Research Article
Research Article
Segregation of the genus Parahypoxylon (Hypoxylaceae, Xylariales) from Hypoxylon by a polyphasic taxonomic approach
expand article infoMarjorie Cedeño-Sanchez§, Esteban Charria-Girón§, Christopher Lambert§, J. Jennifer Luangsa-ard|, Cony Decock, Raimo Franke#, Mark Brönstrup#¤, Marc Stadler§
‡ Helmholtz Centre for Infectiion Research, Braunschweig, Germany
§ Technische Universität Braunschweig, Braunschweig, Germany
| National Center for Genetic Engineering and Biotechnology, Pathumthani, Thailand
¶ Mycothéque de l’ Universite catholique de Louvain, Louvain-la-Neuve, Belgium
# Helmholtz Centre for Infection Research, Braunschweig, Germany
¤ German Center for Infection Research, Braunschweig, Germany
Open Access


During a mycological survey of the Democratic Republic of the Congo, a fungal specimen that morphologically resembled the American species Hypoxylon papillatum was encountered. A polyphasic approach including morphological and chemotaxonomic together with a multigene phylogenetic study (ITS, LSU, tub2, and rpb2) of Hypoxylon spp. and representatives of related genera revealed that this strain represents a new species of the Hypoxylaceae. However, the multi-locus phylogenetic inference indicated that the new fungus clustered with H. papillatum in a separate clade from the other species of Hypoxylon. Studies by ultrahigh performance liquid chromatography coupled to diode array detection and ion mobility tandem mass spectrometry (UHPLC-DAD-IM-MS/MS) were carried out on the stromatal extracts. In particular, the MS/MS spectra of the major stromatal metabolites of these species indicated the production of hitherto unreported azaphilone pigments with a similar core scaffold to the cohaerin-type metabolites, which are exclusively found in the Hypoxylaceae. Based on these results, the new genus Parahypoxylon is introduced herein. Aside from P. papillatum, the genus also includes P. ruwenzoriense sp. nov., which clustered together with the type species within a basal clade of the Hypoxylaceae together with its sister genus Durotheca.


Ascomycota, metabolite annotation, one new genus, one new species, phylogeny, polythetic taxonomy, Xylariales


The genus Hypoxylon Bull. 1791 remains one of the largest in the Xylariales, even after a turbulent taxonomic history, during which its generic concept has changed drastically. Its early taxonomic history has been reviewed in great detail by Ju and Rogers (1996). Therefore, we largely refer to this monograph for the taxonomic treatments that occurred in the 19th and early 20th century.

The first world monograph of Hypoxylon by Miller (1961) was mainly based on stromatal morphology and ascal micromorphology. He recognized four sections (Hypoxylon, Annulata, Applanata and Papillata, the latter of which was further subdivided into two subsections, Papillata and Primocinerea). Ju and Rogers (1996) then restricted Hypoxylon to sections Hypoxylon and Annulata, and included several species of section Papillata in their emended section Hypoxylon. The main criteria for this taxonomic change were the presence of stromatal pigments and a nodulisporium-like anamorph. Many of the species in sections Applanata and Papillata sensu Miller (1961) do not show the latter mentioned features and were accommodated in other genera (e.g., Biscogniauxia, Nemania, Whalleya), which were later transferred to different families (Wendt et al. 2018). For their current classification, we refer to Hyde et al. (2020).

With the advent of molecular phylogenetic studies, and chemotaxonomy as an additional tool, the taxonomic concepts of Hypoxylon and other stromatic genera of the Xylariales have been further refined. The holomorphic concepts developed by Ju and Rogers, as well as other mycologists who put more emphasis on the anamorphic characters than on stromatal and ascospore morphology, have largely been confirmed. Hsieh et al. (2005) used protein-coding genes of a large number of representative taxa to resolve the phylogeny of Hypoxylon s. lat., which resulted in the recognition of the genus Annulohypoxylon. The composition of the latter genus was then equivalent to that of sect. Annulata sensu Ju and Rogers (1996). Notably, a parallel approach to establish a phylogeny based on ITS nrDNA sequences resulted in a very low resolution of the hypoxyloid taxa (Triebel et al. 2005). Later studies revealed that a multi locus phylogeny involving both protein-coding genes and rDNA are suitable to achieve a sufficient phylogenetic resolution within Hypoxylon and its allies (Kuhnert et al. 2014b, 2015, 2017a; Sir et al. 2015) in scope of a polythetic concept. Concurrent chemotaxonomic studies have aided in establishing correlations between the genotypes and the phenotypes of these pyrenomycetes. Their stromatal pigments, as well as certain secondary metabolites of their mycelial cultures, turned out to be informative for taxonomic segregation at the species or even genus level (cf. Helaly et al. 2018; Becker and Stadler 2021).

Based on the above accomplishments, Wendt et al. (2018) proposed a rearrangement of the families of the stromatic Xylariales, as well as the further segregation of genera from the mainstream of Hypoxylon. The Hypoxylaceae were resurrected to accommodate Hypoxylon and its closely related allies, and the Xylariaceae were restricted to the genera with geniculosporium-like anamorphs, which had already been recognized as phylogenetically distinct in earlier studies (e.g., Hsieh et al. 2010). Annulohypoxylon was further subdivided and largely restricted to those species that have ostiolar rings and do not produce cohaerin-type azaphilones. The genus Jackrogersella was erected to accommodate those species of Annulohypoxylon sensu Hsieh et al. (2005) that produce the aforementioned compounds and have papillate ostioles without rings. In addition, the genus Pyrenopolyporus was erected for species of Hypoxylon sensu Ju and Rogers (1996) that have massive stromata, long tubular perithecia, contain naphthopyrones in their stromata and (where this is known) produce a characteristic virgariella-like anamorph. A follow-up study by Lambert et al. (2019) provided evidence that the species of the H. monticulosum complex differ from Hypoxylon by the production of antifungal sporothriolides in culture. In addition, these fungi also lack the typical stromatal pigments of Hypoxylon (Fig. 1) and appear in a basal clade in the molecular phylogeny. The genus Hypomontagnella was therefore introduced to accommodate them.

Figure 1. 

Characteristic stromatal pigments and other secondary metabolites of Hypoxylon species. (+)-mitorubrin (1); (+)-6˝-hydroxymitorubrinol acetate (2); (+)-mitorubrinol acetate (3); (+)-6˝-hydroxymitorubrinol (4); (+)-mitorubrinol (5); sporothriolide (6); dihydroisosporothric acid (7); cohaerin E (8); 8-methoxy naphthol (9); 1,8-naphthol (10); hypoxylone (11).

The genus Hypoxylon in the current sense still appears heterogeneous and paraphyletic in the recently established phylogenies, also because its type species, H. fragiforme clustered in a relatively small clade comprising only a few species such as H. howeanum, H. ticinense and H. rickii (Wendt et al. 2018; Lambert et al. 2021). The latter species have in common that their stromatal pigments are of the mitorubrin type.

