Research Article
Research Article
New species and new combinations in the genus Paraisaria (Hypocreales, Ophiocordycipitaceae) from the U.S.A., supported by polyphasic analysis
expand article infoRichard M. Tehan§, Connor B. Dooley, Edward G. Barge|, Kerry L. McPhail, Joseph W. Spatafora
‡ Oregon State University, Corvallis, United States of America
§ Utica University, Utica, United States of America
| Seed Testing Laboratory, Idaho State Department of Agriculture, Boise, United States of America
Open Access


Molecular phylogenetic and chemical analyses, and morphological characterization of collections of North American Paraisaria specimens support the description of two new species and two new combinations for known species. P. cascadensis sp. nov. is a pathogen of Cyphoderris (Orthoptera) from the Pacific Northwest USA and P. pseudoheteropoda sp. nov. is a pathogen of cicadae (Hemiptera) from the Southeast USA. New combinations are made for Ophiocordyceps insignis and O. monticola based on morphological, ecological, and chemical study. A new cyclopeptide family proved indispensable in providing chemotaxonomic markers for resolving species in degraded herbarium specimens for which DNA sequencing is intractable. This approach enabled the critical linkage of a 142-year-old type specimen to a phylogenetic clade. The diversity of Paraisaria in North America and the utility of chemotaxonomy for the genus are discussed.

Key words

Ascomycota, chemotaxonomy, Cicada, Cordyceps, Cyphoderris, entomopathogen, Ophiocordyceps, Prionus


Paraisaria is an asexual morph-typified genus of entomopathogenic fungi, originally described by Samson and Brady in 1983, characterized by synnemata with verticillately-branched conidiophores and flask-shaped sympodially proliferating phialides (Samson and Brady 1983). These asexual morphs were derived from larvae (Delacroix 1893) and from cultured isolates of the sexual morphs of species in the genus Cordyceps (Samson and Brady 1983; Li et al. 2004), which were later transferred to Ophiocordyceps (Sung et al. 2007). Paraisaria was later proposed for suppression, along with four other genera then in use, in favor of recognizing a broad concept of Ophiocordyceps (Quandt et al. 2014). This limited the number of new combinations required to accommodate 1F1N rules following the abolition of the dual system of nomenclature in which sexual states and asexual states of fungi were classified separately. In molecular analyses, Paraisaria has been recovered as a distinct monophyletic clade, being referred to as the “gracilis subclade” within the ravenelii subclade” of Ophiocordyceps by Sanjuan et al. (2015). Paraisaria was ultimately resurrected in 2019, segregated from Ophiocordyceps, and amended to include sexual morphology (Mongkolsamrit et al. 2019). Paraisaria species possess distinctive sexual morphs characterized by a globose fertile terminal portion of the stroma with immersed perithecia. Thus, Paraisaria constitutes a distinct, and robustly monophyletic clade deserving a unique genus classification, though its segregation from Ophiocordyceps rendered Ophiocordyceps into several paraphyletic clades. Ultimately, a comprehensive analysis of Ophiocordyceps sensu Sung et al. (2007), is needed to establish robust generic concepts and restore global monophyly. A major sticking point for this action is the uncertain placement of the type of Ophiocordyceps, O. blattae, among the paraphyletic subclades of Ophiocordyceps.

In North America, Paraisaria species are unique among most Cordyceps sensu lato in that they form fruiting bodies in the spring, whereas most other insect pathogens fruit in the summer, fall, or winter months, which is evident in herbarium records on MycoPortal (MycoPortal 2023) and observations on the community science platform iNaturalist ( Most Paraisaria species, and thus far, all known Paraisaria species occurring in North America, form fruiting bodies on subterranean insect hosts.

Some of the insect hosts of Paraisaria species are sought as food and their contamination by Paraisaria species could pose a human health concern. Doan et al. (2017) reported a series of poisonings and one fatality in Southern Vietnam, among people who had consumed cicadae infected with a fungus identified as Paraisaria heteropoda (=Cordyceps heteropoda, Ophiocordyceps heteropoda), between 2008 and 2015. The toxicity was attributed to the presence of mycotoxins in the otherwise edible cicadae, and the toxic agent was putatively identified as ibotenic acid. The potential role of entomopathogenic fungi in causing food-borne mycotoxin poisonings underscores the need to describe the biological and chemical diversity present in this group of fungi.

In addition to their impact on human and animal health, fungal natural products can be highly useful phenotypic characters for taxonomic purposes. Chemical fingerprints can be used to identify chemical families that constitute a generic chemotype for a taxonomic group, and also unique suites of compounds within a chemical family can be used to resolve species. For example, Cedeño-Sanchez et al. (2023) profiled chemical extracts from stromata to characterize and distinguish species and genera in the family Hypoxylaceae.

Only two studies (Krasnoff et al. 2005; Umeyama et al. 2011) have reported a total of five natural products from Paraisaria species, both of which investigated fungi identified as the cicada pathogen, Paraisaria heteropoda. A third study reports leucinostatin analogs from an organism reported as Ophiocordyceps heteropoda (=Parasiara heteropoda) (Kil et al. 2020), but which is evidently a Purpureocillium species based on ITS phylogeny and chemotaxonomy. Doan et al. (2017) also report the amino acid, ibotenic acid from this species, but no analytical chemistry data are presented to confirm this. There are currently no published genome sequences available to mine the specialized metabolic potential of Paraisaria species, although the sequenced genomes of other Ophiocordycipitaceae species display a familial trend of high biosynthetic capacity for specialized metabolites. The first chemical study of a member of this genus resulted in the discovery of the new 8-residue antimicrobial peptaibiotics, cicadapeptins I and II, which possessed a unique two consecutive 4-hydroxyproline residues at the N-terminus (Krasnoff et al. 2005). The known antifungal and immunosuppressant sphingosine analog, myriocin was also isolated in this study. Heteropodamides A and B are N-methylated cyclic heptapeptides reported as cytotoxins from P. heteropoda (Umeyama et al. 2011). Their absolute structures are yet to be determined. The further discovery of Paraisaria species and their natural products presents fertile grounds for investigation.

In the course of ongoing investigations for the discovery of biologically active natural products from Paraisaria species (Tehan 2022), it became critical to perform a taxonomic analysis of North American Paraisaria to better understand the biological diversity present in this group. In this study, we examined 29 recent collections of Paraisaria to investigate the diversity of North American Paraisaria. We also analyzed the type collections of Ophiocordyceps insignis and O. monticola, both of which were anticipated to belong in Paraisaria based on morphological description, ecology, and phenology. One phylogenetically informative DNA sequence was afforded from the 87-year old O. monticola specimen. The 142-year old O. insignis type did not permit successful DNA sequencing, however, chemical analysis of the newly characterized paraisariamide family of compounds by LC-HRMS provided robust support for the combination of both species into Paraisaria, as well as the correct identification of a species of importance to human health, as P. insignis. This study provides a novel framework for the use of minimally destructive chemical analysis in taxonomic assessment of type specimens where DNA sequencing is not possible. The combined analysis of molecular data, morphology, ecology, phenology, and chemical data support the circumscription of two new species and two new combinations, and provides an initial overview of the diversity of American Paraisaria species.

