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Research Article
A taxonomic revision of the genus Conidiobolus (Ancylistaceae, Entomophthorales): four clades including three new genera
expand article infoYong Nie‡§, De-Shui Yu§, Cheng-Fang Wang§, Xiao-Yong Liu|, Bo Huang§
‡ Anhui University of Technology, Ma’anshan, China
§ Anhui Agricultural University, Hefei, China
| Institute of Microbiology, Chinese Academy of Sciences, Beijing, China

Corresponding author: Xiao-Yong Liu ( liuxiaoyong@im.ac.cn )

Corresponding author: Bo Huang ( bhuang@ahau.edu.cn )

Academic editor: Marc Stadler
© 2020 Yong Nie, De-Shui Yu, Cheng-Fang Wang, Xiao-Yong Liu, Bo Huang.
This is an open access article distributed under the terms of the Creative Commons Attribution License (CC BY 4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Citation: Nie Y, Yu D-S, Wang C-F, Liu X-Y, Huang B (2020) A taxonomic revision of the genus Conidiobolus (Ancylistaceae, Entomophthorales): four clades including three new genera. MycoKeys 66: 55-81. https://doi.org/10.3897/mycokeys.66.46575
Open Access

Abstract

The genus Conidiobolus is an important group in entomophthoroid fungi and is considered to be polyphyletic in recent molecular phylogenies. To re-evaluate and delimit this genus, multi-locus phylogenetic analyses were performed using the large and small subunits of nuclear ribosomal DNA (nucLSU and nucSSU), the small subunit of the mitochondrial ribosomal DNA (mtSSU) and the translation elongation factor 1-alpha (EF-1α). The results indicated that the Conidiobolus is not monophyletic, being grouped into a paraphyletic grade with four clades. Consequently, the well-known Conidiobolus is revised and three new genera Capillidium, Microconidiobolus and Neoconidiobolus are proposed along with one new record and 22 new combinations. In addition, the genus Basidiobolus is found to be basal to the other entomophthoroid taxa and the genus Batkoa locates in the Entomophthoraceae clade.

Keywords

Zygomycetes, Entomophthorales, Morphology, Phylogeny, New taxa

Introduction

More than 250 species of entomophthoroid fungi were isolated from insects, soil and litter throughout the world (Gryganskyi et al. 2013). For a long time, this group has been considered to be polyphyletic (Nagahama et al. 1995; Jensen et al. 1998; James et al. 2006; Liu and Voigt 2010) and was classified into a subphylum Entomophthoromycotina and a pending taxon Basidiobolus (Hibbett et al. 2007). However, a recent phylogeny using the multi-gene dataset, 18S rDNA, 28S rDNA, mtSSU and RPB2, indicated that this group formed a monophyletic lineage including Basidiobolus and it was consequently reclassified as a new fungal phylum Entomophthoromycota. More recently, a phylogenomic analysis (192 clusters of orthologous proteins) has divided traditional zygomycotan into two phyla Mucoromycota and Zoopagomycota and the entomophthoroid fungi have been re-assigned into the subphylum Entomophthoromycotina under the latter phylum (Spatafora et al. 2016). This taxonomic scheme was supported by the phylogeny of mitochondrial genomes (Nie et al. 2019).

Together with other two genera Ancylistes and Macrobiotophthora, the genus Conidiobolus belongs to Ancylistaceae, Entomophthorales, Entomophthoromycetes, Entomophthoromycotina (Humber 2012). There are six and two accepted species within the Ancylistes and Macrobiotophthora, respectively, while Conidiobolus, one of the largest groups in the entomophthoroid fungi, contains 76 names (http://www.indexfungorum.org/). The genus Conidiobolus is typified by C. utriculosus Bref. 1884 and characterised morphologically by simple sporophores, globose to pyriform multinucleate primary conidia, various types of secondary conidia and resting spores (Brefeld 1884; Humber 1997). Up to the 1940s, for half a century, only three more species were reported, C. minor Bref., C. villosus Martin and C. brefeldianus Couch (Brefeld 1884; Martin 1925; Couch 1939). In the 1950s–1960s, 38 Conidiobolus species and a variety were isolated from the United States and India (Drechsler 1952, 1953a, b, 1954, 1955a, b, c, 1956, 1957a, b, c, 1960, 1961, 1962, 1965; Srinivasan and Thirumalachar 1961, 1962a, b, 1965, 1967, 1968a, b). Based on a numerical taxonomy, King (1976a, b, 1977) recognised 27 definitive species. Since then, along with some new combinations, 10 more species have been added to Conidiobolus (Bałazy et al. 1987; Waters and Callaghan 1989; Bałazy 1993; Huang et al. 2007; Waingankar et al. 2008; Nie et al. 2012, 2016, 2017, 2018). A total of 37 species are currently accepted in this genus (Nie et al. 2018).

Three subgenera – Capillidium, Conidiobolus and Delacroixia – were proposed within the Conidiobolus, based on shape of the secondary conidia and, amongst them, the subgenus Delacroixia was reduced from generic rank (Ben-Ze’ev and Kenneth 1982). This subgeneric criterion provided a valuable contribution for the taxonomy of the genus Conidiobolus (Humber 1989). Since the 1990s, molecular analysis has become an increasingly important tool for fungal taxonomy (Bruns et al. 1991; Taylor et al. 2000). The nucLSU rDNA and EF-1α regions proved to be distinguishable amongst Conidiobolus species (Nie et al. 2012), while nucSSU rDNA indicated the genus Conidiobolus might be a polyphyletic group (Jensen et al. 1998). The subgeneric circumscription was not defined because of instability to form a certain type of secondary conidia for each phylogenetic clade (Callaghan et al. 2000; Gryganskyi et al. 2013; Nie et al. 2018). Besides, the phylogenetic relationships amongst species of Conidiobolus have not been fully resolved due to the absence of types. The genus Batkoa, morphologically similar to Conidiobolus, was phylogenetically closely related to Entomophthoraceae rather than Ancylistaceae (Gryganskyi et al. 2012, 2013).

In the present study, a reclassification of the entomophthoroid fungi, including as many as available Conidiobolus types, was constructed based on four loci (nucSSU, nucLSU, EF-1α and mtSSU) to present the taxonomic delimitation of the genus Conidiobolus and to re-evaluate the phylogenetic relationship between Basidiobolus and Batkoa.

Materials and methods

Isolates and morphology

A total of 26 ex-types of Conidiobolus were purchased from the American Type Culture Collection, Manassas, USA (ATCC) and collected from the China General Microbiological Culture Collection Center, Beijing, China (CGMCC) and the Research Center for Entomogenous Fungi of Anhui Agricultural University, Anhui Province, China (RCEF). Dried cultures were deposited in the Herbarium Mycologicum Academiae Sinicae, Beijing, China (HMAS). Morphology was observed with an Olympus BX51 research microscope and photographed by an Olympus DP25 microscope-camera system. Growth diameter on PDA (potato 200 g, dextrose 20 g, agar 20 g, H2O 1 l), Mycelia, primary conidiophores, primary conidia, microconidia, capilliconidia and resting spores were measured and described with the method of King (1976a).