Another species that was retained in Hypoxylon, even though the DNA sequences of the only available strain formed an aberrant clade in the phylogeny by Wendt et al. (2018) is H. papillatum Ellis & Everh. This species is characterized by effused-pulvinate stromata featuring long tubular perithecia. Therefore, its stromata somewhat resemble those of Pyrenopolyporus and certain Daldinia species such as D. placentiformis that do not have internal concentric zones. Ju and Rogers (1996) have studied the type material and concluded that the syntypes they studied from BPI and NY (i.e., the specimens listed in the protologue by Ellis and Everhart (see Smith 1893) did not all correspond to the same taxon. They identified some of the specimens as Hypoxylon placentiforme (now: Daldinia placentiformis), which was confirmed by Stadler et al. (2014) in the Daldinia world monograph, and selected a lectotype from Ohio (Commons No. 2160) which showed a characteristic morphology and could easily be distinguished from the former taxon. They also listed several other specimens from North America and Trinidad that showed the same characteristics.

Rogers (1985) cultured this fungus and provided a detailed description of its nodulisporium-like anamorph and culture. The corresponding specimen was collected by him in West Virginia, USA, and could have served as epitype. The culture is deposited in ATCC and showed the typical characteristics of H. papillatum sensu Rogers (1985) and was included in the phylogeny by Wendt et al. (2018) as a representative of this taxon. However, it showed an aberrant phylogenetic position in a clade that appeared basal to the others in which the DNA sequences of Hypoxylon species were located. We have come across a very similar fungus that was collected in Central Africa and have studied it, along with several extant type and authentic specimens for comparison. The results of this study, which also relies on state-of-the art metabolomics, are reported herein.

Materials and methods

Sample sources

All scientific names of fungi are given without authorities or publication details, according to Index Fungorum ( Type and reference specimens were provided by Washington State University herbarium (WSP), U.S. National Fungus Collections (BPI) and the New York Botanical Garden (NY), USA. Fungal cultures were provided from the Belgian Coordinated Collections of Microorganisms (MUCL), Belgium and the Westerdijk Fungal Biodiversity Institute (CBS), The Netherlands.

Morphological characterization

The microscopic characteristics of the teleomorph were carried out as described by Pourmoghaddam et al. (2020). To observe the macro-morphology of the cultures, the strains were grown on Difco Oatmeal Agar (OA), 2% Malt Extract Agar (MEA) and Yeast Malt agar (YM agar; malt extract 10 g/L, yeast extract 4 g/L, D-glucose 4 g/L, agar 20 g/L, pH 6.3 before autoclaving) and the cultures checked at 15 days after inoculation. Pigment colors were determined following the color-codes by Rayner (1970).

DNA extraction, PCR and sequencing

The DNA was extracted from pure cultures grown on plates with YM agar. Small amounts of mycelia were harvested after five days of growth and transferred to a 1.5 ml homogenization tube filled with six to eight Precellys Ceramic beads (1.4 mm, Bertin Technologies, Montigny-le-Bretonneux, France).

DNA extraction was performed using the commercially available Fungal gDNA Miniprep Kit EZ-10 spin column (NBS Biologicals, Cambridgeshire, UK) following the manufacturer’s instructions. The tub2 (partial β-tubulin) gene region was amplified using the primers T1 and T22 (O’Donnell and Cigelnik 1997); ITS (nuc rDNA internal transcribed spacer) region using the primers ITS4 and ITS5 (White et al. 1990); LSU (Large subunit nuc 28S rDNA) using LR0R and LR7 (Vilgalys and Hester 1990) and rpb2 (partial second largest subunit of the DNA-directed RNA polymerase II) using fRPB2-5F and fRPB2-7cR (Liu et al. 1999).

PCR reactions were performed by mixing template gDNA (2–3 µL), 12.5 µL JumpStart Taq Ready Mix (Sigma Aldrich, Deisenhofen, Germany), 0.5 µL of both forward and reverse primers (10 mM) and 8.5 to 9.5 μl of sterile filtered and sterilized water to a final volume of 25 µL. Amplification was achieved using a Mastercycler nexus Gradient (Eppendorf, Hamburg, Germany). Thermocycling for ITS commenced with an initial denaturation at 94 °C for 5 min followed by 34 cycles of denaturation (30 s at 94 °C), annealing (30 s at 52 °C), and elongation (1 min at 72 °C). The program concluded with a 10 min lasting elongation at 72 °C and reaction tubes were stored at 4 °C until further use. In the case of the other loci, the following steps were modified: LSU denaturation (1 min at 94 °C), annealing (1 min at 52 °C), and elongation (2 min at 72 °C); For tub2 the cycle repetitions were raised to 38, annealing (30 s at 47 °C) and elongation (2 min 30 s at 72 °C); for rpb2, the cycle repetitions were raised to 38, annealing (1 min at 54 °C) and elongation (1 min 30 s at 72 °C).

Molecular phylogenetic analyses

Sequences were analyzed and processed in Geneious 7.1.9 (Kearse et al. 2012). The generated sequence data were complemented by available sequence data from GenBank and the data sets for each genetic marker were aligned using MAFFT online (, Katoh et al. 2019), and manually curated in MEGA 11 (Tamura et al. 2021). A maximum-likelihood phylogenetic tree was constructed using IQ-TREE v. 2.1.3 [-b 1000 -abayes -m MFP -nt AUTO] (Minh et al. 2020), The selection of the appropriate nucleotide exchange model was selected by ModelFinder (Chernomor et al. 2016; Kalyaanamoorthy et al. 2017) based on Bayesian inference criterion. Branch support was calculated with non-parametric bootstrap (Felsenstein 1985 and approximate Bayes test (Anisimova et al. 2011). The total 1000 bootstrap replicates were mapped onto the ML tree with the best (highest) ML score. Single locus trees were calculated following the identical methodology and checked for congruence with the multigene phylogenetic tree.

A second phylogenetic inference was carried out following a Bayesian approach using MrBayes 3.2.7a (Ronquist et al. 2012) with algorithm options set to the ones reported by Matio Kemkuignou et al. (2022). The data matrix was subjected to PartitionFinder2 (Lanfear et al. 2016) as implemented in the program package phylosuite v. 1.2.2 (Zhang et al. 2020) with settings set to an un-linked determination of the best-fitting nucleotide substitution models following Bayesian information criterion (BIC) for the different partitions, restricted to the ones available in MrBayes. Posterior probabilities (PP) above 95% were regarded as significant. To determine the congruence of the topologies of ML and Bayes, an approximate unbiased (AU) topology test was carried out in IQ-TREE [iqtree -s example.phy -z example.treels -n 0 -zb 10000 -zw -au](Shimodaira 2022). All sequences used for the pyhlogeny are listed in Table 1.

Table 1.

Strains used in the phylogenetic analyses, including the strain IDs, GenBank accession numbers, and the references where the sequence data have been originally generated. Type specimens are labeled with T (holotype), IT (isotype), PT (paratype) and ET (epitype).