Materials and methods

Specimens and isolates

Twenty nine new collections of Paraisaria specimens and their insect hosts were examined. Macroscopic characters were examined from fresh stromata, and microscopic characters were examined from fresh and dried stromata, including ascospores discharged from fresh stromata when possible and sections of dried specimens. Colors are in general terms of the senior author. Specimens are deposited in the Oregon State University Herbarium mycological collection. Culture isolates of fungi were made from tissue dissected from the context of stromata, placed on PDA with 50 µg/ml ampicillin and 100 µg/ml streptomycin, or from ascospores germinated on PDA. Agar plugs were taken from outgrowth of stromatic tissue and subcultured onto PDA and CMA at 20 °C. Cultures are deposited at the USDA ARS Collection of Entomopathogenic Fungal Cultures (ARSEF).

Morphological observations

Fruiting bodies were examined for morphological measurements using a Vernier caliper (Fowler). Sections of ascogenous tissue were mounted in lactophenol cotton blue, 5% KOH, or distilled water, and microanatomical characters were examined with light microscopy using a Leica DM2500. Twenty each, perithecia, asci, and part-spores were measured at magnifications of 10×, 20×, 40×, 63×, or 100×.

DNA extraction and sequencing

DNA was extracted from the ascogenous portion of dried stromata, ground with mortar and pestle in CTAB buffer (1.4 M NaCl, 100 mM Tris–HCl pH 8.0, 20 mM EDTA pH 8.0, 2% CTAB w/v) and processed following the method of Kepler et al. (2012). Samples were extracted with 25:24:1 phenol:chloroform:isoamyl alcohol, (affymetrix), and DNA was precipitated with 3 M sodium acetate (pH 5.2) and 95% ethanol. PCR amplification was performed on the Internal Transcribed Spacer (ITS), amplified using ITS4 and ITS5 primers (White et al. 1990). Alternatively, ITS1F (Gardes and Bruns 1993) was used as a forward primer for samples where ITS4 did not work. For samples in which amplification of the ITS region did not succeed, individual amplification of the ITS1 and ITS2 loci was attempted using primer sets ITS5 and ITS2 (White et al. 1990) for the ITS1 locus, and ITS3 (White et al. 1990) and ITS4 for the ITS2 locus. Nuclear small subunit (nucSSU) was amplified using nucSSU131 and NS24 (Kauff and Lutzoni 2002), nuclear large subunit (nucLSU) using LROR (Rehner and Samuels 1994) and LR7 (Vilgalys and Hester 1990), subunit 1 of RNA polymerase II (RPB1) using RPB1-Af and RPB1-6R1asc (Hofstetter et al. 2007). Alternatively, CRPB-1 (Castlebury et al. 2004) was used as a forward primer for samples where RPB1-Af did not work. Elongation factor 1α (EF-1α) was amplified using 983F and 2218R (Castlebury et al. 2004). PCR was performed with an iCycler (Bio- Rad, USA), with a total of 20 μl reaction mixture containing 1× PCR Buffer (Promega), 1× TBTpar prepared as in Samarakoon et al. (2013), 2.5 mM MgCl2, 0.5 µM each forward and reverse primers, 200 µM of each of the four dNTPs, and 0.5 U Taq polymerase. For ITS, SSU, LSU, and TEF, the PCR thermal cycle consisted of an initial 1 min denaturation at 95 °C; 34 cycles of 30 s at 94 °C, 1 min at 52 °C, 1.5 min at 72 °C, and a termination with an elongation 7 min at 72 °C. For RPB1 and RPB2, the PCR thermal cycle consisted of an initial 1.5 min denaturation at 95 °C; 39 cycles of 30 s at 94 °C, 1 min at 47 °C, 2 min at 72 °C, and a termination with an elongation 4 min at 72 °C. Sequencing was performed by the Sanger method at the Center for Quantitative Life Sciences at Oregon State University. The sequences obtained in this study were deposited to GenBank (Table 1).

Table 1.

Sequences used in phylogenetic tree construction.