DNA extraction, PCR amplification and sequencing

Fungal strains were incubated on PDA for 7 d at 21 °C. Total genomic DNA was extracted from the fresh fungal mycelia by using modified CTAB method (Watanabe et al. 2010). Four gene portions from cell nuclei and mitochondria and one protein coding gene were used in this study: the large subunit of nuclear ribosomal RNA genes (nucLSU), the small subunit of nuclear ribosomal RNA genes (nucSSU), the small subunit of mitochondrial ribosomal RNA genes (mtSSU) and the translation elongation factor 1-alpha gene (EF-1α). The nucLSU region was amplified with the primers LR0R and LR5 (Vilgalys and Hester 1990), the nucSSU region with nucSSU-0021-5’ (Gargas and DePriest 1996) and nucSSU-1780-3’ (DePriest 1993) and EF-1α region with the primers EF983 and EF1aZ-1R (http://www.aftol.org/primers.php). These PCR reactions have been described by Liu et al. (2005), Jensen et al. (1998) and Nie et al. (2012). The primers used for the mtSSU region were mtSSU1 and mtSSU2R and the PCR reaction was performed using the following cycling parameters: denaturation at 94 °C for 3 min, followed by 35 cycles of denaturation at 94 °C for 1 min, annealing at 52 °C for 1 min, extension at 72 °C for 1 min and finalised with an extra extension at 72 °C for 7 min (Zoller et al. 1999). PCR products were purified and sequenced by Shanghai Genecore Biotechnologies Company (Shanghai, China) with the same primers as relative PCR. The nucleotide sequence data have been deposited in the GenBank (Table 1).

Table 1.
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The species used in phylogenetic analyses.

Species Strains* GenBank accession numbers
nucSSU nucLSU EF-1α mtSSU
Allomyces arbusculus AFTOL 300 AY552524 DQ273806 DQ275334 –
Basidiobolus haptosporus ARSEF 261 JX242606 JX242586 – JX242626
B. heterosporus CBS 311.66 JX242607 JX242587 – JX242627
B. magnus CBS 205.64 JX242608 JX242588 – JX242628
B. meristosporus CBS 931.73 JX242609 JX242589 – JX242629
B. microsporus CBS 130.62 (T) JX242610 JX242590 – JX242630
B. ranarum NRRL 34594 AY635841 DQ273807 DQ275340 EF392490
Batkoa apiculata ARSEF 3130 DQ177437 EF392404 – EF392513
B. gigantea ARSEF 214 JX242611 JX242591 – JX242631
B. major ARSEF 2936 EF392559 EF392401 – EF392511
B. obscurus** CBS 182.60 JX242614 JX242595 – JX242635
B. pseudapiculata** ARSEF 395 EF392557 EF392378 – EF392508
Coemansia reversa AFTOL 140 AY546685 AY546689 DQ282615 –
Conidiobolus adiaeretus ARSEF 451 (T) – KC461182 – –
C. adiaeretus CGMCC 3.15888 – MN061284 MN061481 MN061287
C. antarcticus ARSEF 6913 (T) – DQ364207 – DQ364227
C. bangalorensis ARSEF 449 (T) – DQ364204 – DQ364225
C. brefeldianus ARSEF 452 (T) AF368506 EF392382 – EF392495
C. chlamydosporus ATCC 12242 (T) – JF816212 JF816234 MK301178
C. coronatus NRRL 28638 AF113418 AY546691 DQ275337 –
C. coronatus RCEF 4518 – JN131537 JN131543 –
C. couchii ATCC 18152 (T) – JN131538 JN131544 MK301179
C. dabieshanensis CGMCC 3.15763 (T) – KY398125 KY402206 MK301180
C. denaeosporus ATCC 12940 (T) – JF816215 JF816228 MK301181
C. firmipilleus ARSEF 6384 JX242612 JX242592 – JX242632
C. gonimodes ATCC 14445 (T) – JF816221 JF816226 MK301182
C. heterosporus RCEF 4430 – JF816225 JF816239 MK301183
C. humicolus ATCC 28849 (T) – JF816220 JF816231 MK301184
C. incongruus NRRL 28636 AF113419 AF113457 – –
C. iuxtagenitus ARSEF 6378 (T) – KC788410 – –
C. iuxtagenitus RCEF 4445 – JX946695 JX946700 MK333391
C. khandalensis ATCC 15162 (T) – KX686994 KY402204 MK301185
C. lachnodes ARSEF 700 – KC788408 – –
C. lamprauges ARSEF 2338 AF296754 DQ364206 – DQ364226
C. lichenicolus ATCC 16200 (T) – JF816216 JF816232 MK301186
C. lobatus ATCC 18153 (T) – JF816218 JF816233 MK301187
C. marcosporus ATCC 16578 (T) – KY398124 KY402209 MK301188
C. megalotocus ATCC 28854 (T) – MF616383 MF616385 MK301189
C. mirabilis CGMCC 3.17763 (T) – MH282852 MH282853 MK333389
C. mycophagus ATCC 16201 (T) – JX946694 JX946698 MK301190
C. mycophilus ATCC 16199 (T) – KX686995 KY402205 MK301191
C. nodosus ATCC 16577 (T) – JF816217 JF816235 MK333388
C. osmodes ARSEF 79 AF368510 EF392371 – DQ364219
C. osmodes RCEF4447 – JN131539 JN131545 MK333392
C. pachyzygosporus CGMCC 3.17764 (T) – KP218521 KP218524 MK333390
C. paulus ARSEF 450 (T) – vv – –
C. polyspermus ATCC 14444 (T) – MF616382 MF616384 MK301193
C. polytocus ATCC 12244 (T) – JF816213 JF816227 MK301194
C. pumilus ARSEF 453 (T) JX242615 EF392383 – EF392496
C. rhysosporus ATCC 12588 (T) – JN131540 JN131546 MK301195
C. sinensis RCEF 4952 (T) – JF816224 JF816238 MK301196
C. stilbeus RCEF 5584 (T) – KP218522 KP218525 MK301197
C. stromoideus ATCC 15430 (T) – JF816219 JF816229 MK301198
C. terrestris ATCC 16198 (T) – KX752050 KY402208 MK301199
C. thromboides ATCC 12587 (T) – JF816214 JF816230 MK301200
C. thromboides FSU 785 JX242616 JX242597 – JX242637
C. thromboides RCEF 4492 – JF816223 JF816236 MK333393
C. undulatus ATCC 12943 (T) – JX946693 JX946699 MK301201
Dimargaris bacillispora AFTOL 136 AB016020 DQ273791 DQ282609 –
Endogone pisiformis AFTOL 539 DQ322628 DQ273811 DQ282618 –
Entomophaga aulicae ARSEF 172 EF392542 EF392372 – EF392487
E. conglomerata ARSEF 2273 AF368509 – – –
E. maimaga ARSEF 1400 EF392556 EF392395 – EF392505
Eryniopsis caroloniana ARSEF 640 EF392552 EF392387 – EF392500
Entomophthora chromaphidis ARSEF 1860 AF353725 – – –
E. culicis ARSEF 387 AF368516 – – –
E. grandis ARSEF 6701 – DQ481229 – –
E. scatophaga ARSEF 6704 – DQ481226 – –
E. muscae ARSEF 3074 AY635820 DQ273772 DQ275343 –
E. planconiana ARSEF 6252 AF353723 GQ285878 – –
E. schizophorae ARSEF 5348 AF052402 GQ285883 – –
E. syrphi ARSEF 5595 – DQ481230 – –
E. tripidium ARSEF 6518 AF296755 – – –
Erynia conica ARSEF 1439 AF368513 EF392396 – EF392506
E. ovispora ARSEF 400 JX242620 JX242601 – JX242641
E. rhizospora ARSEF 1441 AF368514 EF392397 – EF392507
E. sciarae ARSEF 1870 AF368515 EF392399 – EF392509
Furia americana ARSEF 742 EF392554 EF392389 – –
F. gastropachae ARSEF 5541 EF392562 EF392407 – EF392516
F. ithacensis ARSEF 663 EF392553 EF392388 – EF392501
F. neopyralidarum ARSEF 1145 AF368518 EF392394 – EF392504
F. pieris ARSEF 781 AF368519 EF392390 – EF392502
F. virescens ARSEF 1129 EF392555 EF392393 – EF392503
Gaertneriomyces semiglobiferus AFTOL 34 AF164247 DQ273778 DQ275338 –
Macrobiotophthora vermicola ARSEF 650 AF052400 – – –
Massospora cicadina ARSEF 374 EF392548 EF392377 – EF392492
Mortierella verticillata AFTOL 141 AF157145 DQ273794 – –
Pandora blunckii ARSEF 217 (T) JX242621 JX242602 – –
P. delphacis ARSEF 459 AF368521 EF392384 – EF392497
P. dipterigena ARSEF 397 AF368522 EF392380 – EF392565
P. kondoiensis CBS 642.92 JX242622 JX242603 – JX242642
P. neoaphidis ARSEF 3240 EF392560 EF392405 – EF392514
Piptocephalis corymbifera AFTOL 145 AB016023 AY546690 DQ282619 –
Rhizophagus intraradices AFTOL 845 DQ322630 FJ461839 DQ282611 –
Rozella allomycis AFTOL 297 AY635838 DQ273803 DQ275342 –
Schizangiella serpentis ARSEF 2237 AF368523 EF392428 – EF392488
Strongwellsea castrans – AF052406 – – –
Zancudomyces culisetae AFTOL 29 AF277007 DQ273773 – –
Zoophthora anglica ARSEF 396 – EF392379 – EF392493
Z. lanceolata ARSEF 469 EF392550 EF392385 – EF392498
Z. phalloides ARSEF 2281 EF392558 EF392400 – EF392510
Z. radicans ARSEF 388 JX242624 JX242605 – JX242644