Species Strain number GenBank Accession Number Origin References
ITS LSU rpb2 tub2
Annulohypoxylon annulatum CBS 140775 KY610418 KY610418 KY624263 KX376353 USA (ET) Kuhnert et al. (2017a; tub2), Wendt et al. (2018: ITS, LSU, rpb2)
Annulohypoxylon michelianum CBS 119993 KX376320 KY610423 KY624234 KX271239 Spain Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Annulohypoxylon truncatum CBS 140778 KY610419 KY610419 KY624277 KX376352 USA (ET) Kuhnert et al. (2017a; tub2), Wendt et al. (2018; ITS, LSU, rpb2)
Daldinia bambusicola CBS 122872 KY610385 KY610431 KY624241 AY951688 Thailand (T) Hsieh et al. (2005; tub2), Wendt et al. (2018; ITS, LSU, rpb2)
Daldinia childiae CBS 122881 KU683757 MH874773 KU684290 KU684129 France (T) U’Ren et al. (2016; ITS, tub2, rpb2), Vu et al. (2019; LSU)
Daldinia concentrica CBS 113277 AY616683 KY610434 KY624243 KC977274 Germany Triebel et al. (2005; ITS), Kuhnert et al. (2014a; tub2), Wendt et al. (2018; LSU, rpb2)
Daldinia dennisii CBS 114741 JX658477 KY610435 KY624244 KC977262 Australia (T) Stadler et al. (2014; ITS), Kuhnert et al. (2014a; tub2), Wendt et al. (2018; LSU, rpb2)
Daldinia eschscholtzii MUCL 45435 JX658484 KY610437 KY624246 KC977266 Benin Stadler et al. (2014a; ITS), Kuhnert et al. (2014a; tub2), Wendt et al. (2018; LSU, rpb2)
Daldinia petriniae MUCL 49214 AM749937 KY610439 KY624248 KC977261 Austria (ET) Bitzer et al. (2008; ITS), Kuhnert et al. (2014a; tub2), Wendt et al. (2018; LSU, rpb2)
Daldinia placentiformis MUCL 47603 AM749921 KY610440 KY624249 KC977278 Mexico Stadler et al. (2014a; ITS), Kuhnert et al. (2014a; tub2), Wendt et al. (2018; LSU, rpb2)
Daldinia vernicosa CBS 119316 KY610395 KY610442 KY624252 KC977260 Germany (ET) Kuhnert et al. (2014a; tub2), Wendt et al. (2018; ITS, LSU, rpb2)
Durotheca rogersii YMJ 92031201 EF026127 JX507794 EF025612 Taiwan Ju et al. (2007) as Theissenia
Durotheca comedens YMJ 90071615 EF026128 JX507793 EF025613 Taiwan (T) Ju et al. (2003) as Theissenia
Durotheca crateriformis GMBC0205 MH645426 MH645425 MH645427 MH049441 China (T) de Long et al. (2019)
Durotheca guizhouensis GMBC0065 MH645423 MH645421 MH645422 MH049439 China (T) de Long et al. (2019)
Durotheca rogersii GMBC0204 MH645433 MH645434 MH645435 MH049449 China de Long et al. (2019)
Graphostroma platystomum CBS 270.87 JX658535 DQ836906 KY624296 HG934108 France (T) Zhang et al. (2006; LSU), Stadler et al. (2014; ITS), Koukol et al. (2015; tub2), Wendt et al. (2018; rpb2)
Hypomontagnella barbarensis STMA 14081 MK131720 MK131718 MK135891 MK135893 Argentina (T) Lambert et al. (2019)
Hypomontagnella monticulosa MUCL 54604 KY610404 KY610487 KY624305 KX271273 French Guiana Wendt et al. (2018)
Hypomontagnella submonticulosa CBS 115280 KC968923 KY610457 KY624226 KC977267 France Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon addis MUCL 52797 KC968931 ON954141 OP251037 KC977287 Ethiopia (T) Kuhnert et al. (2014a; ITS, tub2), This study
Hypoxylon aveirense MUM 19.40 MN053021 ON954142 OP251028 MN066636 Portugal (T) Vicente et al. (2021; ITS, tub2), This study
Hypoxylon baruense UCH9545 MN056428 ON954143 MK908142 Panama (T) Cedeño–Sanchez et al. (2020; ITS, tub2); This study
Hypoxylon canariense MUCL 47224 ON792787 ON954140 OP251029 ON813073 Spain, Canary Islands (PT) This study. (Species described by Stadler et al. 2008)
Hypoxylon carneum MUCL 54177 KY610400 KY610480 KY624297 KX271270 France Wendt et al. (2018)
Hypoxylon cercidicola CBS 119009 KC968908 KY610444 KY624254 KC977263 France Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon chionostomum STMA 14060 KU604563 ON954144 OP251030 ON813072 Argentina Sir et al. (2016; ITS); This study
Hypoxylon chrysalidosporum FCATAS2710 OL467294 OL615106 OL584222 OL584229 China (T) Ma et al. (2022)
Hypoxylon crocopeplum CBS 119004 KC968907 KY610445 KY624255 KC977268 France Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon cyclobalanopsidis FCATAS2714 OL467298 OL615108 OL584225 OL584232 China (T) Ma et al. (2022)
Hypoxylon erythrostroma MUCL 53759 KC968910 ON954154 OP251031 KC977296 Martinique Kuhnert et al. (2014a; ITS2, TUB), This study
Hypoxylon eurasiaticum MUCL 57720 MW367851 MW373852 MW373861 Iran (T) Lambert et al. (2021)
Hypoxylon fendleri MUCL 54792 KF234421 KY610481 KY624298 KF300547 French Guiana Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon ferrugineum CBS 141259 KX090079 KX090080 Austria Friebes and Wendelin (2016)
Hypoxylon fragiforme MUCL 51264 KC477229 KM186295 MK887342 KX271282 Germany (ET) Stadler et al. (2013; ITS), Daranagama et al. (2015; LSU, rpb2), Wendt et al. (2018; tub2)
Hypoxylon fuscoides MUCL 52670 ON792789 ON954145 OP251038 ON813076 France (T) This study. (Species described by Fournier et al. 2010a)
Hypoxylon fuscum CBS 113049 KY610401 KY610482 KY624299 KX271271 Germany (ET) Wendt et al. (2018)
Hypoxylon gibriacense MUCL 52698 KC968930 ON954146 OP251026 ON813074 France (T) Kuhnert et al. (2014a; ITS). This study
Hypoxylon griseobrunneum CBS 331.73 KY610402 KY610483 KY624300 KC977303 India (T) Kuhnert et al. (2014a; tub2), Wendt et al. (2018; ITS, LSU, rpb2)
Hypoxylon guilanense MUCL 57726 MT214997 MT214992 MT212235 MT212239 Iran (T) Pourmoghaddam et al. (2020)
Hypoxylon haematostroma MUCL 53301 KC968911 KY610484 KY624301 KC977291 Martinique (ET) Wendt et al. (2018; LSU, rpb2), Kuhnert et al. (2014a; ITS, tub2),
Hypoxylon hainanense FCATAS2712 OL467296 OL616132 OL584224 OL584231 China (T) Ma et al. (2022)
Hypoxylon hinnuleum ATCC 36255, MUCL 3621 MK287537 MK287549 MK287562 MK287575 USA (T) Sir et al. (2019)