Species Code Host ITS SSU LSU EF1a RPB1 RPB2 Reference
Cordyceps kyushuensis EFCC 5886 Lepidoptera EF468960 EF468813 EF468754 EF468863 EF468917 Sung et al. 2007
Cordyceps militaris OSC.93623 Lepidoptera JN049825 AY184977 AY184966 DQ522332 DQ522377 Kepler et al. 2017
Drechmeria balanoides CBS 250.82 Nematoda MH861495 AF339588 AF339539 DQ522342 DQ522388 DQ522442 Vu et al. 2019
Drechmeria sinensis CBS 567.95 Nematoda MH862540 AF339594 AF339545 DQ522343 DQ522389 DQ522443 Spatafora et al. 2007
Harposporium anguillulae ARSEF 5407 Nematoda AY636080 Chaverri et al. 2005
Harposporium helicoides ARSEF 5354 Nematoda AF339577 AF339527 Sung et al. 2001
Ophiocordyceps australis HUA186147 Hymenoptera KC610784 KC610764 KC610734 KF658678 Sanjuan et al. 2015
Ophiocordyceps australis HUA186097 Hymenoptera KC610786 KC610765 KC610735 KF658662 Sanjuan et al. 2015
Ophiocordyceps curculionum OSC 151910 Coleoptera KJ878918 KJ878885 KJ878999 Quandt et al. 2014
Ophiocordyceps irangiensis NBRC101400 Hymenoptera JN943335 JN941714 JN941426 JN992449 Schoch et al. 2012
Ophiocordyceps kimflemingiae SC30 Hymenoptera KX713629 KX713622 KX713699 KX713727 Araújo et al. 2018
Ophiocordyceps konnoana EFCC 7315 Coleoptera EF468959 EF468753 EF468861 EF468916 Mongkolsamrit et al. 2019
Ophiocordyceps longissima TNS F18448 Hemiptera KJ878925 KJ878892 KJ878971 KJ879005 Quandt et al. 2014
Ophiocordyceps melolonthae OSC.110993 Coleoptera DQ522548 DQ518762 DQ522331 DQ522376 Mongkolsamrit et al. 2019
Ophiocordyceps monticola BPI 634610 Orthoptera OQ709246 This Study
Ophiocordyceps nigrella EFCC 9247 Coleoptera JN049853 EF468963 EF468818 EF468758 EF468866 EF468920 Mongkolsamrit et al. 2019
Ophiocordyceps nutans OSC 110994 Hemiptera DQ522549 DQ518763 DQ522333 DQ522378 Quandt et al. 2014
Ophiocordyceps pulvinata TNS-F 30044 Hymenoptera GU904208 GU904209 GU904210 Kepler et al. 2011
Ophiocordyceps ravenelii OSC 151914 Coleoptera KJ878932 KJ878978 KJ879012 KJ878950 Quandt et al. 2014
Ophiocordyceps sinensis EFCC 7287 Lepidoptera JN049854 EF468971 EF468827 EF468767 EF468874 EF468924 Quandt et al. 2014
Ophiocordyceps stylophora OSC_111000 Coleoptera JN049828 DQ522552 DQ518766 DQ522337 DQ522382 DQ522433 Quandt et al. 2014
Ophiocordyceps variabilis OSC 111003 Diptera EF468985 EF468839 EF468779 EF468885 EF468933 Mongkolsamrit et al. 2019
Ophiocordyceps variabilis ARSEF 5365 Diptera DQ522555 DQ518769 DQ522340 DQ522386 DQ522437 Mongkolsamrit et al. 2019
Paraisaria alba HKAS_102484 Orthoptera MN947219 MN943843 MN943839 MN929085 MN929078 MN929082 Wei et al. 2021
Paraisaria amazonica HUA 186143 Orthoptera KJ917562 KJ917571 KM411989 KP212902 KM411982 Sanjuan et al. 2015
Paraisaria amazonica HUA 186113 Orthoptera KJ917566 KJ917572 KP212903 KM411980 Sanjuan et al. 2015
Paraisaria arcta HKAS_102553 Lepidoptera MN947221 MN943845 MN943841 MN929087 MN929080 Wei et al. 2021
Paraisaria arcta HKAS 102552 Lepidoptera MN947220 MN943844 MN943840 MN929086 MN929079 MN929083 Wei et al. 2021
Paraisaria blattarioides HUA186093 Blattodea KJ917559 KJ917570 KM411992 KP212910 Sanjuan et al. 2015
Paraisaria blattarioides HUA 186108 Blattodea KJ917558 KJ917569 KP212912 KM411984 Sanjuan et al. 2015
Paraisaria cascadensis OSC-M-052010 Orthoptera OQ709237 OQ800918 OQ708931 OR199814 OR199828 OR199838 This Study
Paraisaria cascadensis OSC-M-052012 Orthoptera OQ709239 OQ800920 OQ708933 OR199816 OR199830 This Study
Paraisaria cascadensis OSC-M-052017 Orthoptera OQ709240 OQ800921 OQ708934 OR199817 OR199831 This Study
Paraisaria coenomyia NBRC 106964 Diptera AB968397 AB968385 AB968413 AB968571 AB968533 Ban et al. 2015
Paraisaria coenomyia NBRC 108993 Diptera AB968396 AB968384 AB968412 AB968570 AB968532 Ban et al. 2015
Paraisaria gracilioides HUA186095 Coleoptera KJ917556 KM411994 KP212914 Sanjuan et al. 2015
Paraisaria gracilioides HUA 186092 Coleoptera KJ917555 KJ130992 KP212915 Sanjuan et al. 2015
Paraisaria gracilis EFCC 3101 Lepidoptera EF468955 EF468810 EF468750 EF468858 EF468913 Sung et al. 2007
Paraisaria gracilis EFCC 8572 Lepidoptera JN049851 EF468956 EF468811 EF468751 EF468859 EF468912 Ban et al. 2015
Paraisaria heteropoda OSC 106404 Hemiptera AY489690 AY489722 AY489617 AY489651 Quandt et al. 2014
Paraisaria heteropoda EFCC 10125 Hemiptera JN049852 EF468957 EF468812 EF468752 EF468860 EF468914 Quandt et al. 2014
Paraisaria heteropoda NBRC 100643 Hemiptera JN941719 JN941422 AB968595 JN992453 AB968556 Ban et al. 2015
Paraisaria heteropoda BCC 18235 Hemiptera JN941720 JN941421 AB968594 JN992454 AB968555 Ban et al. 2015
(NBRC 100642)
Paraisaria heteropoda BCC 18246 Hemiptera AB968411 AB113352 MK214083 MK214087 Ban et al. 2015
(NBRC 33060)
Paraisaria insignis OSC.164134 Coleoptera OQ709231 OQ800911 OQ708924 OR199807 OR199822 This Study
Paraisaria insignis OSC.164135 Coleoptera OQ709232 OQ800912 OQ708925 OR199808 OR199823 This Study
Paraisaria insignis OSC.164137 Coleoptera OQ709233 OQ800913 OQ708926 OR199809 OR199824 This Study
Paraisaria insignis OSC-M-052004 Coleoptera OQ709234 OQ800914 OQ708927 OR199810 This Study
Paraisaria insignis OSC-M-052008 Coleoptera OQ709236 OQ800917 OQ708930 OR199813 OR199827 This Study
Paraisaria insignis OSC-M-052013 Coleoptera OQ709244 OQ800924 OQ708938 OR199820 OR199834 This Study
Paraisaria orthopterorum BBC 88305 Orthoptera MH754742 MK332583 MK214080 MK214084 Mongkolsamrit et al. 2019
Paraisaria orthopterorum TBRC 9710 Orthoptera MH754743 MK332582 MK214081 MK214085 Mongkolsamrit et al. 2019
Paraisaria phuwiangensis TBRC 9709 Coleoptera MK192015 MK192057 MK214082 MK214086 Mongkolsamrit et al. 2019
Paraisaria phuwiangensis BBH 43492 Coleoptera MH188541 MH201169 MH211355 MH211352 Mongkolsamrit et al. 2019
Paraisaria pseudoheteropoda OSC-M-052005 Hemiptera OQ800915 OQ708928 OR199811 OR199825 OR199836 This Study
Paraisaria pseudoheteropoda OSC-M-052007 Hemiptera OQ709235 OQ800916 OQ708929 OR199812 OR199826 OR199837 This Study
Paraisaria pseudoheteropoda OSC-M-052022 Hemiptera OQ709245 OQ800925 OQ708939 OR199821 OR199835 OR199841 This Study
Paraisaria pseudoheteropoda OSC-M-052020 Hemiptera OQ709243 OQ800923 OQ708937 OR199819 OR199833 This Study
Paraisaria pseudoheteropoda OSC-M-052009 Hemiptera OQ709241 OQ800922 OQ708935 OR199818 OR199832 OR199840 This Study
Paraisaria rosea HKAS_102546 Coleoptera MN947222 MN943846 MN943842 MN929088 MN929081 MN929084 Wei et al. 2021
Paraisaria sp. OSC-M-052011 Insecta OQ709238 OQ800919 OQ708932 OR199815 OR199829 OR199839 This Study
Paraisaria sp. OSC-M-052026 Insecta OQ709242 OQ708936 This Study
Paraisaria tettigonia GZUH CS14062709 Orthoptera KT345954 KT345955 KT375440 KT375441 Wen et al. 2016
Paraisaria yodhathaii BBH 43163 Coleoptera MH188539 MK332584 MH211353 MH211349 Mongkolsamrit et al. 2019
Paraisaria yodhathaii TBRC 8502 Coleoptera MH188540 MH201168 MH211354 MH211350 Mongkolsamrit et al. 2019
Perennicordyceps cuboideus CEM 1514 Coleoptera KF049609 KF049628 KF049683 Kepler et al. 2013
Perennicordyceps prolifica TNS-F-18547 Hemiptera KF049660 KF049613 KF049632 KF049687 KF049649 KF049670 Kepler et al. 2013
Pleurocordyceps nipponicus BCC_2325 Neuroptera KF049665 KF049622 KF049640 KF049696 KF049655 KF049677 Kepler et al. 2013
Pleurocordyceps sinensis ARSEF_1424 Coleoptera KF049661 KF049615 AY259544 DQ118754 DQ127245 KF049671 Kepler et al. 2013
Pleurocordyceps yunnanensis NBRC 101760 Hemiptera MN586827 MN586818 MN586836 MN598051 MN598042 MN598060 Wang et al. 2021
Polycephalomyces formosus CGMCC_5.2204 Coleoptera MN586831 MN586821 MN586839 MN598054 MN598045 MN598061 Wang et al. 2021
Polycephalomyces formosus CGMCC_5.2208 Coleoptera MN586835 MN586825 MN586843 MN598058 MN598049 MN598065 Wang et al. 2021
Purpureocillium atypicola CEM 1185 Araneae KJ878907 KJ878872 KJ878955 Quandt et al. 2014
Purpureocillium atypicola OSC 151901 Araneae KJ878914 KJ878880 KJ878961 KJ878994 Quandt et al. 2014
Purpureocillium takamizusanensis NHJ_3497 Hemiptera EU369096 EU369033 EU369014 EU369053 EU369074 Johnson et al. 2009
Tolypocladium capitatum OSC 71233 Fungi (Eurotiales) AY489689 AY489721 AY489615 AY489649 DQ522421 Spatafora et al. 2007
Tolypocladium inflatum OSC 71235 Coleoptera JN049844 EF469124 EF469077 EF469061 EF469090 EF469108 Kepler et al. 2012
Tolypocladium ophioglossoides OSC 106405 Fungi (Eurotiales) AY489691 AY489723 AY489618 AY489652 DQ522429 Castlebury et al. 2004
Tolypocladiumn japonicum OSC 110991 Fungi (Eurotiales) JN049824 DQ522547 DQ518761 DQ522330 DQ522375 DQ522428 Quandt et al. 2014
Torrubiellomyces zombiae NY04434801 Fungi (Hypocreales) ON493543 ON493602 ON513396 ON513398 ON513402 Araújo et al. 2022