* AFTOL, Assembling the Fungal Tree of Life; ARSEF, ARS Entomopathogenic Fungus Collection (Ithaca, U.S.A.); ATCC, American Type Culture Collection (Manassas, U.S.A); CGMCC, China General Microbiological Culture Collection Center (Beijing, China); FSU, Jena Microbial Resource Collection (Friedrich-Schiller-University of Jena, Germany); NRRL, ARS Culture Collection (Peoria, U.S.A); RCEF, Research Center for Entomogenous Fungi (Hefei, China). T = ex-type. **Batkoa sp. CBS 182.60 was received as Conidiobolus obscurus, while B. pseudapiculata ARSEF 395 was received as C. pseudapiculatus.

Phylogenetic analyses

More available nucLSU, nucSSU, mtSSU and EF-1α sequences of 14 Conidiobolus species and 47 other entomophthoroid fungi were obtained from GenBank. Ten species of Glomeromycotina, Mortierellomycotina, Mucoromycotina, Kickxellomycotina, Zoopagomycotina, Blastocladiomycota, Chytridiomycota and Cryptomycota, were chosen as outgroups. Alignments were constructed separately for each locus with MUSCLE 3.8.31 (Edgar 2004) and the concatenated matrices were assembled by SequenceMatrix 1.7.8 (Vaidya et al. 2011). The best model for the phylogenetic analysis was selected with Akaike Information Criterion (AIC) by using Modeltest 3.7 (Posada and Crandall 1998). Phylogenetic relationships were inferred using Maximum Likelihood (ML) and Bayesian Inference (BI). The best-scoring ML tree analysis was performed using raxmlGUI 1.5b1 with GTRGAMMA model and 1000 replicates (Silvestro and Michalak 2012). The BI analysis was performed using MrBayes 3.2.2 (Ronquist and Huelsenbeck 2003). Markov Chain Monte Carlo (MCMC) chains ran until the convergences met and the standard deviation fell below 0.01. The first 25% of trees were discarded as burn-in. The combined dataset was deposited at TreeBase (No. S25064). Phylogenetic trees were checked and modified in FigTree 1.4 (Rambaut 2012).

Results

Phylogenetic analyses

The combined dataset contained 4521 characters of nucLSU (1–1326), nucSSU (1327–3424), EF-1α (3425–4062) and mtSSU (4063–4521) after alignment. With the optimal model GTR+I+G and random starting trees, four Markov chains were run for 7 million generations and every 100th generation was sampled once. ML and BI analyses of the combined dataset resulted in phylogenetic reconstructions with almost similar topologies and the average standard deviation of split frequencies was 0.006721 (BI).

In the ML phylogenetic tree (Figure 1), the Basidiobolaceae lineage (88/0.94) is located at the base of the entomophthoroid fungi and is closely related to the Ancylistaceae group (56/0.91). The Batkoa lineage is grouped within the Entomophthoraceae Clade (60/0.89). All Conidiobolus lineages are clustered into a paraphyletic grade and therefore cannot be considered congeneric. Moreover, the Conidiobolus grade consists of four well supported clades. In detail, there are 7, 10, 16 and 3 species in Clade I (100/1.00), II (77/1.00), III (100/1.00) and IV (99/1.00), respectively.

Figure 1. 

Phylogenetic tree constructed by maximum likelihood analyses of nucLSU, nucSSU, EF-1α and mtSSU sequences for Entomophthoromycotina, with some chytrid and mucoralean fungi as outgroups. Three new genera and one Chinese new record are shown in red. Maximum likelihood bootstrap values (≥ 50%) / Bayesian posterior probabilities (≥ 0.50) of main clades are indicated along branches. Scale bar indicates substitutions per site.

Taxonomy

In order to provide a more natural taxonomic classification, four genera (Capillidium, Conidiobolus, Microconidiobolus and Neoconidiobolus) and their type species (Ca. heterosporum, C. utriculosus, M. paulus and N. thromboides) are described here in this paper. Additionally, a new record Ca. adiaeretum, C. coronatus and C. iuxtagenitus with new isolates from China and C. khandalensis being first reported to produce microconidia are illustrated herein.

Capillidium B. Huang & Y. Nie, gen. nov.

MycoBank No: 831596

Etymology

Referring to unique ellipsoidal secondary conidia (capilliconidia).

Type species

Capillidium heterosporum (Drechsler) B. Huang & Y. Nie.

Description

Mycelia colourless. Primary conidiophores simple, bearing a single primary conidia. Primary conidia forcibly discharged multinucleate, colourless, globose, pyriform to obovoid. Two kinds of replicative conidia, the first one is similar and smaller than primary conidia, the second one (capilliconidia) arises from elongate and slender conidiophores. Zygospores present or absent, formed in axial alignment with conjugating segments, globose to subglobose, often smooth, sometimes rough, colourless or yellowish.

Notes

Conidiobolus subgen. Capillidium Ben-Ze’ev & Kenneth was firstly established to include species with capilliconidia (Ben-Ze’ev and Kenneth 1982). In this phylogenetic analysis, all members of the subgenus Capillidium grouped with good support (100/1.00) and, therefore, it was raised from subgenus to genus status based on the monophyly, as well as the stability to form ellipsoidal secondary conidia (capilliconidia). In addition to capilliconidia, C. adiaeretus also produces microconidia.

Capillidium heterosporum (Drechsler) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831601
Figure 2

Conidiobolus heterosporus Drechsler, Am. J. Bot. 40: 107 (1953). Basionym.

=Conidiobolus rugosus Drechsler, Am. J. Bot. 42: 437 (1955).

Specimens examined

China, Anhui Province, Plant detritus, 8 Nov 2008, C.F. Wang, RCEF 4430.