Hypoxylon howeanum MUCL 47599 AM749928 KY610448 KY624258 KC977277 Germany Bitzer et al. (2008; ITS), Kuhnert et al. (2014a; tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon hypomiltum MUCL 51845 KY610403 KY610449 KY624302 KX271249 Guadeloupe Wendt et al. (2018)
Hypoxylon invadens MUCL 51475 MT809133 MT809132 MT813037 MT813038 France (T) Becker et al. (2020)
Hypoxylon investiens CBS 118183 KC968925 KY610450 KY624259 KC977270 Malaysia Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon isabellinum MUCL 53308 KC968935 ON954155 OP251032 KC977295 Martinique (T) Kuhnert et al. (2014a; ITS, tub2), This study
Hypoxylon laschii MUCL 52796 JX658525 ON954147 OP251027 ON813075 France Stadler et al. (2014; ITS), This study
Hypoxylon lateripigmentum MUCL 53304 KC968933 KY610486 KY624304 KC977290 Martinique (T) Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon lechatii MUCL 54609 KF923407 ON954148 OP251033 KF923405 French Guiana Kuhnert et al. (2014b; ITS, tub2), This study
Hypoxylon lenormandii CBS 119003 KC968943 KY610452 KY624261 KC977273 Ecuador Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon lienhwacheense MFLUCC 14-1231 KU604558 MK287550 MK287563 KU159522 Thailand Sir et al. (2016; ITS, tub2), Sir et al. (2019; LSU, rpb2)
Hypoxylon lividipigmentum STMA14045 ON792788 ON954149 ON813077 Argentina This study
Hypoxylon lividipigmentum BCRC 34077 JN979433 AY951735 Mexico (IT) Hsieh et al. (2005)
Hypoxylon macrocarpum CBS119012 ON792785 ON954151 OP251034 ON813071 Germany This study
Hypoxylon munkii MUCL 53315 KC968912 ON954153 OP251035 KC977294 Martinique Kuhnert et al. (2014a; ITS, tub2), This study
Hypoxylon musceum MUCL 53765 KC968926 KY610488 KY624306 KC977280 Guadeloupe Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon ochraceum MUCL 54625 KC968937 KY624271 KC977300 Martinique (ET) Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; rpb2)
Hypoxylon olivaceopigmentum DSM 107924 MK287530 MK287542 MK287555 MK287568 USA (T) Sir et al. (2019)
Hypoxylon perforatum CBS115281 KY610391 KY610455 KY624224 KX271250 France Wendt et al. (2018)
Hypoxylon petriniae CBS 114746 KY610405 KY610491 KY624279 KX271274 France (T) Wendt et al. (2018)
Hypoxylon pilgerianum STMA 13455 KY610412 KY610412 KY624308 KY624315 Martinique Wendt et al. (2018)
Hypoxylon porphyreum CBS 119022 KC968921 KY610456 KY624225 KC977264 France Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon pseudofuscum DSM112038 MW367857 MW367848 MW373858 MW373867 Germany (T) Lambert et al. (2021)
Hypoxylon pulicicidum CBS 122622 JX183075 KY610492 KY624280 JX183072 Martinique (T) Bills et al. (2012; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon rickii MUCL 53309 KC968932 KY610416 KY624281 KC977288 Martinique (ET) Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon rubiginosum MUCL 52887 KC477232 KY610469 KY624266 KY624311 Germany (ET) Stadler et al. (2013; ITS), Wendt et al. (2018; tub2, LSU, rpb2)
Hypoxylon samuelsii MUCL 51843 KC968916 KY610466 KY624269 KC977286 Guadeloupe (ET) Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon sporistriatatunicum MN056426 ON954150 OP251036 MK908140 Panama (T) Cedeño-Sanchez et al. (2020; ITS, tub2); This study
Hypoxylon subticinense MUCL 53752 KC968913 ON954152 KC977297 French Guiana Kuhnert et al. (2014a; ITS, tub2), This study
Hypoxylon texense DSM 107933 MK287536 MK287548 MK287561 MK287574 USA (T) Sir et al. (2019)
Hypoxylon ticinense CBS 115271 JQ009317 KY610471 KY624272 AY951757 France Hsieh et al. (2005; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon trugodes MUCL 54794 KF234422 KY610493 KY624282 KF300548 Sri Lanka (ET) Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon vogesiacum CBS 115273 KC968920 KY610417 KY624283 KX271275 France Kuhnert et al. (2014a; ITS), Kuhnert et al. (2017a; tub2), Wendt et al. (2018; LSU, rpb2)
Hypoxylon wuzhishanense FCATAS2708 OL467292 OL615104 OL584220 OL584227 China (T) Ma et al. (2022)
Jackrogersella cohaerens CBS 119126 KY610396 KY610497 KY624270 KY624314 Germany Wendt et al. (2018)
Jackrogersella multiformis CBS 119016 KC477234 KY610473 KY624290 KX271262 Germany (ET) Kuhnert et al. (2014a ; ITS), Kuhnert et al. (2017a; tub2), Wendt et al. (2018; LSU, rpb2)
Natonodosa speciosa CLM-RV86 MF380435 MF380435 MH745150 Mexico (T) Heredia et al. (2020)
Parahypoxylon papillatum comb. nov. ATCC 58729 KC968919 KY610454 KY624223 KC977258 USA (T) Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Parahypoxylon ruwenzoriense sp. nov. MUCL51392 ON792786 ON954156 OP251039 ON813078 D. R. Congo (T) This study
Pyrenopolyporus hunteri MUCL 52673 KY610421 KY610472 KY624309 KU159530 Ivory Coast (ET) Kuhnert et al. (2017a; tub2), Wendt et al. (2018; ITS, LSU, rpb2)
Pyrenopolyporus laminosus MUCL 53305 KC968934 KY610485 KY624303 KC977292 Martinique (T) Kuhnert et al. (2014a; ITS, tub2), Wendt et al. (2018; LSU, rpb2)
Pyrenopolyporus nicaraguense CBS 117739 AM749922 KY610489 KY624307 KC977272 Burkina_Faso Bitzer et al. (2008; ITS), Kuhnert et al. (2014a; tub2), Wendt et al. (2018; LSU, rpb2)
Rhopalostroma angolense CBS 126414 KY610420 KY610459 KY624228 KX271277 Ivory Coast Wendt et al. (2018)
Rostrohypoxylon terebratum CBS 119137 DQ631943 DQ840069 DQ631954 DQ840097 Thailand (T) Tang et al. (2007), Fournier et al. (2010b)
Ruwenzoria pseudoannulata MUCL 51394 KY610406 KY610494 KY624286 KX271278 D. R. Congo (T) Wendt et al. (2018)
Thamnomyces dendroidea CBS 123578 FN428831 KY610467 KY624232 KY624313 French Guiana (T) Stadler et al. (2010; ITS), Wendt et al. (2018; tub2, LSU, rpb2)
Xylaria arbuscula CBS 126415 KY610394 KY610463 KY624287 KX271257 Germany Fournier et al. (2011; ITS), Wendt et al. (2018; tub2, LSU, rpb2)
Xylaria hypoxylon CBS 122620 KY610407 KY610495 KY624231 KX271279 Sweden (ET) Wendt et al. (2018)