Data analysis

Sequences derived from the SSU, LSU, TEF, RBP1, RPB2, and ITS were aligned with MUSCLE 5.1 (Edgar 2004). Ambiguous and phylogenetically uninformative regions were manually removed and the trimmed alignments were concatenated for analysis using Geneious Prime® 2023.0.4. A Maximum Likelihood Tree was made using the GTR+I+A algorithm and 1000 bootstrap replicates.

Chemical extraction and LCMS analysis

Excisions (0.4–6.7 mg) were made from the endosclerotia of nineteen dried Paraisaria collections, individually placed in MeOH (1 ml, HPLC-grade), sonicated for 5 min, and extracted for 1 hr at 35 °C, then 24 h at ambient temperature. The twenty separate extracts were filtered through syringe filters (0.2 µm PTFE) and dried in vacuo before dissolution in MeOH (0.1 mg/ml, LC-MS-grade) for analysis by LC-MS, injecting 3 µl on a Phenomenex Kinetex column (2.6 µm C18 100 Å, 50 × 2.1 mm), with H2O + 0.1% Formic Acid (A) MeCN + 0.1% Formic Acid (B) as mobile phase solvents at 0.4 ml/min. The LC method was as thus: 0.5 mins at 20% B, a linear gradient from 20–90% B over 14 mins, 4 min at 90% B, a linear gradient from 90–100% B over 0.5 mins, 4.5 mins at 100% B, followed by a linear return to 20% B over 3 mins, and re-equilibration at 20% B for 5 mins, before the next injection. High resolution (Agilent 6545 QToF) mass data were acquired for 26 mins from m/z 100–3200, with MS/MS spectra obtained using data-dependent ion selection for up to five precursor ions per duty cycle, excluding precursor ions with m/z less than 210, and fragmenting with collision energies of 20, 40, and 60 eV. LCMS data files were converted to mzML format and deposited on the public repository MassIVE (MSV000092591). Extracted ion chromatograms were produced for m/z 690–875, corresponding to the mass range for the paraisariamide peptide family (Tehan 2022).

Molecular networking

Unprocessed LC-MS files were converted to mzML format and uploaded to the GNPS online molecular networking platform (version 30) (Wang et al. 2016) using the default network settings but with minimum peak intensity set to 3000. The resulting network was downloaded as a graphML file, analyzed, and visualized using (Ctyoscape ver. 3.9.1). The GNPS job is accessible at

Principal component analysis

LC-MS data were processed in MZmine v2.53 (Pluskal et al. 2010). Feature detection was performed with noise level set to 1×104. Chromatograms were built using a minimum group size of 5, group intensity threshold set to 1×102, minimum highest intensity set to 2×104, and m/z tolerance was set to m/z 0.001 or 10 ppm. Chromatogram deconvolution was performed with minimum peak height set to 1×104, peak duration was set to 0.1–10 mins, and the baseline was set to 5×102. Isotope peaks were grouped with mass tolerance m/z 0.001 or 15 ppm, RT tolerance was set to 1, with the most intense ion taken as the representative, and max charge was set to 2. Peaks were aligned with mass tolerance m/z 0.001 or 12 ppm, RT tolerance set to 0.8 mins, with m/z weighted 75% and RT weighted 25%. Feature list rows were filtered for features falling within the range m/z 690–875, and RT 5–14 mins, with a minimum of 2 peaks per row, and a minimum of 2 peaks in an isotopic pattern. Gap filling was performed with an intensity tolerance of 10%, mass tolerance m/z 0.001 or 15 ppm, and RT tolerance 0.6 mins. The resulting feature list was subjected to Principal Component Analysis (PCA).


Molecular phylogeny

We generated 82 new sequences (16 SSU, 16 LSU, 15 TEF, 14 RPB1, 6 RPB2, and 16 ITS). The combined dataset of 79 taxa afforded a concatenated multi-locus alignment comprising 5,317 bp (1,030 SSU, 955 LSU, 977 TEF, 702 RPB1, 1,037 RPB2, 616 ITS) which was deposited on TreeBASE (accession URL: In the resulting phylogenetic tree (Fig. 1), ten genera in the family Ophiocordycipitaceae are represented. Cordyceps kyushuensis and C. militaris (Cordycipitaceae) were designated as outgroup taxa. All genera, with the exception of Ophiocordyceps, are supported as monophyletic clades. A clade comprising several species morphologically similar to the well-known cicada pathogen, P. heteropoda, referred to here as the “P. heteropoda complex”, is resolved into five well-defined species as well as additional samples revealing cryptic diversity. Two new species within the P. heteropoda complex, Paraisaria cascadensis and Paraisaria pseudoheteropoda, are supported as monophyletic clades, and are described below. Ophiocordyceps insignis samples produced a monophyletic clade within the P. heteropoda complex supporting its combination into Paraisaria, and is redescribed based on a fresh collection, which is designated here as an epitype. The type collection of Ophiocordyceps monticola also occurred within the genus Paraisaria, grouping closely with P. yodhathaii and P. alba. It was the only North American Paraisaria species analyzed in this study which did not fall within the P. heteropoda complex.

Figure 1. 