Description

Colonies on PDA at 25 °C after 3 d, white, reaching ca. 21 mm in diameter. Mycelia colourless, 5–9 μm wide. Primary conidiophores, colourless, unbranched and producing a single globose conidium with widening upwards, extending to a length of 30–245 μm into the air, 8–17 μm wide. Primary conidia forcibly discharged, colourless, globose to subglobose, measuring 12–37 μm in greatest length and 11–31 μm in total width, including a basal papilla 1.5–5 μm high and 5–12 μm wide. After discharging on to 2% water-agar, similar and smaller secondary conidia arise from primary conidia, 1–6 ellipsoidal secondary conidia (capilliconidia, 10–20 × 12–38 μm) arise from slender conidiophores (50–250 × 2.5–4 μm). Resting spores not observed.

Notes

The ex-type living culture is ATCC 12941 (United States, Maryland, 18 Mar 1952, Drechsler).

Figure 2. 

Capillidium heterosporum a colony on PDA after 3 d at 25 °C b primary conidiophores bearing primary conidia c primary conidia d, e, f ellipsoidal secondary conidia arising from slender conidiophores g, h production of secondary conidia. Scale bars: 10 mm (a); 20 μm (b, c, d, g, h); 100 μm (e, f).

Capillidium adiaeretum (Drechsler) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831602
Figure 3

Conidiobolus adiaeretus Drechsler, J. Wash. Acad. Sci. 43: 42 (1953). Basionym.

Specimens examined

China, Jiangsu Province, Nanjing City, Laoshan Forest Park, 32°5'58"N, 118°35'53"E, Plant detritus, 1 Dec 2018, Y. Nie and Y. Gao, HMAS 248358, culture CGMCC 3.15888 (=RCEF 6550).

Description

Colonies on PDA at 25 °C after 3 d, white, reaching ca. 7–10 mm in diameter. Mycelia colourless, 3–4.5 μm wide. Primary conidiophores, colourless, unbranched and producing a single globose conidium with widening upwards; they offer a pronounced dimensional contrast with the mycelial filaments, extending to a length of 50–210 μm into the air, 3–25 μm wide. Primary conidia forcibly discharged, colourless, globose, measuring 15–45 μm in greatest length and 13–42 μm in total width, including a basal papilla 2–6 μm high and 5–17 μm wide. After discharging on to 2% water-agar, similar and smaller secondary conidia arise from primary conidia, two generations of multiple spherical units forming on the parent globose conidia Microconidia only formed from the second set, 5–12 × 9–10 μm. Capilliconidia formed readily from discharged microconidia, 16–24 × 5–6 μm. Chlamydospores formed within the substratum, colourless, globose to ellipsoidal, 13–40 × 15–45 μm.

Notes

The species was firstly reported from America (Drechsler 1953a). The ex-type living culture is ATCC 12589 isolated by Drechsler (1953a). It is mainly characterised and differs from other Capillidium species by its ability to form both microconidia and capilliconidia (Callaghan et al. 2000). The Chinese specimen CGMCC 3.15888 clusters completely (100/1.00) with an isotype ARSEF 451 (98% sequence similarity in nucLSU) and fits well with its morphological descriptions. It is reported in China for the first time.

Figure 3. 

Capillidium adiaeretum a colony on PDA after 3 d at 25 °C b mycelia c, d primary conidiophores bearing primary conidia e, f primary conidia g Production of secondary conidia h first stage of forming microconidia i second stage of forming microconidia j, k ellipsoidal secondary conidia arising from slender conidiophores l chlamydospores. Scale bars: 10 mm (a); 100 μm (b); 20 μm (c–l).

Conidiobolus Bref., Mykol. Untersuch. 6(2): 35 (1884), emend.

MycoBank No: 20144

= Delacroixia Sacc. & P. Syd., Syll. fung. (Abellini) 14(1): 457 (1899).

Conidiobolus subgen. Delacroixia (Sacc. & P. Syd.) Tyrrell & Macleod, J. Invert. Pathol. 20: 12 (1972).

Type species

Conidiobolus utriculosus Bref.

Description

Mycelia colourless. Primary conidiophores simple or branched dichotomously, positively phototropic, bearing a single or 2–4 primary conidia. Primary conidia forcibly discharged, multinucleate, colourless, pyriform, obovoid, globose to subglobose. Secondary conidia usually with shape of primary conidia but smaller, formed singly on short secondary conidiophores. Microspores arising from primary or secondary conidia. Villose appendaged globose conidia and formed villose conidia. Chlamydospores formed intercalarily within assimilative hyphae. Zygospores formed in axial alignment with one or two (homothallic or heterothallic) conjugating segments.

Notes

C. utriculosus, the type species of the genus Conidiobolus, has not been re-collected since Brefeld isolated it in 1884 and most taxonomists working on entomophthoroid fungi now universally recognised it as C. coronatus (Gryganskyi et al. 2013). However, the smaller pear-shaped conidia of C. utriculosus are different from the larger globose conidia of C. coronatus and villose spores in C. coronatus are not observed in C. utriculosus (Brefeld 1884; King 1977). Consequently, C. coronatus is not synonymised with C. utriculosus in this study. Instead, this study agrees with Srinivasan and Thirumalachar (1967) and King (1977) to place C. minor in synonymy with C. utriculosus because the small conidia of C. minor were probably replicative conidia of C. utriculosus. Nevertheless, neither C. utriculosus nor C. minor has available living cultures. Therefore, we have not yet designated an epitype and thus no DNA sequences for explaining this type. Fortunately, we are able to recognise clade III (Fig. 1) as Conidiobolus on the basis of its synapomorph, namely microspores.

Conidiobolus utriculosus Bref., Mykol. Untersuch. 6(2): 35 (1884)

MycoBank No: 144259 (MBT391377)

= Conidiobolus minor Bref., Mykol. Untersuch. 6(2): 35, 68 (1884).

Specimens examined

No ex-type.

Description

Refer to Brefeld (1884) and King (1977).

Notes

Due to the lack of ex-type, plates 3, 4, and 5 in Brefeld, Mykol. Untersuch. 6(2): 35 (1884) are designated here as the lectotype for Conidiobolus utriculosus.

Conidiobolus coronatus (Costantin) A. Batko, Entomophaga, Mémoires hors série 2: 129 (1964)

MycoBank No: 283037
Figure 4

Boudierella coronata Costantin, Bull. Soc. mycol. Fr. 13: 40 (1897). Basionym.

Delacroixia coronata (Costantin) Sacc. & P. Syd., Syll. fung. (Abellini) 14(1): 457 (1899).

Entomophthora coronata (Costantin) Kevorkian, J. Agric. Univ. Puerto Rico 21(2): 191 (1937).

= Conidiobolus villosus G.W. Martin, Bot. Gaz. 80(3): 317 (1925).

Specimens examined

China, Shandong Province, Plant detritus, 20 Mar 2009, C.F. Wang, RCEF 4518.

Description

Colonies grown on PDA for 3 d at 21 °C, reaching ca. 65 mm in diameter. Mycelia colourless, 8–20 μm wide. Primary conidiophores, positively phototropic, colourless, unbranched and producing a single globose conidium, extending to a length of 53–287 μm into the air, 7.5–20.5 μm wide. Primary conidia forcibly discharged, colourless, globose, measuring 36–52 μm in greatest width and 42–65 μm in total length, including a basal papilla 12–18 μm high and 6.5–14 μm wide. After discharging on to 2% water-agar, similar and smaller secondary conidia arise from primary conidia. Microconidia produced readily from primary conidia, globose or almond-shaped, 13–19 × 11–15 μm. Villose spores formed after 4–5 d, globose, 20–42 μm.

