented by N/A.

Type species: Phaeosphaeriopsis glaucopunctata (Grev.) M.P.S. Câmara, M.E. Palm & A.W. Ramaley

Classification: Dothideomycetes, Pleosporales, Phaeosphaeriaceae

Background: Phaeosphaeriopsis was established by Câmara et a l. (2003) to include several species which could not be categorized as Paraphaeosphaeria based on morphological characters and SSU molecular data. Câmara et al. (2003) transferred Paraphaeosphaeria agavensis, Pa. glauco-punctata, Pa. nolinae and Pa. obtusispora to Phaeosphaeriopsis and described a new species P. amblyspora. Phaeosphaeriopsis represents species that produce uni- or multi-loculate stromata, 4–5-septate, punctate or verrucose ascospores and have asexual morphs characterized by 0–3-septate, cylindrical, brown conidia or sometimes bacillar microconidia (Câmara et al. 2003, Phookamsak et al. 2014, Thambugala et al. 2014, Marin-Felix et al. 2019a).

Distribution: worldwide (Farr & Rossman 2025)

Disease symptoms: Phaeosphaeriopsis species can cause leaf spots and leaf on several hosts (Fig. 27). Phaeosphaeriopsis glauco-punctata has been reported as the causative agent (as “Paraphaeosphaeria glauco-punctata” in earlier studies) of leaf spot and necrosis on Ruscus (Camara et al. 2001). The symptoms are 2 mm wide, discoloured lesions that gradually develop into larger brown necrotic lesions. When infection is severe, these lesions coalesce and form large necrotic regions sometimes leading to leaf necrosis (Camara et al. 2001, Golzar & Wang 2012).

Hosts: Phaeosphaeriopsis glauco-punctata and Pa. triseptata have been reported on various Ruscus spp. from Europe. Phaeosphaeriopsis amblyspora was described from dead leaves of Yucca baccata (Câmara et al. 2003). Phaeosphaeriopsis agapanthi was isolated from Agapanthus precox. Phaeosphaeriopsis agavacearum and Pa. pseudoagavacearum have been reported from Agave sp. (Crous et al. 2016a, b). Phaeosphaeriopsis aloes and Pa. aloicola, were both recorded from leaves of Aloe sp. in the USA (Marin-Felix et al. 2019a). Phaeosphaeriopsis musae was associated with leaf spots on Musa sp. (Arzanlou and Crous 2006). Phaeosphaeriopsis dracaenicola was reported from Dracaena loureiroi in Thailand (Phookamsak et al. 2014), while Phaeosphaeriopsis grevilleae was collected on leaves of Grevillea sp. from Queensland, Australia (Marin-Felix et al. 2019a). Phaeosphaeriopsis beaucarneae was isolated from dead leaves of Beaucarnea recurvata (Tennakoon et al. 2020). Phaeosphaeriopsis omaniana, Pa. sansevieriae and Pa. triseptata were isolated from the leaves of Dracaena serrulata, Sansevieria hyacinthoides and Ruscus aculeatus, respectively (Thambugala et al. 2014, Al-Jaradi et al. 2020, Crous et al. 2021b, Farr & Rossman 2025).

Fig. 27. a Symptoms caused by Phaeosphaeriopsis sp. on Agapanthus precox. b Conidiomata sporulating on pine needle agar. c Conidiogenous cells giving rise to conidia. d Conidia. Scale bars: c-d = 10 µm. This picture is copyright of Pedro Crous (Crous et al. (2016b) and Marin-Felix et al. (2019a)).

Phaeosphaeriopsis consists mostly of saprobic species. However, Phaeosphaeriopsis glaucopunctata has been reported to cause leaf spot and necrosis in Ruscus aculeatus (Câmara et al. 2003, Golzar & Wang 2012). Phaeosphaeriopsis agapanthi on Agapanthus precox and P. dracaenicola on Dracaena lourieri have been associated with causing necrotic leaf spots on these hosts (Phookamsak et al. 2014, Crous et al. 2016a). Phaeosphaeriopsis musae is assumed to be a weak pathogen or secondary invader (Arzanlou & Crous 2006) as it has often been found associated with leaf lesions of Mycosphaerella fijiensis, Sphaerulina musae and Cordana musae. The first report of Pa. glaucopunctata causing leaf spot and foliage necrosis on Ruscus aculeatus in Australia was reported by Golzar & Wang (2012).