UHPLC profiling and dereplication

The secondary metabolites were extracted using a small piece of the stromata (approx. 1 mm3). Each piece was placed in 1.5 ml reaction tubes, covered with 1000 µl of methanol and placed for 30 min at 40 °C in an ultrasonic bath. The tubes were centrifuged at 14 000 rpm for 10 min. The methanol extract was separated from the remaining stromata, which was extracted again under the same procedure. Finally, both organic phases were combined and dried under nitrogen. Each sample was analyzed at a concentration of 450 µg/mL on an ultrahigh performance liquid chromatography system (Dionex Ultimate3000RS, Thermo Scientific, Dreieich, Germany), using a C18 column (Kinetex 1.7 µm, 2.1 × 150 mm, 100 Å; Phenomenex, Aschaffenburg, Germany) with a sample injection volume of 2 µL. The mobile phase consisted of A (H2O + 0.1% formic acid) and B (ACN + 0.1% formic acid) with a constant flow rate of 0.3 mL/min. The gradient began with 1% B for 0.5 min, increasing to 5% B in 1 min, then to 100% B in 19 min and holding at 100% B for 5 min. The temperature of the column was kept at 40 °C and UV-Vis data were recorded with a DAD at 190–600 nm.

MS spectra were collected using a trapped ion mobility quadrupole time-of-flight mass spectrometer (timsTOF Pro, Bruker Daltonics, Bremen, Germany) with the following parameters: tims ramp time 100 ms, spectra rate 9.52 Hz, PASEF on, cycle time 320 ms, MS/MS scans 2, scan range (m/z, 100–1800 Da; 1/k0, 0.55–2.0 V∙s/cm2). For the stromatal extracts and the standards ESI mass spectra were acquired in positive ion mode. Raw data were pre-processed with MetaboScape 2022 (Bruker Daltonics, Bremen, Germany) in the retention time range of 0.5 to 25 min. The obtained features were dereplicated against our in-house database comprising MS/MS spectra of standards from characteristic secondary metabolites of hypoxylaceous species (e.g. azaphilones, asterriquinones, binaphthalenes, cytochalasins, macrolides and sesquiterpenoids) in MetaboScape. A molecular network was created with the Feature-Based Molecular Networking (FBMN) (Nothias et al. 2020) and the Spec2Vec (Huber et al. 2021) workflows on the GNPS platform (Wang et al. 2016) using the pre-processed feature table from MetaboScape. Fragmentation ions resulting from the MS/MS spectra of cohaerin E, cohaerin H, and minutellin A were assigned using CFM-ID 4.0 web server (Wang et al. 2021) and validated with the SmartFormula 3D tool from MetaboScape. The datasets generated/analyzed for this study are included in Suppl. material 1.


Phylogenetic analyses

The final data matrix for the molecular phylogenetic analysis (Fig. 2) comprised 345 sequences (44 generated in this study, and complemented by sequences available from GenBank, NCBI) derived from 89 strains and four loci, namely ITS, LSU, rpb2 and tub2. The final MAFFT alignments consisted of 4018 nucleotides for the ITS alignment, 3642 for the LSU alignment, 2238 for the tub2 alignment and 4023 positions for the rpb2 alignment. The alignment of each locus is available in the Suppl. material 1: table S3–S6. Sequences of representatives for each molecularly well-established genus of the Hypoxylaceae were included: Annulohypoxylon (3 strains), Daldinia (8 strains), Durotheca (5 strains), Hypomontagnella (3 strains), Hypoxylon (58 strains), Jackrogersella (2 strains), Natonodosa (1 strain), Pyrenopolyporus (3 strains), as well as Rhopalostroma, Rostrohypoxylon, Ruwenzoria, and Thamnomyces (1 strain each). Three members of Xylariaceae and Graphostromataceae (Xylaria hypoxylon, X. arbuscula and Graphostroma platystomum) served as outgroup.

Figure 2. 

Inferred molecular phylogenetic maximum Likelihood (lLn = -122825.7921) tree of selected Hypoxylaceae, Graphostromataceae and Xylariaceae sequences. The analysis was calculated by using IQ-Tree with posterior probability support calculated from Bayesian inference methodology and support values generated from 1000 bootstrap replicates using a multigene alignment (ITS, LSU, tub2 and rpb2). The tree was rooted with Xylaria hypoxylon CBS 122620, X. arbuscula CBS 126415 (Xylariaceae) and Graphostroma platystomum CBS 27087 (Graphostromataceae). Type material is highlighted in bold letters. Bayesian posterior probability scores ≥ 0.95 / Bootstrap support values ≥ 70 are indicated along branches.

The inference of phylogenetic relationship using a Maximum-Likelihood and Bayesian approach yielded two different, discongruent topologies. An approximate unbiased (AU) topology test implemented in IQTree indicated that the tree resulting from Bayesian inference received a significantly (p < 0.05) lower maximum likelihood score, suggesting its rejection. Hence, we included support values of the approximate Bayes test implemented in IQTree to access posterior probability support values of the inferred phylogenetic tree. The combined rooted phylogenetic tree showed a clade consisting of the core members of the Hypoxylaceae, such as Hypoxylon, Daldinia, Pyrenopolyporus, Hypomontagnella, Jackrogersella, Rostrohypoxylon, Thamnomyces and Ruwenzoria with medium BS and high PP support (1/90), which was placed in a sister position to a clade consisting of members of Parahypoxylon gen. nov., and Durotheca (Hypoxylaceae) at the base of the tree with strong support (1/100). The genus Hypoxylon could be confirmed as paraphyletic, as has been described already by Wendt et al. (2018), Lambert et al. (2019), and Becker et al. (2020). The sequences assigned to Parahypoxylon ruwenzoriense formed a highly supported (1/100) cluster with the sequences derived from Parahypoxylon papillatum. The topology of Durotheca and the newly described genus Parahypoxylon as a basal lineage in the Hypoxylaceae are further reflected upon in the taxonomic part of this study.


Lecto- and epitypification

Hypomontagnella monticulosa (Mont.) Sir, L. Wendt & C. Lamb.


French Guiana, Cayenne, Leprieur, C. 1176, dead wood (PC, holotype; FH, isotype of H. monticulosum).


(designated here). France. French Guyana, Sinnamary, Paracou, Amazonian rain forest, bark of unknown tree, June 2012, leg J. Fournier (LIP, ex-epitype culture MUCL 54604). GenBank acc. nos for DNA sequences: KY610404 and KJ810556 (ITS), KY610487 (LSU), KY624305 (rpb2), KX271273 (tub2); MT889334 (sporothriolide gene cluster published by Tian et al. 2020).

MBT no: 10010042.


The strain designated here as epitype was used by Lambert et al. (2019) and the subsequent publications on genome analysis (Stadler et al. 2020; Tian et al. 2020; Kuhnert et al. 2021; Wibberg et al. 2021). The specimen and culture are perfectly suitable, because it was collected from the same geographic area as the holotype.