Notes

The ex-type living culture is ATCC 28691 (United States, Louisiana, Plant detritus, 3 January 1972). Due to the absence of molecular data of ex-type strain ATCC 28691, the molecular data of the authentic strain NRRL 28638, which has been applied in many other phylogenetic analysis (James et al. 2006; Liu and Voigt 2011; Gryganskyi et al. 2012; Tretter et al. 2014; Spatafora et al. 2016), was used in this study instead. The monotypic genus Delacroixia was typified by D. coronata which was transferred from an ascomycete Boudierella coronata Costantin (Costantin 1897; Saccardo and Sydow 1899). After that, it was reclassified as a subgenus of Conidiobolus, namely Conidiobolus sub. Delacroixia (Sacc. & P. Syd.) Tyrrell & MacLeod to define all those Conidiobolus species capable of forming microspores and, consequently, D. coronata was recombined as C. coronatus (Tyrrell and MacLeod 1972; Ben-Ze’ev and Kenneth 1982).

Figure 4. 

Conidiobolus coronatus a colony on PDA after 3 d at 21 °C b primary conidia c production of secondary conidia d, e primary conidiophores bearing primary conidia f, g microconidia h villose spores. Scale bars: 10 mm (a); 20 μm (b–h).

Conidiobolus iuxtagenitus S.D. Waters & Callaghan, Mycol. Res. 93(2): 223 (1989)

MycoBank No: 135617
Figure 5a–g

Specimens examined

China, Anhui Province, Plant detritus, 8 Nov 2008, C.F. Wang, RCEF 4445.

Description

Colonies on PDA at 21 °C after 3 d white, flat, slow-growing, reaching ca. 13 mm in diameter. Mycelia colourless, 5.5–11 μm wide. Primary conidiophores, positively phototropic, arising from hyphal segments, colourless, 28–75 × 7.5–10 μm, unbranched and producing a single globose conidium. Primary conidia forcibly discharged, globose, 27–37 × 21–28 μm, with a basal papilla 6–10 μm wide. Secondary conidia arising from primary conidia, similar to, but smaller than the primary ones, forcibly discharged. Tertiary conidium fusiform arising from primary conidia, 30–45 × 16–22 μm. Zygospores in a position separated by a short beak near a lateral conjugation, globose to subglobose, smooth, 21–25 × 17–24 μm, with a 1–2 μm thick wall.

Notes

The ex-type living culture is ARSEF 6378 (United Kingdom, Staffordshire, Plant detritus, 31 October 1983, M. F. Smith).

Figure 5. 

a–g Conidiobolus iuxtagenitus h Conidiobolus khandalensis a colony on PDA after 3 d at 21 °C b primary conidiophores bearing primary conidia c primary conidia d tertiary fusiform conidium from a globose spore e zygospore formation with the beak almost emptied of protoplasm f production of secondary conidia g zygospores h microconidia produced from global conidia. Scale bars: 10 mm (a); 20 μm (b–h).

Conidiobolus khandalensis Sriniv. & Thirum., Mycologia 54(6): 692 (1963) [1962]

MycoBank No: 328754
Figure 5h

Specimens examined

India, Khandala, Dec. 1961, Srinivasan and Thirumalachar, ATCC 15162.

Description

Refer to Srinivasan and Thirumalachar (1962b). Microconidia produced from global conidia on the 2% water-agar at 16 °C (Fig. 5h).

Notes

According to the original morphological description (Srinivasan and Thirumalachar 1962b) and the re-examination by King (1977), microconidia have not been reported. However, we observed the microconidia produced from global conidia on 2% water-agar at 16 °C. Moreover, this specimen was located in the Conidiobolus lineage (Figure 1) which was supported by our morphological analyses.

Microconidiobolus B. Huang & Y. Nie, gen. nov.

MycoBank No: 831597

Etymology

Referring to small discharged primary conidia.

Type species

Microconidiobolus paulus (Drechsler) B. Huang & Y. Nie.

Description

Mycelia colourless. Primary conidiophores simple and short, bearing a single primary conidia. Primary conidia forcibly discharged, multinucleate, colourless, globose to obovoid, usually small, mostly less than 20 μm. Only globose replicative conidia produced, similar and smaller than primary conidia. Chlamydospores globose, formed terminally on hyphae or from globose cells by thickening of the wall. Zygospores formed in axial alignment with two conjugating segments, globose to ellipsoidal, smooth and yellowish.

Notes

This genus includes three species producing smaller primary conidia (mostly less than 20 μm) without microspores or capilliconidia compared to other Conidiobolus spp. These three species are C. nodosus, C. paulus and C. terrestris. According to the taxonomic scheme of Conidiobolus by King (1977), C. undulatus is a synonym of C. paulus, which is supported herein by molecular evidence (Figure 1). However, the phylogeny does not support C. nodosus and C. terrestris as synonyms of C. lachnodes, since the former two were located in clade IV and the latter in clade II (Figure 1). Therefore, we accept the taxonomic status at species level for C. nodosus and C. terrestris, based on the morphological and phylogenetic analyses.

Microconidiobolus paulus (Drechsler) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831605

Conidiobolus paulus Drechsler, Bull. Torrey bot. Club. 84: 269 (1957). Basionym.

= Conidiobolus undulatus Drechsler, Bull. Torrey bot. Club. 84: 275 (1957).

= Conidiobolus parvus Drechsler, Bull. Torrey bot. Club. 89: 233 (1962).

Description

Refer to Drechsler (1957a).

Notes

The ex-type living culture is ATCC 12942 (United States, Wisconsin, 18 November 1954, Drechsler).

Neoconidiobolus B. Huang & Y. Nie, gen. nov.

MycoBank No: 831598

Etymology

Referring to the subgenus Conidiobolus raised to generic rank.

Type species

Neoconidiobolus thromboides (Drechsler) B. Huang & Y. Nie.

Description

Mycelia colourless. Primary conidiophores simple, sometimes branched from hyphal knots or differentiated from aerial hyphae, positively phototropic, bearing a single primary conidium. Primary conidia forcibly discharged, multinucleate, colourless, globose, pyriform to obovoid. Replicative conidia similar and smaller than primary conidia. Chlamydospores globose, formed terminally on hyphae or from globose cells by thickening of the wall. Zygospores formed in axial alignment with two conjugating segments, globose to ellipsoidal, smooth, colourless, rarely pale yellowish.

Notes

The genus Neoconidiobolus is strikingly similar to the subgenus Conidiobolus which produces neither microconidia nor capilliconidia. All members in the clade of Neoconidiobolus share the following characteristics: forcibly discharged, colourless, globose, pyriform to obovoid primary conidia. Two kinds of replicative conidia produced. One is discharged, similar and smaller than primary conidia and the other is elongate and forcibly discharged. Two types of resting spores produced: zygospores and chlamydospores.

Neoconidiobolus thromboides (Drechsler) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831606
Figure 6

Conidiobolus thromboides Drechsler, J. Wash. Acad. Sci. 43: 38 (1953). Basionym.

Specimens examined

China, Anhui Province, Plant detritus, 21 Feb 2009, C.F. Wang, RCEF 4492.

Description

Colonies grown on PDA for 3 d at 25 °C, white, reaching ca. 30 mm diameter. Mycelium colourless, filamentous, 5–7.5 µm wide. Primary conidiophores colourless, unbranched and producing a single conidium, 50–122.5 × 6–16.5 µm. Primary conidia forcibly discharged, colourless, globose to subglobose, 20–26.5 µm wide, 26.5–34 µm long, including a basal papilla 6–10 µm wide. Secondary conidia globose, forming from the primary conidia. Zygospores most often formed between segments of separate hyphae. Mature zygospores smooth, globose to subglobose, 25–30 μm in diameter with wall 2–3 μm thick.