Pathogen biology, disease cycle and epidemiology: Many Phaeosphaeriopsis species are associated with leaf spots, blights, or necrotic lesions (Camara et al. 2001, Golzar & Wang 2012). Phaeosphaeriopsis are generally considered necrotrophic pathogens and they often affect stressed or senescing plant material (Phookamsak et al. 2014, Thambugala et al. 2014, Marin-Felix et al. 2019a). Phaeosphaeriopsis species release spores under moist conditions, which are spread by rain splash, wind, or mechanical means (van Niekerk et al. 2010). The disease cycle begins when spores land on susceptible plant surfaces and germinate under high humidity or prolonged leaf wetness. Infection often starts at the leaf margins or tips, especially where mechanical damage or stress is present (Adaskaveg et al. 2008, Gonçalves et al. 2013). Lesions typically expand during periods of wet weather, favouring rapid disease development. The fungus survives between growing seasons as saprobes on crop residues or in infected plant tissue (Adaskaveg et al. 2008, van Niekerk et al. 2010).

Morphological-based identification: Phaeosphaeriopsis species produce uni- or multi-loculate stromata, scattered, semi-immersed to erumpent, globose to subglobose, ostiolate ascomata, cylindrical to broadly fusoid or clavate asci and 4–5-septate, yellowish to light brown, verrucose ascospores. The asexual morph is coniothyrium-like or phaeostagonospora-like (Câmara et al. 2003, Quaedvlieg et al. 2013, Thambugala et al. 2014, Marin-Felix et al. 2019a). Morphologically, the conidia are similar to Paraphaeosphaeria species but can be distinguished based on the different number of septa in ascospores (4–5-septate conidia vs. 2-septate ascospores) and asexual morphs (Câmara et al. 2003). Only a few species have both sexual and asexual morphs e.g. Pa. beaucarneae isolated from Beaucarnea recurvata (Tennakoon et al. 2020). For species such as Pa. agapanthi, Pa. aloes, Pa. grevilleae, Pa. pseudoagavacearum, Pa. sansevieriae and Pa. triseptata, only the asexual morph is known and for example in Pa. aloicola and Pa. obtusispora only the sexual morph is known. The asexual morphs of Pa. amblyspora and Pa. agavensis produce hyaline, bacillar conidia from simple phialides. The asexual morphs of Pa. glauco-punctata, Pa. obtusispora, and Pa. nolinae produce brown conidia from percurrently proliferating conidiogenous cells. In Pa. nolinae, the conidia are 3-septate and in Pa. glauco-punctata often 1-septate, while those in Pa. obtusispora are usually aseptate (Câmara et al. 2003, Thambugala et al. 2014). Thambugala et al. (2014) suggested that it was unlikely that Pa. phacidiomorpha belonged to Phaeosphaeriopsis due to its different asci characters, hyaline ascospores and microsphaeropsis-like asexual morphs. This requires further investigation. Thus, Phaeosphaeriopsis is a genus that requires more collections in order to establish its taxonomic boundaries, sexual-asexual connections and diversity.

Molecular-based identification: Thambugala et al. (2014) accepted seven species in Phaeosphaeriopsis based on multi-gene analysis of LSU, SSU, ITS and rpb2 regions but excluded P. musae. However, in our phylogenetic analysis, the type strain of P. musae was included, and it grouped with other species of the genus. Marin-Felix et al. (2019a) accepted 15 species, including four new species in Phaeosphaeriopsis based on multi-gene analysis of ITS, LSU, rpb2, tef and tub2 genes.

Phaeosphaeriopsis pseudoagavacearum is closely related to Pa. agavacearum (Marin-Felix et al. 2019a) and both were recorded from Agave sp. However, Pa. aloes and Pa. aloicola, which were isolated from Aloe sp. are phylogenetically distant, with Pa. aloes more closely related to Pa. obtusispora and Pa. aloicola to Pa. agapanthi (Marin-Felix et al. 2019a). A similar result was also observed in our phylogenetic analysis. This study provides an updated phylogeny of Phaeosphaeriopsis based on combined ITS, LSU, rpb2 and tef sequence data (Fig. 28, Table 14).

Recommended genetic markers (genus level): LSU

Recommended genetic markers (species level): ITS, LSU, rpb2 and tef

Accepted number of species: 20

References: Câmara et al. (2003), Thambugala et al. (2014), Marin-Felix et al. (2019a), Sun et al. (2025) (morphology and phylogeny); Golzar & Wang (2012) (pathogenicity)