Maximum likelihood tree based on the combined dataset of SSU, LSU, TEF, RPB1, RPB2, and ITS sequences displaying the relationship of Paraisaria species within family Ophiocordycipitaceae.

LC-MS analysis

Molecular Network Analysis of nineteen Paraisaria endosclerotium extracts revealed a prominent subnetwork identified as the paraisariamide family of cyclopeptides, with constituent molecular ion masses ([M+H]+) ranging from m/z 694.49–860.56 (Fig. 2A.). All endosclerotium extract samples were observed to possess a subset of paraisariamide congeners with partial overlap between species. Production of paraisariamide cyclopeptides in host/endosclerotium is thus supported as a conserved chemotype for Paraisaria. Paraisariamides can thus potentially be used as a generic diagnostic character. Chromatograms generated from the extracted ion range m/z 690–875, corresponding to the mass range for the peptide family of paraisariamides, were unique to and consistent within each species (Fig. 2B). From the processed mass data, a feature list was produced comprising 59 LC-MS ion features (Suppl. material 1). A PCA plot generated from this feature list afforded three major clusters (Fig. 2C). Samples derived from P. insignis and P. pseudoheteropoda were resolved in distinct clusters. Samples derived from P. cascadensis together with samples from its sister clade, “Paraisaria sp. 1”, grouped apart from other samples. Ophiocordyceps monticola afforded two prominent ion peaks with quasimolecular ions, m/z 708.502 and 722.518 eluting at 8.0 and 8.7 min respectively, and grouped most closely with P. insignis in the PCA plot. Qualitatively, the general shape of ion chromatograms was highly conserved within each species and distinct between species. The resolution of species by LC-MS analysis overall accorded very well with the phylogenetic analysis.

Figure 2. 

Chemical comparison of paraisariamide content in the endosclerotia of Paraisaria species collected in the USA A molecular network of the paraisariamide molecular family of cyclic peptides detected in methanol extracts of endosclerotia of Paraisaria specimens. Nodes are displayed as pie charts conveying the relative abundance of paraisariamide mass ion features in each Paraisaria species (Orange = P. cascadensis, Purple = P. pseudoheteropoda, Green = P. insignis, Yellow = “Paraisaria sp. 1”, Red = P. monticola) B extracted ion chromatograms of m/z 690–875 for methanol extracts of endosclerotia of Paraisaria specimens C principal component analysis of mass features m/z 690–875 from methanol extracts of endosclerotia of Paraisaria specimens, color-coded by phylogenetic clade.


Paraisaria cascadensis Tehan, Dooley & Spatafora, sp. nov.

MycoBank No: 849757
Fig. 3

Type material

Holotype. U.S.A., Washington. Skamania County, Gifford Pinchot National Forest, Mt. St. Helens, at approximately 46.1771, -121,9224. 1,042 m alt., 9 June 2021, on adult Cyphoderris monstrosa buried in the ground, in mixed coniferous forest comprising Pinus contorta, Pseudotsuga menziesii, and Abies sp., collected by R. Tehan, C. Dooley (RMT-2021-072, OSC-M-052017, ex-holotype living culture: ARSEF 14609.

Figure 3. 

Paraisaria cascadensis A OSC-M-052017 B fertile head C cross section of fertile head showing arrangement of perithecia D perithecia E ascus F Ascus apex G–I ascospores J part-spores K, L colonies on PDA 61 d (K obverse, L reverse).


cascadensis occurring in the Cascade Mountain range in the Pacific Northwest, USA.


Stroma capitate, solitary, rhizoids solitary arising from heads of adult Cyphoderris monstrosa buried in soil. Ascogenous portion globose or subglobose, 8–9 × 6–9 mm, chestnut brown. Stipe white to light brown, inside hollow, fibrous, white, 15–17 mm long, 3–4 mm wide, papillate with ostioles of perithecia. Perithecia obclavate, immersed, ordinally arranged, 800–970 × 105–150 µm. Asci hyaline, cylindrical, eight-spored, observed up to 350 µm long × 4.5–7 µm wide, possessing abruptly thickened apex. Ascospores hyaline, filiform, multiseptate, breaking into 64 cylindrical part-spores, (6.3–)7.5–9.5(–10.3) × 1.6–2.2(–2.4) µm.

Culture characteristics

Colonies on PDA 61 days at 20 °C, 28 mm, white to yellow, reverse reddish brown to orange. Mycelium septate, smooth-walled hyaline. No conidial state was observed.


Cyphoderris monstrosa (Prophalangopsidae, Orthoptera).


Specimens occur on hypogeous adult hump-winged grigs, Cyphoderris monstrosa, in coniferous forest.

Additional materials examined

U.S.A., Washington: Skamania County, at approximately 46.177, -121.9167, elevation: 974 m, 29 May 2018, on cf. Cyphoderris monstrosa buried in soil, collected by Josh Grefe (OSC-M-052003). U.S.A., Washington: Chelan County, 47.9761, -120.7811, elevation: 865 m, 15 June 2020, on adult Cyphoderris monstrosa, buried in soil, collected by Daniel Winkler, Hans Drabicki (OSC-M-052010). U.S.A., Washington: Skamania County, at approximately 46.1848, -122.1139, elevation: 12332 m, 12 June 2020, on cf. Cyphoderris monstrosa, collected by Ben McCormick (OSC-M-052012). U.S.A., Washington: Skamania County, Gifford Pinchot National Forest, Mt. St. Helens, at approximately 46.1771, -121,9224. 1,042 m alt., 9 June 2021, on adult Cyphoderris monstrosa buried in soil, in mixed coniferous forest comprising Pinus contorta, Pseudotsuga menziesii, and Abies sp., collected by Richard Tehan, Connor Dooley (RMT-2021-071, OSC-M-052016).


This species is uncommon and has thus far only been collected in the Cascade Mountains of Washington State in the vicinity of Mount St. Helens at elevations above 850 m. It might be expected to have a broader range on the basis of the range of its host, Cyphoderris monstrosa, which is known to occur in coniferous forest in several Western U.S. states and Canada (The Orthopterists’ Society 2023).

Paraisaria pseudoheteropoda Tehan & Spatafora, sp. nov.

MycoBank No: 849758
Fig. 4

Type material

Holotype. U.S.A. Arkansas: Searcy County, Grinder's Ferry, 35.985, -92.732, elevation: 252 m, 15 May 2022, on nymphs of cicadidae (Hemiptera) buried in soil, in near Quercus sp., Carya sp., and Juniperus virginiana, collected by Kerri McCabe (OSC-M-052022, ex-type culture: ARSEF 14616).

Figure 4. 

Paraisaria pseudoheteropoda A OSC-M-052022 B fertile head C, D cross section of fertile head showing arrangement of perithecia E perithecia F ascus G ascus apex H, I ascospores J ascospore tip K part-spores L, K colonies on PDA 61 d (L obverse, M reverse).


pseudoheteropoda resembling another cicada-pathogenic species, Paraisaria heteropoda.