Notes

The ex-type living culture is ATCC 12587 (United States, New Hampshire, September 1957, Drechsler).

Figure 6. 

Neoconidiobolus thromboides a colony on PDA after 3 d at 25 °C b, c primary conidiophores bearing primary conidia d production of secondary conidia e zygospores f primary conidia. Scale bars: 10 mm (a); 20 μm (b–d, f); 40 μm (e).

More new combinations

In addition to previously described taxa, more new combinations were proposed herein and their descriptions refer to relevant protologues.

Capillidium bangalorense (Sriniv. & Thirum.) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831607

Conidiobolus bangalorensis Sriniv. & Thirum., Mycologia 59(4): 702 (1967). Basionym.

Capillidium denaeosporum (Drechsler) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831608

Conidiobolus denaeosporus Drechsler, J. Wash. Acad. Sci. 47: 309 (1957). Basionym.

Capillidium lobatum (Sriniv. & Thirum.) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831609

Conidiobolus lobatus Sriniv. & Thirum., J. Elisha Mitchell scient. Soc. 84: 212 (1968). Basionym.

Capillidium pumilum (Drechsler) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831610

Conidiobolus pumilus Drechsler, J. Wash. Acad. Sci. 45: 115 (1955). Basionym.

= Conidiobolus globuliferus Drechsler, Am. J. Bot. 43: 783 (1957) [1956].

= Conidiobolus inordinatus Drechsler, J. Wash. Acad. Sci. 47: 312 (1957).

Capillidium rhysosporum (Drechsler) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831611

Conidiobolus rhysosporus Drechsler, Am. J. Bot. 41: 567 (1954). Basionym.

Microconidiobolus nodosus (Sriniv. & Thirum.) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831624

Conidiobolus nodosus Sriniv. & Thirum., Mycologia 59(4): 705 (1967). Basionym.

Microconidiobolus terrestris (Sriniv. & Thirum.) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831625

Conidiobolus terrestris Sriniv. & Thirum., Mycopathol. Mycol. appl. 36(3–4): 344 (1968). Basionym.

Neoconidiobolus couchii (Sriniv. & Thirum.) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831626

Conidiobolus couchii Sriniv. & Thirum., J. Elisha Mitchell scient. Soc. 84: 211 (1968). Basionym.

Neoconidiobolus lachnodes (Drechsler) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831627

Conidiobolus lachnodes Drechsler, Am. J. Bot. 42: 442 (1955). Basionym.

Neoconidiobolus mirabilis (Y. Nie & B. Huang) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831628

Conidiobolus mirabilis Y. Nie & B. Huang, Mycol. Progr. 17(10): 1204 (2018). Basionym.

Neoconidiobolus osmodes (Drechsler) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831629

Conidiobolus osmodes Drechsler, Am. J. Bot. 41: 571 (1954). Basionym.

= Conidiobolus antarcticus S. Tosi, Caretta & Humber, Mycotaxon 90(2): 344 (2004).

Neoconidiobolus pachyzygosporus (Y. Nie & B. Huang) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831630

Conidiobolus pachyzygosporus Y. Nie & B. Huang, Mycol. Progr. 17(10): 1206 (2018). Basionym.

Neoconidiobolus sinensis (Y. Nie, X.Y. Liu & B. Huang) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831631

Conidiobolus sinensis Y. Nie, X.Y. Liu & B. Huang, Mycotaxon 120: 432 (2012). Basionym.

Neoconidiobolus stilbeus (Y. Nie & B. Huang) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831632

Conidiobolus stilbeus Y. Nie & B. Huang, Mycosphere 7(6): 804 (2016). Basionym.

Neoconidiobolus stromoideus (Sriniv. & Thirum.) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831633

Conidiobolus stromoideus Sriniv. & Thirum., Sydowia 16(1–6): 65 (1963) [1962]. Basionym.

Neoconidiobolus vermicola (J.S. McCulloch) B. Huang & Y. Nie, comb. nov.

MycoBank No: 831634

Entomophthora vermicola J.S. McCulloch, Trans. Br. mycol. Soc. 68(2): 173 (1977). Basionym.

Macrobiotophthora vermicola (J.S. McCulloch) B.E. Tucker, Mycotaxon 13(3): 499 (1981).

Discussion

The phylogenetic position of Basidiobolus in the Kingdom Fungi has been problematic for a long time. Previous phylogenetic analyses of the rDNA (18S, 28S and 5.8S) sequences grouped Basidiobolus outside or basal in the Entomophthorales (Nagahama et al. 1995; Jensen et al. 1998; White et al. 2006). Combined with the study of other protein-coding molecular markers, Basidiobolus was located inside the Entomophthorales (James et al. 2006). Recently, according to the phylogeny of much more available molecular data of entomophthoroid fungi in three families, Basidiobolus was grouped basal to other entomophthoroid taxa (Gryganskyi et al. 2012) which was also supported by the phylogenomic analyses of zygomycete fungi (Spatafora et al. 2016) and by the multi-gene analyses in this study. Although the morphological characteristics of Batkoa were similar to Conidiobolus, the Batkoa lineage appeared to be most closely related to the other taxa in the Entomophthoraceae Clade and should be distinguished from Conidiobolus lineage by its obligate pathogenicity for invertebrates and by staining readily, while most members of Conidiobolus are saprobic and non-staining.

The phylogenetic relationship of the genus Conidiobolus has been unclear for a long time, because of its high heterology (Gryganskyi et al. 2013). This article used more available ex-type strains to revise this genus, based on phylogeny and morphology. According to Figure 1, four main clades were reconstructed and the results showed that Conidiobolus s.l. is not a monophyletic group but paraphyletic with Macrobiotophthora vermicola. The M. vermicola was originally placed in Entomophthora (Mcculloch 1977) and transferred to Macrobiotophthora, based on the morphological characters of primary spores, secondary spores and zygospores (Tucker 1981). The paraphyletic relationship between Macrobiotophthora vermicola and Conidiobolus s.l. was also revealed by Gryganskyi et al. (2012). In this paper, we treated it as a new combination and, therefore, proposed a monophyletic group of the new genus Neoconidiobolus.

In Clade I of the genus Capillidium, seven species grouped in a monophyletic clade with good support (100/1.00) and the synapomorph of producing capilliconidia: Conidiobolus adiaeretus (= Capillidium adiaeretum), Co. bangalorensis (= Ca. bangalorensis), Co. denaeosporus (= Ca. denaeosporum), Co. heterosporus (= Ca. heterosprum), Co. lobatus (= Ca. lobatum), Co. pumilus (= Ca. pumilum) and Co. rhysosporus (= Ca. rhysosporum). As a note, Co. denaeosporus was synonymised with Co. pumilus (King 1976b), but herein its taxonomic status of species level was accepted according to the phylogeny. Co. adiaeretus forms not only capilliconidia but also microspores (Callaghan et al. 2000).

In Clade II of the genus Neoconidiobolus, all 14 strains comprising 10 species produce neither microspores nor capilliconidia. Amongst these, C. antarcticus was identified as a synonym of C. osmodes (Chen and Huang 2018), which was confirmed here as they grouped into a robust clade.