Parahypoxylon M. Cedeño-Sanchez, E. Charria-Girón & M. Stadler, gen. nov.

MycoBank No: 845463


Refers to the morphological similarity to Hypoxylon, from which the genus is phylogenetically distinct.


Differs from the genus Durotheca by the presence of greenish KOH-extractable pigments and by having an amyloid ascal apical apparatus. Differs from the genus Hypoxylon by containing yet unknown cohaerin-type azaphilones and by its basal position in the molecular phylogenetic inference using am ITS, LSU, rpb2 and tub2 matrix.

Parahypoxylon papillatum (Ellis & Everh) M. Cedeño-Sanchez, E. Charria-Girón & M. Stadler, comb. nov.

MycoBank No: 845462
Figs 3, 4

Hypoxylon papillatum Ellis & Everh. in Smith, Bull. Lab. Nat. Hist. Iowa State Univ. 2: 408 (1893). Syn.


USA. Ohio, Delaware, 21 Jul 1893, A. Commons 2160, rotten wood of Carya (NY [2 pks.], selected by Ju and Rogers (1996).


USA. West Virginia, Mason Co., Bruce’s Chapel, 18 Aug 1983, wood of Acer, J.D. Rogers (WSP 7557; ex-epitype culture ATCC 58729).

MBT no: 10011515.


Stromata superficial, effused-pulvinate to plane, with inconspicuous to conspicuous perithecial mounds, up to 12.5 cm long × up to 4 cm broad × 1.8–4.0 mm thick; surface Honey (64) to Isabelline (65), Isabelline (65) to Gray Olivaceous (107), or Isabelline (65) to Olivaceous (48); blackish granules immediately beneath surface and between perithecia, with KOH-extractable pigments Isabelline (65); the tissue below the perithecial layer conspicuous, black, 1.0–2.5 mm thick. Perithecia long-tubular, 0.3–0.4 mm diam × 0.8–1.5 mm high. Ostioles umbilicate. Asci with amyloid, discoid apical apparatus, 1–2 µm high × 3.5 µm wide, stipe up 137–180 µm long × 8–10 µm broad, the spore-bearing parts 93–110 µm long, the stipes 30–80 µm long. Ascospores brown to dark brown, unicellular, ellipsoid, nearly equilateral, with broadly to narrowly rounded ends, 12.0–18.5 × 6.5–9.0 µm, with straight germ slit spore-length; perispore indehiscent in 10% KOH; epispore smooth.

Figure 3. 

Parahypoxylon papillatum comb. nov. A stroma B ostioles C KOH extractable stromatal pigments D perithecia (cross section) E ascospores with straight germ slits F amyloid apical apparatus in a mature ascus treated with Melzer’s reagent G amyloid apical apparatus in an immature ascus treated with Melzer’s reagent. Scale bars: 1 cm (A); 10 μm (E–F); 10 μm (G).

Figure 4. 

Parahypoxylon papillatum comb. nov. (ATCC 58729) Colonies after 2 weeks (A, B) on 2% MEA (C, D) on OA (E, F) on YM.

Cultures and anamorph

Colonies on MEA, OA, and YM covering a 9 cm Petri plate in 2 weeks, with white, flat, mycelium, margins filamentous. Reverse at first white, becoming yellowish at the center. The anamorph has been described by Rogers (1985), but we were unable to confirm the presence of conidial structures when we studied the strain more than 30 years later.

Secondary metabolites

Stromata contain BNT and cohaerin type azaphilones according to the MS/MS analysis.


We were not only able to confirm the morphometric results of Ju and Rogers (1996) but even established that this species is characterized by a rather specific metabolite profile. This species has to our knowledge still not been reported from outside America and seems to be most frequently encountered in the Eastern USA.

Further specimens examined

USA. Kansas, on decorticated wood, Feb 1884, F.W. Cragin 257 (NY00830462, syntype of H. papillatum); Pennsylvania, Allegheny Co., on dead wood, 14 Aug 1941, Henry, L.K. 4885 (BPI 591033); Pennsylvania, Meadville, old log, 17 Oct 1922, E.C. Smith 353 (BPI 591030); CANADA., on wood, J. Dearness (BPI 591035A, syntype of H. papillatum).

Parahypoxylon ruwenzoriense M. Cedeño-Sanchez, E. Charria-Girón & M. Stadler, sp. nov.

MycoBank No: 845457
Figs 5, 6


Democratic Republic Of The Congo. North Kivu: Mt. Ruwenzori, about 00°33.961'N, 29°81.795'E, between 2,138 and 2,400 m alt., 3–5 Feb 2008, tropical mountain forest, C. Decock (MUCL 51392, ex-holotype culture MUCL 51392).


Named after the Ruwenzori Mountains, where the species was collected.


Stromata superficial, incomplete, effused-pulvinate, 60 mm long × 40 mm broad × 3–5 mm thick; surface Fawn (87), with inconspicuous perithecial mounds, with a black, shiny hard crust 100–150 µm thick above perithecia, without visible granules, with KOH-extractable pigments Hazel (88); the pruina hyphae turn violet in KOH; the tissue below the perithecia 2–4 mm thick, vertically fibrose, dark grey. Perithecia tubular, 0.90–1.50 mm high × 0.2–0.3 mm diam (n=18). Ostioles umbilicate, surrounded by a white substance. Asci cylindrical, 8-spored, the spore-bearing parts 82–105 µm long × 5.5–6.0 µm broad, the stipes 38–130 µm long, with amyloid, discoid apical ring 0.7–2.0 µm high × 2.5–3.5 µm (n=21) broad. Ascospores smooth, unicellular, brown to dark brown, narrowly ellipsoid, nearly equilateral with narrowly rounded ends, 10.5–13.8 × 4.0–5.6 µm (n=40), with a faint, straight germ slit; perispore indehiscent in 10% KOH.

Figure 5. 

Parahypoxylon ruwenzoriense sp. nov. (MUCL 51392). A stroma B ostioles with white ring C KOH extractable stromatal pigments D perithecia (cross section) E ascospores F amyloid apical apparatus (blueing in Melzer’s reagent) indicated by arrowheads G asci. Scale bars: 1 cm (A); 2 mm (D); 10 μm (E, F); 50 μm (G).

Figure 6. 

Parahypoxylon ruwenzoriense sp. nov. (MUCL 51392) Colonies after 2 weeks (A, B) on 2% MEA (C, D) on OA (E, F) on YM.

Cultures and anamorph

Colonies on MEA, OA, and YM covering a 9 cm Petri plate in 2 weeks, with mycelium white at first, flat to raised in some zones, to becoming greenish in the center. Reverse at first yellowish, to become orange with a black spot at the center. Conidiophores not produced.

Secondary metabolites

Stromata contain BNT and cohaerin type azaphilones according to the MS/MS analysis.


P. ruwenzoriense is phylogenetically close to P. papillatum but differs by its KOH-extractable pigments Hazel (88) and by smaller ascospores.