Stromata capitate or subclavate, unbranched, growing singly or up to two stromata attached by rhizoids to hypogeous nymphs of Cicadidae (Hemiptera). Ascogenous portion globose or subglobose, 9–11 × 7–8 mm, cream to chestnut brown. Stipe white to light brown, inside fibrous, white, 20–53 mm long, 4–5 mm wide, papillate with ostioles of perithecia. Perithecia obclavate, immersed, ordinally arranged 680–745(–760) × (310–)330–420 µm. Asci hyaline, cylindrical, eight-spored, observed up to 420 µm long × 5.5–6.5 µm wide, possessing abruptly thickened apex. Ascospores hyaline, filiform, multiseptate, breaking into 64 cylindrical part-spores, (5.6–)6.2–7.9(–8.7) × 1.6–2.1(–2.4) µm.

Culture characteristics

Colonies on PDA 61 days at 20 °C, 29 mm, white, reverse yellow to orange. Mycelium septate, smooth-walled hyaline. No conidial state was observed.


Nymphs of Cicadidae (Hemiptera).


Specimens occur on hypogeous nymphs of cicadae at the base of coniferous and deciduous trees, especially oaks.

Additional materials examined

U.S.A. Missouri: Barry County, Cassville, at approximately 36.5586, -93.6833, elevation: 301 m, 26 May 2019, on nymph of cicada buried in soil, collected by Aaron Peters, (OSC-M-052005) U.S.A. Missouri: Barry County, Cassville, at approximately 36.6501, -93.7031, elevation: 382 m, 16 May 2019, on nymph of cicada buried in soil, collected by Aaron Peters (OSC-M-052007) U.S.A. Missouri: Barry County, Cassville, at approximately 36.5586, -93.6833, elevation: 301 m, 4 April 2020, on nymph of cicada buried in soil, collected by Aaron Peters (OSC-M-052009, living culture: ARSEF 14610). U.S.A. Kentucky: Lincoln County, Crab Orchard, at approximately 36.464, -84.51, elevation: 290 m, 19 April 2021, on nymph of cicada buried in soil, collected by Michael Roberts (OSC-M-052015). U.S.A. Tennessee: Putnam County, Cookerville, at approximately 36.163, -85.501, elevation: 337 m, 17 April 2022, on nymph of cicada buried in soil in mixed hardwood forest comprising Quercus sp., Fagus sp., Populus sp. and Arundinaria gigantea, collected by Jamie Newman (OSC-M-052019). U.S.A. Tennessee: Putnam County, Silver Point, at approximately 36.1409, -85.7374, elevation: 180 m, 17 April 2022, on nymph of cicada buried in soil among Acer negundo, Carpinus caroliniana, Carya sp., Quercus rubra, Lindera sp., Amphicarpaea bracteata, Phlox divaricata, Salvia lyrata. collected by Holly Taylor (OSC-M-052020). U.S.A. Arkansas: Searcy County, Grinder's Ferry, at approximately 35.983, -92.719, elevation: 222 m, 14 May 2022, on nymphs of cicadae buried in soil, in near Quercus sp., Carya sp., and Juniperus virginiana, collected by Kerri McCabe (OSC-M-052021). U.S.A. Missouri: Barry County, Roaring River, at approximately 36.5593, -93.683, elevation: 296 m, 24 May 2022, on nymphs of cicadae buried in soil, collected by Aaron Peters, (OSC-M-052023). U.S.A. Virginia: Albemarle County, Charlottesville, at approximately 38.0812, -78.4657, elevation: 133 m, 31 May 2022, on nymph of cf. Neotibicen sp. (Cicadidae, Hemiptera) buried in soil near Acer rubrum, collected by Amelio Little (OSC-M-052024). U.S.A. Missouri: Barry County, Roaring River, at approximately 36.5583, -93.6836, elevation: 305 m, 25 May 2022, on nymphs of cicadae buried in soil, collected by Aaron Peters, (OSC-M-052025). U.S.A. Alabama: St. Clair County, Leeds, at approximately 33.5540, -86.5382, elevation: 198 m, 12 March 2023, on nymphs of cicadae buried in soil, collected by Courtney Mynick, (OSC-M-053266). U.S.A. Alabama: Jefferson County, Birmingham, at approximately 33.4402, -86.8894, elevation: 195 m, 16 March 2023, on nymphs of cicadae buried in soil, collected by Bucky Raeder, (OSC-M-053267).


This species is the only Paraisaria species known to occur on cicadas in North America. In morphology and geographic distribution, it overlaps with P. insignis but that species is distinguished by its strict occurrence on Coleoptera. P. pseudoheteropoda sometimes has a pallid stroma which is not observed in P. insignis.

Paraisaria insignis (Cooke & Ravenel) Tehan & Spatafora, comb. nov.

MycoBank No: 849763
Fig. 5

Cordyceps insignis Cooke & Ravenel, Grevillea 12(no. 61): 38 (1883). Basionym.

Ophiocordyceps insignis (Cooke & Ravenel) G.H. Sung, J.M. Sung, Hywel-Jones & Spatafora, Stud. Mycol. 57: 43 (2007). Synonym.


U.S.A. South Carolina, “seaboard”, 4 January 1881, on larva coleoptera, collected by H. W. Ravenel. (Holotype: Ravenel 3251, K-M 1434269).

Figure 5. 

Paraisaria insignis A OSC-M-052013 Epitype B fertile head C, D cross section of fertile head showing arrangement of perithecia E rhizomorphs F perithecia G ascus H, I asci apices J–L ascospores M part-spores N, O colony on PDA 70 d (N obverse, O reverse).

Epitype designated here: U.S.A. Arkansas: Saline County, Avilla, at approximately 34.713, -92.587, elevation: 169 m, 2 April 2021, on larva of Prionus imbricornis (Cerambycidae, Coleoptera) buried in soil near Quercus sp., collected by Jay Justice (OSC-M-052013, ex-type living culture ARSEF 14611).


Stromata capitate, unbranched, growing singly to gregarious, in groups of up to four stromata on a single host. Stromata 20–52.5 mm long. Ascogenous portion brown, globose to oblong, 8–22 mm long × 7–16 mm wide, papillate with ostioles of perithecia. Stipe golden yellow to reddish orange, sometimes furfuraceous toward upper half, 14–25 × 4–9 mm long, attached to hypogeous host by thick mats of fibrous, tangled, yellow to reddish orange rhizomorphs, extending 25–45 mm. Mycelial growth occurring between, and sometimes over, larval segments, forming a thin membrane. Perithecia embedded, obclavate, brown, (520–)640–800(840) × (160–)185–250(–270) µm. Asci hyaline, cylindrical, up to 380 µ long × (3.8–)4.0–5.9(–7.5) µm, possessing abruptly thickened apex. Ascospores hyaline, filiform, smooth, disarticulating into 64 part-spores. Part-spores, cylindrical, 6.3–9.0(–10.5) × 2.5–3.5 µm. Growing on larvae of Prionus cf. imbricornis. (Cerambycidae, Coleoptera).

Culture characteristics

Colonies on PDA 70 days at 20 °C, 37.5 mm, white, reverse reddish brown to yellow. Mycelium septate, smooth-walled hyaline. No conidial state was observed.


larvae of Prionus cf. imbricornis. (Cerambycidae, Coleoptera)


Specimens occur on hypogeous larvae of coleoptera typically at the base of oak trees.