Considering its long history and significant impact, we kept and emended the genus Conidiobolus and the original illustrations of the type species C. utriculosus (Brefeld 1884) were designated as its lectotype. Thus, we were able to recognise clade III under the genus name Conidiobolus on the basis of its synapomorph, namely microspores. In Clade III of the genus Conidiobolus, all species definitely produce microspores, except Conidiobolus dabieshanensis, C. iuxtagenitus, C. khandalensis and C. lichenicolus. Microspores have never been observed in C. dabieshanensis and C. iuxtagenitus (King 1977; Waters and Callaghan 1989; Nie et al. 2017), but cases for C. khandalensis and C. lichenicolus are somewhat different. For C. khandalensis, the protologue did not document any microspores (Srinivasan and Thirumalachar 1962b; King 1977), but they can be observed on 2% water-agar at 16 °C (Fig. 5h). Although the microspore of C. lichenicolus was not mentioned in the original description, the ability to produce microspores has been exhibited in accordance with original illustrations (Srinivasan and Thirumalachar 1968a). The phylogeny also resulted in the following taxonomic treatments. On the one hand, some previously synonymised taxa recover their specific status, for example, C. gonimodes, C. megalotocus and C. mycophagus should be separated from C. incongruus, C. macrosporus and C. mycophilus, respectively. On the other hand, C. chlamydosporus is synonymised with C. firmipilleus.

In Clade IV of the genus Microconidiobolus, Conidiobolus undulatus was identified as a synonym of C. paulus (= M. paulus) by King (1976b), which is supported by our molecular data. Otherwise, C. nodosus (= M. nodosus) and C. terrestris (= M. terristris) were classified as synonyms of C. lachnodes (= Neoconidiobolus lachnodes) in the study of King (1976b). Morphologically, C. lachnodes bears larger primary conidia (9–25 × 10–27 μm) than C. nodosus (13–16 × 17–22 μm) and C. terrestris (8–12 μm in width) (Drechsler 1955b; Srinivasan and Thirumalachar 1967, 1968a). Furthermore, C. lachnodes was located in Clade II and is distantly related to C. nodosus and C. terrestris. Therefore, C. nodosus and C. terrestris are accepted as two distinct species. This clade comprises four ex-type strains, all producing smaller primary conidia (mostly less than 20 μm) and can be morphologically easily distinguished from other Conidiobolus species.

Phylogenetically, Conidiobolus lamprauges does group with Clade III and received strong bootstrap support (100/1.00). Morphologically, this species produces small primary conidia (12.5–20 × 15–22 μm) without microconidia or capilliconidia and is similar to species within Clade IV. Its taxonomic status remains unclear in the present study.

Acknowledgements

We thank Dr. Z.F. Yu (Yunnan University) for improving the manuscript. We also thank Y. Gao (Jiangxi Agricultural University) for collecting some Conidiobolus strains. This project was supported by the National Natural Science Foundation of China (Nos. 31900008, 30770008 and 31670019).