Metabolomic profiling of stromata

As explained in the Experimental section, stromata of five herbarium specimens assignable to Parahypoxylon were extracted and analysed by UHPLC-DAD-IM-MS/MS. The raw data sets were pre-processed and the obtained feature table dereplicated using high resolution m/z, MS/MS spectra, retention time, CCS value, and UV/Vis spectra and reference data obtained from our in-house library of common secondary metabolites of the Hypoxylaceae (data not shown).

From the base peak chromatograms (BPC) of the stromatal extracts of the studied specimens, six major peaks could be distinguished (Fig. 7). An additional MS/MS similarity search without matching the precursor mass against our in-house library in MetaboScape yielded a MS/MS score > 700 for compounds 2 and 5 when compared with cohaerin E, cohaerin H, and minutellin A standards, which were not contained in the stromatal extracts (Suppl. material 1: fig. S2). This tentatively advocated a structural relation to the azaphilone family (Fig. 8a). Molecular formulae for compounds 16 were predicted as C23H24O7, C23H22O7, C23H20O8, C23H22O6, C23H20O6, and C23H21NO5 (Suppl. material 1: table S7), with a lower number of carbons than cohaerin E (C28H30O6), cohaerin H (C28H32O6), and minutellin A (C28H30O7). To further validate the presence of cohaerin E-like azaphilones in the stromatal extracts of the Parahypoxylon spp. a molecular networking (MN) approach was pursued. The above mentioned tool can be employed to organize in an automatic basis MS/MS spectra into groups based on similarities in their fragmentation patterns and the hypothesis that structurally related molecules will yield similar MS/MS spectra (Duncan et al. 2015). For this analysis, we compared the MS/MS spectra of cohaerin E, cohaerin H, and minutellin A (Suppl. material 1: table S7, fig. S2) with all MS/MS spectra obtained from the Parahypoxylon gen. nov. stromatal extracts by means of the unsupervised machine learning approach Spec2Vec. As a result, the molecular cluster containing the cohaerin standards consisted of 29 consensus spectra (nodes), which included compounds 16 (Fig. 8b). In addition, cohaerin E and H have UV/Vis absorptions at λmax 226–223 and 344–380 nm, which are resembling UV/Vis absorptions from compounds 1, 3, 4, and 6. Minutellin A displayed UV/Vis absorptions at λmax 224, 271, and 343 nm, a pattern identified also for compounds 2 and 5 (Fig. 8c).

Figure 7. 

Base peak chromatograms (BPCs) from UHPLC-MS analysis of the stromatal extracts of P. papillatum (BPI 591029), P. papillatum (BPI 591035), P. papillatum (NYGB 830462), P. papillatum (NYGB 830463), and Parahypoxylon ruwenzoriense sp. nov. (STMA 08016). Compounds common between several species (numbered 16) are highlighted in red.

Figure 8. 

A Reference MS/MS spectra of cohaerin E, cohaerin H, and minutellin A standards, and the six major azaphilones identified in the UHPLC-MS chromatograms of stromatal extracts from the Parahypoxylon spp. B azaphilone cluster in a molecular network created from the Parahypoxylon spp. stromatal extracts and MS/MS spectra from selected standards C UV/Vis profile comparison from compound 16, cohaerin E, cohaerin H, and minutellin A.

Cohaerin-type azaphilones present as well a distinct MS fragmentation pattern. In MS/MS experiments, cohaerin E generated fragment ions at 393.207 Da, 323.092 Da, 281.085 Da, and 253.086 Da, while minutellin A generated fragment ions at 397.201 Da, 341.102 Da, 299.091 Da, and 271.097 Da. The most abundant fragments were annotated using the CFM-ID 4.0 peak assignment module. In both cases, the most abundant fragments were traced down to the azaphilone backbone (Fig. 9). For instance, the mass difference of 18 Da between 323.092 Da and 341.102 Da could be interpreted as H2O, reflecting the different substitution of the 3-methylphenol moiety. Fragment ions at 281.085 Da and 253.086 Da for cohaerin E represent the tricyclic portion of the molecule, while fragment ions at 299.091 Da and 271.097 Da represent the same part of the molecule in minutellin A. Analogously, MS fragmentation patterns for cohaerin H (Fig. 8) resembles the generated fragments for cohaerin E. As some typical cohaerin-type azaphilones fragmentation patterns were conserved, we assume that the changes found for the stromatal metabolites of 16 occur in the side chain of the molecules. In summary, the UHPLC-DAD-IM-MS/MS and UV/Vis data, combined with a comparison of molecular networking analyses, indicated the presence of novel azaphilones related to the cohaerin family in the stromatal extracts from the Parahypoxylon spp., in contrast to the absence of other common secondary metabolites of the Hypoxylaceae.

Figure 9. 

A most abundant fragment ions observed in MS/MS spectra for cohaerin E and associated structures as predicted by CFM-ID 4.0 B most abundant fragment ions observed in MS/MS spectrum for minutellin A and associated structures as predicted by CFM-ID 4.0.


The genus Hypoxylon in the current taxonomic concept has frequently been shown to be paraphyletic (Wendt et al. 2018; Lambert et al. 2019), which has once more been confirmed in this study, foreshadowing again future rearrangements for a thorough revision of its systematics. This is especially apparent because the type species H. fragiforme forms a relatively small clade clustering with a small subset of closely related taxa. Therefore, further segregation will eventually be unavoidable once more data to safely delineate the different lineages becomes available. Here, we gathered chemotaxonomic, morphological and sequence data to enable a polyphasic characterization of a basal clade formerly phylogenetically resolved inside Hypoxylon, containing specimen closely related to H. papillatum, for which we propose the erection of the new genus Parahypoxylon, sharing many salient features with Hypoxylon in the “traditional” definition.

The investigation of the stromatal metabolite extracts by HPLC has proven to be a valuable resource to achieve a more natural classification of hypoxylaceous taxa (Kuhnert et al. 2015; Wendt et al. 2018; Lambert et al. 2019). Recent advances in the analytics for in-depth characterization of natural products, mainly driven by metabolomics-based strategies, have enabled a better understanding of complex natural systems (Van der Hooft et al. 2020). The current MS-based techniques can help as a predictor for the discovery of new carbon skeletons to help and prioritize their isolation and description instead of the isolation of new derivatives of already known metabolite scaffolds. Nevertheless, relying mainly on MS/MS fragmentation spectra could lead to an underestimation of chemical diversity. The complex chemical space produced by a single BGC may result in completely different fragmentation patterns only by the addition of small structural changes (McCaughey et al. 2022). Still, a general methodology for characterizing and classifying structural analogs with a common biosynthetic origin is absent particularly in the field of fungal natural products (Almeida et al. 2020).

However, in many occasions and applications, the isolation and structure elucidation of yet unidentified compounds is not possible, such as in the example of isolating pigments from natural sources, as is the case in the genus Hypoxylon. Even very old specimens have been reported to harbor intact secondary metabolites, as has been described for fossilized stromata assigned to Hypoxylon fragiforme in a study of archeological samples by Surup et al. (2018). Here, fortunately the original species could be recollected in German woods, but for rarer specimens, or specimens only producing scarce amounts of stromata, this is not a practicable option. Instability of the contained metabolites during e.g. purification further complicates the issue (Stadler et al. 2008; Kuhnert et al. 2014b; Sir et al. 2019). In this study, we demonstrated the value of integrating metabolomics-based tools to characterize the secondary metabolite profile of the type and authentic specimens of P. papillatum and the new species from the D.R. Congo.