Additional materials examined

U.S.A. Arkansas: Saline County, Avilla, at approximately 34.713, -92.587, elevation: 169 m, 18 March 2018, on larva of Prionus imbricornis (Cerambycidae, Coleoptera) buried in soil near Quercus sp., collected by Jay Justice (OSC.164134). U.S.A. Arkansas: Saline County, Avilla, at approximately 34.713, -92.587, elevation: 169 m, 2 April 2018, on larva of Prionus imbricornis (Cerambycidae, Coleoptera) buried in soil near Quercus sp., collected by Jay Justice (OSC.164135, living culture: ARSEF 14615). U.S.A. Arkansas: Saline County, Avilla, at approximately 34.713, -92.587, elevation: 169 m, 21 April 2018, on larva of Prionus imbricornis (Cerambycidae, Coleoptera) buried in soil near Quercus sp., collected by Jay Justice (OSC.164136). U.S.A. Arkansas: Pulaski County, North Little Rock, at approximately 34.7989, -92.312, elevation: 99 m, 17 April 2018, on larva of Prionus imbricornis (Cerambycidae, Coleoptera) buried in soil near Quercus sp., and Ulmus sp., collected by Sheila Griffin (OSC.164137). U.S.A. Missouri: Barry County, Cassville, at approximately 36.6116, -93.6938, elevation: 381 m, 16 April 2019, on larva of Prionus imbricornis (Cerambycidae, Coleoptera) buried in soil, collected by Aaron Peters (OSC-M-052004). U.S.A. TEXAS: Harris County, Friendswood, at approximately 29.5501, -95.1972, 19 m, 15 February 2020, on larva of Coleoptera, cf. Prionus imbricornis buried in soil, collected by Brett Jackson (OSC-M-052008). U.S.A. Mississippi: Otibbeha County, at approximately 33.4576, -88.7859, elevation: 109 m, 29 March 2021, on larva of Coleoptera buried in soil near Quercus sp., collected by Carol Siniscalchi (OSC-M-052014) U.S.A. Arkansas: Saline County, Avilla, at approximately 34.713, -92.587, elevation: 169 m, 21 April 2018, on larva of Prionus imbricornis (Cerambycidae, Coleoptera) buried in soil near Quercus sp., collected by Jay Justice (OSC-M-052018, living culture: ARSEF 14617). U.S.A. Georgia: Greene County, Greensboro, at approximately 33.556, -83.262, elevation 152 m, 25 March 2023, on larva of coleoptera, buried in soil, collected by Patti Chaco (OSC-M-053264). U.S.A. Georgia: Bibb County, Musella, at approximately 32.8491, -83.8886, elevation 145 m, 2 April 2023, on larva of coleoptera, buried in soil near Quercus phellos, collected by Rose Payne (OSC-M-053265).


Recent collections of this species were initially determined to not match any described species and were given the provisional name Paraisaria tortuosa, which was used in a doctoral dissertation (Tehan 2022), and in conference presentations. The conspecificity with Ophiocordyceps insignis (=Cordyceps insignis) was considered but it was difficult to reconcile Cooke’s description of the stroma as “livid purple”. However, that species was described from a dried specimen and the true colors of the fresh specimen were evidently not observed by the authority. Petch (1935) cast doubt on the accurate description of the color of C. insignis and though the original host is not able to be precisely identified, Petch’s analysis here is helpful, suggesting based on morphology that the host is one that pupates in wood, which accords with the host of recent collections identified as Prionus imbricornis. Ultimately, chemical comparison of fresh collections to the holotype was definitive in the identification of the fresh collections, and strongly supports the combination into Paraisaria.

Paraisaria monticola (Mains) Tehan & Spatafora, comb. nov.

MycoBank No: 849764
Fig. 6

Cordyceps monticola Mains, Mycologia 32(3): 310 (1940). Basionym.

Ophiocordyceps monticola (Mains) G.H. Sung, J.M. Sung, Hywel-Jones & Spatafora, Stud. Mycol. 57: 45 (2007). Synonym.

Materials examined

Type: U.S.A. Tennessee, Monroe County, Vonore, June 1936, on adult Neocurtilla hexadactyla. collected by G. L. Williams. (BPI 634610).

Figure 6. 

Paraisaria monticola A holotype BPI 634610 B fertile head C ascus D ascus apex E portion of ascospore F part spores.


P. monticola is known to occur on adult Northern mole cricket, Neocurtilla hexadactyla (= Gryllotalpa hexadactyla, Orthoprtera, Gryllotalpidae). Other pathogens of mole crickets, Gryllotalpidae include Beauveria gryllotalpidicola, Beauveria sinensis, Cordyceps neogryllotalpae, Ophiocordyceps gryllotalpae, Ophiocordyceps krachonicola, and Polycephalomycs albiramus, all of which are only known from east Asia. Lloyd (1920) reported C. gryllotalpae from a mole cricket collected in Louisiana, USA, but that specimen was immature, and bore only cylindrical immature stromata with no ascogenous tissue. Owing to the absence of microanatomical character data available for C. gryllotalpae, and the lack of genetic data available for either species, future studies could compare P. monticola to C. gryllotalpae by chemical means, focusing on paraisariamide content of the fungal endosclerotium. P. monticola is only known from the type collection.

Additional Paraisaria specimens examined

Two additional collections were examined which were phylogenetically closest to P. cascadensis but occurring on undetermined insect hosts, outside of the known geographic distribution of Cyphoderris monstrosa, the host of P. cascadensis. Together they form a clade which is sister to P. cascadensis. We do not consider these collections to be conspecific to P. cascadensis, but their formal description was not within the scope of the present study owing to lack of adequate sampling and host data. We anticipate that they represent two distinct new species, the description of which requires further sampling. U.S.A., California: Mendocino County, Ukiah, at approximately 39.1568, -123.2328, elevation: 352 m, 5 April 2019, on undetermined insect host buried in soil, collected by Warren Cardimona (OSC-M-052011) U.S.A., Iowa: Johnson County, Solon, at approximately 41.7572, -91.5457, elevation: 238 m, 30 June 2022, on undetermined insect host buried in soil, collected by Ross Salinas (OSC-M-052026).