References

  • Bałazy S, Wiśniewski J, Kaczmarek S (1987) Some noteworthy fungi occurring on mites. Bulletin of the Polish Academy of Sciences, Biological Sciences 35: 199–224.
  • Balazy S (1993) Entomophthorales. Flora of Poland (Flora Polska), Fungi (Mycota) 24: 1–356. Polish Academy of Sciences, W. Szafer Institute of Botany, Kraków, Poland.
  • Ben-Ze’ev IS, Kenneth RG (1982) Features-criteria of taxonomic value in the Entomophthorales: I. A revision of the Batkoan classification. Mycotaxon 14: 393–455.
  • Brefeld O (1884) Conidiobolus utriculosus und minor. Untersuchungen aus der Gesammtgebiete der Mykologie 6(2): 35–78.
  • Callaghan AA, Waters SD, Manning RJ (2000) Alternative repetitional conidia in Conidiobolus adiaeretus: development and germination. Mycological Research 104: 1270–1275. https://doi.org/10.1017/S0953756200003063
  • Costantin J (1897) Sur une entomophthoree nouvelle. Bulletin de la Societe Mycologique de France 13: 38–43.
  • Drechsler C (1953a) Three new species of Conidiobolus isolated from leaf mold. Journal of the Washington Academy of Science 43(2): 29–34.
  • Drechsler C (1955a) A small Conidiobolus with globose and with elongated secondary conidia. Journal of Washington Academy of Sciences 45(4): 114–117.
  • Drechsler C (1957a) Two small species of Conidiobolus forming lateral zygospores. Bulletin of the Torrey Botanical Club 84(4): 268–280. https://doi.org/10.2307/2482673
  • Drechsler C (1957b) A new species of Conidiobolus with distended conidiophores. Sydowia Annales Mycologici 9: 189–192.
  • Drechsler C (1957c) Two medium-sized species of Conidiobolus occurring in Colorado. Journal of Washington Academy of Sciences 47: 309–315.
  • Drechsler C (1961) Two species of Conidiobolus often forming zygospores adjacent to antheridium-like distentions. Mycologia 53(3): 278–303. https://doi.org/10.2307/3756275
  • Drechsler C (1962) A small Conidiobolus with resting spores that germinate like zygospores. Bulletin of the Torrey Botanical Club 89(4): 233–240. https://doi.org/10.2307/2483199
  • Drechsler C (1965) A robust Conidiobolus with zygospores containing granular parietal protoplasm. Mycologia 57(6): 913–926. https://doi.org/10.2307/3756891
  • Gargas A, DePriest PT (1996) A nomenclature for fungal PCR primers with examples from intron-containing SSU rDNA. Mycologia 88: 745–748. https://doi.org/10.2307/3760969
  • Gryganskyi AP, Humber RA, Smith ME, Miadlikovska J, Wu S, Voigt K, Walther G, Anishchenko IM, Vilgalys R (2012) Molecular phylogeny of the Entomophthoromycota. Molecular Phylogenetics and Evolution 65: 682–694. https://doi.org/10.1016/j.ympev.2012.07.026
  • Hibbett DS, Binder M, Bischo JF, Blackwell M, Cannon PF, Eriksson OE, Huhndorf S, James T, Kirk PM, Lücking R, Lumbsch HT, Lutzoni F, Matheny PB, McLaughlin DJ, Powell MJ, Redhead S, Schoch CL, Spatafora JW, Stalpers JA, Vilgalys R, Aime MC, Aptroot A, Bauer R, Begerow D, Benny GL, Castlebury LA, Crous PW, Dai YC, Gams W, Geiser DM, Grith GW, Gueidan C, Hawksworth DL, Hestmark G, Hosaka K, Humber RA, Hyde KD, Ironside JE, Kõljalg U, Kurtzman CP, Larsson KH, Lichtwardt R, Longcore J, Miadlikowska J, Miller A, Moncalvo JM, Mozley-Standridge S, Oberwinkler F, Parmasto E, Reeb V, Rogers JD, Roux C, Ryvarden L, Sampaio JP, Schüßler A, Sugiyama J, Thorn RG, Tibell L, Untereiner WA, Walker C, Wang Z, Weir A, Weiss M, White MM, Winka K, Yao YJ, Zhang N (2007) A higher-level phylogenetic classification of the Fungi. Mycological Research 111: 509–547. https://doi.org/10.1016/j.mycres.2007.03.004
  • Huang B, Humber RA, Hodge KT (2007) A new species of Conidiobolus from Great Smoky Mountains National Park. Mycotaxon 100: 227–233.
  • James TY, Kau F, Schoch C, Matheny PB, Hofstetter V, Cox CJ, Celio G, Geuidan C, Fraker E, Miadlikowska J, Lumbsch HT, Rauhut A, Reeb V, Arnold AE, Amtoft A, Stajich JE, Hosaka K, Sung GH, Johnson D, O’Rourke B, Crockett M, Binder M, Curtis JM, Slot JC, Wang Z, Wilson AW, Schüßler A, Longcore JE, O’Donnell K, Mozley-Standridge S, Porter D, Letcher PM, Powell MJ, Taylor JW, White MW, Grith GW, Davies DR, Humber RA, Morton JB, Sugiyama J, Rossman A, Rogers JD, Pfister DH, Hewitt D, Hansen K, Hambleton S, Shoemaker RA, Kohlmeyer J, Volkmann-Kohlmeyer B, Spotts RA, Serdani M, Crous PW, Hughes KW, Matsuura K, Langer E, Langer G, Untereiner WA, Lücking R, Büdel B, Geiser DM, Aptroot A, Diederich P, Schmitt I, Schultz M, Yahr R, Hibbett DS, Lutzoni F, McLaughlin DJ, Spatafora JW, Vilgalys R (2006) Reconstructing the early evolution of Fungi using a six-gene phylogeny. Nature 443: 818–822. https://doi.org/10.1038/nature05110
  • Jensen AB, Gargas A, Eilenberg J, Rosendahl S (1998) Relationships of the insect-pathogenic order Entomophthorales (Zygomycota, Fungi) based on phylogenetic analyses of nucleus small subunit ribosomal DNA sequences (SSU rDNA). Fungal Genetics and Biology 24(3): 325–334. https://doi.org/10.1006/fgbi.1998.1063
  • King DS (1976a) Systematics of Conidiobolus (Entomophthorales) using numerical taxonomy I. Taxonomic considerations. Canadian Journal of Botany 54: 45–65. https://doi.org/10.1139/b76-008
  • King DS (1976b) Systematics of Conidiobolus (Entomophthorales) using numerical taxonomy II. Taxonomic considerations. Canadian Journal of Botany 54: 1285–1296. https://doi.org/10.1139/b76-141
  • King DS (1977) Systematics of Conidiobolus (Entomophthorales) using numerical taxonomy III. Descriptions of recognized species. Canadian Journal of Botany 55: 718–729. https://doi.org/10.1139/b77-086
  • Liu M, Rombach MC, Humber RA, Hodge KT (2005) What’s in a name? Aschersonia insperata: a new pleoanamorphic fungus with characteristics of Aschersonia and Hirsutella. Mycologia 97: 249–256. https://doi.org/10.3852/mycologia.97.1.246
  • Liu XY, Voigt K (2011) Molecular Characters of Zygomycetous Fungi. In: Gherbawy Y, Voigt K (Eds) Molecular Identification of Fungi. Springer-Verlag, Heidelberg, Berlin, 461–488. https://doi.org/10.1007/978-3-642-05042-8_20
  • Miller MA, Pfeiffer W, Schwartz T (2010) Creating the CIPRES Science Gateway for Inference of Large Phylogenetic Trees. Proceedings of the Gateway Computing Environments Workshop (GCE), 14 Nov. 2010, New Orleans, 8 pp. https://doi.org/10.1109/GCE.2010.5676129
  • Nagahama T, Sato H, Shimazu M, Sugiyama J (1995) Phylogenetic divergence of the entomophthoralean fungi: evidence from nuclear 18S ribosomal RNA gene sequences. Mycologia 87: 203–209. https://doi.org/10.2307/3760906
  • Nie Y, Yu CZ, Liu XY, Huang B (2012) A new species of Conidiobolus (Ancylistaceae) from Anhui, China. Mycotaxon 120: 427–435. https://doi.org/10.5248/120.427
  • Nie Y, Tang XX, Liu XY, Huang B (2016) Conidiobolus stilbeus, a new species with mycelial strand and two types of primary conidiophores. Mycosphere 7(6): 801–809. https://doi.org/10.5943/mycosphere/7/6/11
  • Nie Y, Wang L, Cai Y, Tao W, Zhang YJ, Huang B (2019) Mitochondrial genome of the entomophthoroid fungus Conidiobolus heterosporus provides insights into evolution of basal fungi. Applied Microbiology and Biotechnology 103(3): 1379–1391. https://doi.org/10.1007/s00253-018-9549-5
  • Saccardo PA, Sydow P (1899) Delacroixia Sacc. et Syd. Sylloge Fungorum 14: 1–457.
  • Spatafora JW, Chang Y, Benny GL, Lazarus K, Smith ME, Berbee ML, Bonito G, Corradi N, Grigoriev I, Gryganskyi A, James TY, O’Donnell K, Roberson RW, Taylor TN, Uehling J, Vilgalys R, White MM, Stajich JE (2016) A phylum-level phylogenetic classification of zygomycete fungi based on genome-scale data. Mycologia 108(5): 1028–1046. https://doi.org/10.3852/16-042
  • Srinivasan MC, Thirumalachar MJ (1961) Studies on species of Conidiobolus from India-I. Sydowia, Annales Mycologici 15: 237–241.
  • Srinivasan MC, Thirumalachar MJ (1962a) Studies on species of Conidiobolus from India-II. Sydowia, Annales Mycologici 16: 60–66.
  • Srinivasan MC, Thirumalachar MJ (1965) Studies on species of Conidiobolus from India-IV. Sydowia 19: 86–91.
  • Srinivasan MC, Thirumalachar MJ (1967) Evaluation of taxonomic characters in the genus Conidiobolus with key to known species. Mycologia 59: 698–713. https://doi.org/10.2307/3757098
  • Srinivasan MC, Thirumalachar MJ (1968a) Studies on species of Conidiobolus from India-V. Mycopathologica et Mycologia Applicata 36: 341–346. https://doi.org/10.1007/BF02050380
  • Srinivasan MC, Thirumalachar MJ (1968b) Two new species of Conidiobolus from India. Journal of the Mitchell Society 84: 211–212.
  • Taylor JW, Jacobson DJ, Kroken S, Kasuga T, Geiser DM, Hibbett DS, Fisher MC (2000) Phylogenetic species recognition and species concepts in fungi. Fungal Genetics and Biology 31: 21–32. https://doi.org/10.1006/fgbi.2000.1228
  • Tretter ED, Johnson EM, Benny GL, Lichtwardt RW, Wang Y, Kandel P, Novak SJ, Smith JF, White MM (2014) An eight-gene molecular phylogeny of the Kickxellomycotina, including the first phylogenetic placement of Asellariales. Mycologia 106(5): 912–935. https://doi.org/10.3852/13-253
  • Tucker BE (1981) A review of the nonentomogenous Entomophthorales. Mycotaxon 13: 481–505.
  • Vaidya G, Lohman DJ, Meier R (2011) SequenceMatrix: concatenation software for the fast assembly of multi-gene datasets with character set and codon information. Cladistics 27(2): 171–180. https://doi.org/10.1111/j.1096-0031.2010.00329.x
  • Vilgalys R, Hester M (1990) Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species. Journal of Bacteriology 172: 4238–4246. https://doi.org/10.1128/JB.172.8.4238-4246.1990
  • Watanabe M, Lee K, Goto K, Kumagai S, Sugita-Konishi Y, Hara-Kudo Y (2010) Rapid and effective DNA extraction method with bead grinding for a large amount of fungal DNA. Journal of Food Protection 73(6): 1077–1084. https://doi.org/10.4315/0362-028X-73.6.1077
  • White MM, James TY, O’Donnell K, Cafaro MJ, Tanabe Y, Sugiyama J (2006) Phylogeny of the Zygomycota based on nuclear ribosomal sequence data. Mycologia 98: 872–884. https://doi.org/10.3852/mycologia.98.6.872
  • Zollera S, Scheideggera C, Sperisena C (1999) PCR primers for the amplification of mitochondrial small subunit ribosomal DNA of lichen-forming ascomycetes. The Lichenologist 31(5): 511–516. https://doi.org/10.1006/lich.1999.0220
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