An MS/MS analysis of the major metabolites suggested the presence of six unknown compounds assignable to the azaphilones related to the cohaerin family, which have been predicted to harbor a smaller carbon skeleton than the known cohaerins, and which still conserve some of the distinctive fragmentation patterns of these secondary metabolites (Suppl. material 1: fig. S3). This phenomenon has been exemplified within the Hypoxylaceae, which present a highly diverse group of PKS-derived pigments, among which the different subfamilies present different attached side chains at the C-8 oxygen (Kuhnert et al. 2021). The above findings suggest that the type of azaphilone produced by the studied species belong to a different type of azaphilones with a shorter side chain, but with a shared backbone in comparison to the cohaerins and minutellins. Additionally, the number of nodes found in the MN analysis suggests that the chemical diversity of the azaphilones produced by the strains belonging to Parahypoxylon gen. nov. is much higher than thought. In general, following a similar approach, the MolNetEnhancer workflow allowed the characterization of triterpenoid metabolites with several distinct phenolic acid modifications (e.g., vanillate, protocatechuate) in a different taxonomic background in the plant family Rhamnaceae (Ernst et al. 2019). The same methodology enabled the annotation of molecular families with known chemical motifs previously unreported for Salinispora, Streptomyces, and Xenorhabdus bacterial extracts (Ernst et al. 2019). Even though the ideal scenario would remain to isolate and elucidate the structures of the secondary metabolites, these tools are a powerful resource to classify chemical structural annotation and enhance our understanding of chemodiversity by adding biological and chemical insights of complex metabolic mixtures. It is worth noting that the stromatal material could eventually become available in the future from forthcoming collection campaigns, and therefore the aforementioned hypothesis might be confirmed through isolation and chemical characterization of the major metabolites.

In this context, the stromatal metabolite profile of the specimens of P. papillatum and the new species P. ruwenzoriense are rather unique, even though it exhibits related chemotaxonomic features more likely to be found in the Hypoxylaceae. The cohaerin type azaphilones (which include also the multiformins and minutellins) have first been reported by Quang et al. (2005a, b, 2006), Surup et al. (2013) and Kuhnert et al. (2017b) and were recently found to possess interesting antiviral effects (Jansen-Olliges et al. 2023). Their producers are now all classified in Jackrogersella (Wendt et al. 2018) and were formerly placed in Hypoxylon sect. Annulata or (Ju and Rogers 1996) Annulohypoxylon (Hsieh et al. 2005), respectively. Kuhnert et al. (2017a) already reported that the species of Annulohypoxylon are divided into two chemotypes, one of which is characterized by stromata with papillate ostioles and cohaerin type azaphilones. The other chemotype is devoid of these compounds and produces binaphthalenes as prevailing stromatal metabolites. It includes A. truncatum, the type species of Annulohypoxylon, and many other species that feature ostiolar discs. Since this coincided with the molecular phylogeny by Wendt et al. (2018), the new genus Jackrogersella was erected for the cohaerin-containing species with papillate, diskless ostioles. There is only one species in Annulohypoxylon (i.e., A. michelianum) that has such ostiolar rings and also produces cohaerins. It was left at interim in Annulohypoxylon, even though its DNA sequence occupied a separate clade in the phylogeny by Wendt et al. (2018). The reason is that the strain studied did not constitute type material, and we felt that the erection of a separate genus should only be carried out by including fresh material from the geographic area and host (Laurus in South Europe) from which the holotype specimen was reported. Aside from the above-mentioned fungi, metabolites with cohaerin-like characteristics (i.e. characteristic mass and diode array spectra) have even been detected in species of Hypoxylon, such as H. pulicicidum (Bills et al. 2012). A recent study based on the analysis of full genomes based on 3rd generation sequencing techniques, such as PacBio and Oxford nanopore (Wibberg et al. 2021), has even revealed the corresponding biosynthetic gene clusters encoding for these azaphilone pigments to be present in the studied Jackrogersella species and H. pulicicidum (Kuhnert et al. 2021). For instance, the identified BGC in H. pulicicidum carries the core set of conserved genes for this family of azaphilones, but the additional presence of additional tailoring enzymes indicates that the produced metabolites might have different structural features compared to the known cohaerins (Kuhnert et al. 2021).

In the future, it will become easier to tell if the genetic information for the successful biosynthesis of such secondary metabolites is present in the genomes of the respective organisms even if the products cannot be detected. Chemotaxonomic evidence can also be used to segregate the new genus from the species that are located in neighboring basal clades in the current phylogeny (i.e., Hypoxylon aeruginosum and Durotheca spp.). Interestingly, these species neither contain azaphilones nor binaphthalenes, with H. aeruginosum and the related genus Chlorostroma reported to have lepraric acid derivatives as major stromatal metabolites (Læssøe et al. 2010), which are otherwise unique and only occur in some lichenized ascomycetes. Durotheca, on the other hand, appears to be poor in stromatal metabolites, and Læssøe et al. (2013) only found traces of lepraric acids in one of the species they studied. The current study has further confirmed the results by de Long et al. (2019), who found that Durotheca is a hypoxylaceous genus, even though its species have a distinctive ascospore morphology and other secondary metabolites.

The integration of state-of-the-art metabolomic-based tools in chemotaxonomic surveys will further accelerate and assist the systematic study of paraphyletic taxa within the concept of polyphasic taxonomy as herein demonstrated for the introduction of Parahypoxylon.


This work was funded by the DFG (Deutsche Forschungsgemeinschaft) priority program “Taxon-Omics: New Approaches for Discovering and Naming Biodiversity” (SPP 1991). It also benefited from the European Union’s H2020 Research and Innovation Staff Exchange program (RISE) [Grant No. 101008129: MYCOBIOMICS], granted to J.J. Luangsa-ard and M. Stadler. MCS gratefully acknowledges a PhD stipend from The National Secretariat of Science, Technology and Innovation of the Republic of Panama (SENACYT) and the Institute for the Development of Human Resources (IFARHU). E. Charria-Girón was funded by the HZI POF IV Cooperativity and Creativity Project Call. Additionally, we gratefully acknowledge support from the curators of the international herbaria, above all Lisa A Castlebury (BPI), Laura Briscoe (NY) and Monique Slipher (WSP), who provided important specimens for this study. Additionally, we are grateful for the expert and technical advice from Ulrike Beutling and Frank Surup.


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Supplementary material

Supplementary material 1 

Supplementary information

Marjorie Cedeño-Sanchez, Esteban Charria-Girón, Christopher Lambert, J. Jennifer Luangsa-ard, Cony Decock, Raimo Franke, Mark Brönstrup, Marc Stadler

Data type: Alignments and MS raw data (PDF file)

This dataset is made available under the Open Database License ( The Open Database License (ODbL) is a license agreement intended to allow users to freely share, modify, and use this Dataset while maintaining this same freedom for others, provided that the original source and author(s) are credited.
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