In this study, two new Paraisaria species are described and two known species are combined into Paraisaria. The entomopathogenic fungal genus Paraisaria thus currently comprises 18 formally described species which occur on six continents, as deduced from a combination of herbarium records (MycoPortal 2023) and citizen science observations (iNaturalist 2023). The extent of Paraisaria diversity both in North America and worldwide is not comprehensively reflected in this study, which warrants future studies of this group. The results of our phylogenetic and chemical analyses support the presence of additional cryptic diversity yet to be elucidated. For such a geographically widespread genus, there has been a relative paucity of sampling and analyses of Paraisaria specimens globally. Continued study of this group promises to reveal additional new Paraisaria species, each with the potential for new specialized metabolite discovery. In this study, Paraisaria populations in North America prove to be enriched in species falling within the Paraisaria heteropoda complex. Species in this clade are characterized by fruiting bodies with yellow, brown, and reddish hues and prodigious orange to brown rhizomorphs attaching to hypogeous insect hosts. Aboveground portions of the fruiting bodies in some respects resemble the truffle parasite, Tolypocladium capitatum, with which they have been compared (Cooke 1883), and with which they are frequently confused. Numerous host shifts have accompanied speciation in the P. heteropoda complex with species occurring on insect hosts in orders Hemiptera, Diptera, Coleoptera, and Orthoptera. Host identification is critical for field identification of North American Paraisaria species. P. insignis and P. pseudoheteropoda overlap extensively in fruiting body morphology and geographic distribution but are easily distinguished by their respective distinct hosts. P. insignis occurs strictly on coleopteran hosts and P. pseudoheteropoda is the only known Paraisaria species to occur on cicadas in North America. P. cascadensis and P. monticola both occur on orthopteran hosts, but the geographic distribution of P. cascadensis appears to be restricted to montane regions of the Pacific Northwest, which accords with the distribution of its host, Cyphoderris monstrosa. P. monticola is only known from the type specimen collected in Vonore, TN. Re-collection efforts for this species would be valuable and could focus on records of its host Neocurtilla hexadactyla, in the vicinity of the type locality. Notably, N. hexadactyla is widely distributed, and may support a wide distribution of P. monticola.

The life cycles of Paraisaria species, including mode of infection of their insect hosts, their possible occurrence in soil, as endophytes, saprophytic, and nematophagous nutritional modes, are not well characterized. Owing to the observation that Paraisaria species produce fruiting bodies in spring months in North America, we hypothesize that they colonize their insect hosts in the prior season and overwinter as endosclerotia which are observed to possess high concentrations of cyclopeptide specialized metabolites. The molecular structures, biological activities, and chemical ecology of Paraisaria specialized metabolites are the focus of ongoing studies (Tehan 2022).

The targeted LC-MS analysis of specialized metabolites from fungi that are only partially represented in phylogenetic analyses represents a robust application of chemotaxonomy to resolve species. Fungi that produce cyclopeptides may be especially good candidates for chemotaxonomic profiling as many cyclopeptides are particularly resistant to degradation by oxidation, heating, or proteolytic cleavage (Haque and Grayson 2020). Chemotaxonomic profiling of stable metabolites also provides a framework for the analysis of fungal groups lacking genetic data for type specimens, whereby type specimens that afford only chemical data can be linked to samples for which both chemical and genetic data are available, if both types of data resolve species groups. The lack of genetic data for type material is especially challenging when type specimens are very old and possess degraded, highly-fragmented DNA, and for which no suitable neotype has been designated. Micromorphological characters lack robustly distinct differences between Paraisaria species for use in reliable species diagnoses. It was thus critical to compare chemical profiles of recent collections of P. insignis to the holotype to rigorously establish their conspecificity. Conservation of the general paraisariamide chemotype also supports paraisariamides as chemotaxonomic markers for genus Paraisaria, as these compounds were detected in the endosclerotia of all Paraisaria specimens analyzed. These markers are substantially more durable than DNA over long periods of time as is evident from the definitive detection of these compounds in the 142-year-old holotype of P. insignis. Notably, the shape of chromatograms was visually identical between old and new specimens, indicating that even the relative abundance of paraisariamide congeners within a sample is preserved. LC-MS/MS profiling surveys should be conducted across Paraisaria species and related groups of fungi to assess the extent of the paraisariamide molecular family and confirm the utility of these metabolites as chemotaxonomic markers.

Other specialized metabolite families may offer promise as critical chemotaxonomic markers, depending on the relative stability of their biosynthetic genes over time, and whether or not they are reliably expressed. For example, genomic analyses show that the cyclosporin genotype is highly conserved within the insect pathogen, Tolypocladium inflatum (Ophiocordycipitaceae), whereas peptaibiotics have evolved rapidly (Olarte et al. 2019) though neither cyclosporins nor peptaibiotics are detected by LCMS in every Tolypocladium strain exhibiting those genotypes (Blount 2018; Tehan et al. 2022).

Ophiocordyceps blattae, the type species of the large genus Ophiocordyceps, presents another system for potential chemotyping to compare with the various paraphyletic clades of Ophiocordyceps. Grounding of genus Ophiocordyceps in a type species to strictly define a core Ophiocordyceps clade and circumscribe other clades, has remained a longstanding problem owing to the rarity of the type species, and age of its holotype specimen. Increasingly routine chemical profiling by high resolution LC-MS and metabolomics analysis applied to the characterization of fungi in taxonomic studies adds an additional layer of phenotypic assessment that could be indispensable for taxon circumscriptions. Increasing efforts to profile and characterize specialized metabolites in fungi will not only provide useful data for taxonomists but is critical for understanding fungal ecology and may also guide pharmaceutical drug discovery efforts. These pursuits are highly complementary, as demonstrated here and in ongoing research. The isolation, structure elucidation, organic synthesis, biosynthesis, biological characterization, and chemical ecology of the paraisariamides are the focus of ongoing research.


Any opinions, findings, and conclusions or recommendations expressed in this material are those of the author(s) and do not necessarily reflect the views of the National Science Foundation. We thank Dr. Jessie Uehling and Kyle Gervers at the Oregon State University herbarium for assistance with processing loans and accessioning specimens, Lee Davies at the Fungarium of Royal Botanic Gardens, Kew (K), and Shannon Dominick at the U.S. National Fungus Collections (BPI) for assistance processing herbarium loans. We thank Dr. Kathryn Bushley, Michael Wheeler, and Nin Knight at the USDA ARSEF collection for assistance with culture curation. We thank Dr. Chris Marshall at Oregon State University for assistance in the identification of insect hosts.

Additional information

Conflict of interest

The authors have declared that no competing interests exist.

Ethical statement

No ethical statement was reported.


This research was supported in part by the National Institutes of Health via NCCIH 1T32 AT010131 (support of RMT) and NIGMS 1R01GM132649 (KLM), the National Science Foundation (DEB-135944 to JWS, KLM), The Sonoma County Mycological Society, The Oregon Mycological Society, and The Cascade Mycological Society.

Author contributions

Conceptualization: RMT, JWS, KLM. Methodology: RMT, JWS, KLM. Formal analysis: RMT. Investigation: RMT, CBD. Resources: JWS, KLM. Data Curation: RMT, EGB, CBD. Writing - Original draft: RMT. Writing - Review and Editing: RMT, KLM, JWS. Visualization: RMT, EGB. Supervision: JWS, KLM. Project administration: RMT. Funding Acquisition: RMT, KLM, JWS.

Author ORCIDs

Richard M. Tehan

Connor B. Dooley

Edward G. Barge

Kerry L. McPhail

Joseph W. Spatafora

Data availability

All of the data that support the findings of this study are available in the main text or Supplementary Information.


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Supplementary material

Supplementary material 1 

Endosclerotia LCMS feature list

Author: Richard M. Tehan

Data type: csv

Explanation note: This table comprises processed LCMS data for methanol extracts of the endosclerotia of 19 vouchered specimens.

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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