Redefining Ceratocystis and allied genera.

The genus Ceratocystis was established in 1890 and accommodates many important fungi. These include serious plant pathogens, significant insect symbionts and agents of timber degradation that result in substantial economic losses. Virtually since its type was described from sweet potatoes, the taxonomy of Ceratocystis has been confused and vigorously debated. In recent years, particulary during the last two decades, it has become very obvious that this genus includes a wide diversity of very different fungi. These have been roughly lumped together due to their similar morphological structures that have clearly evolved through convergent evolution linked to an insect-associated ecology. As has been true for many other groups of fungi, the emergence of DNA-based sequence data and associated phylogenetic inferences, have made it possible to robustly support very distinct boundaries defined by morphological characters and ecological differences. In this study, DNA-sequence data for three carefully selected gene regions (60S, LSU, MCM7) were generated for 79 species residing in the aggregate genus Ceratocystis sensu lato and these data were subjected to rigorous phylogenetic analyses. The results made it possible to distinguish seven major groups for which generic names have been chosen and descriptions either provided or emended. The emended genera included Ceratocystis sensu stricto, Chalaropsis, Endoconidiophora, Thielaviopsis, and Ambrosiella, while two new genera, Davidsoniella and Huntiella, were described. In total, 30 new combinations have been made. This major revision of the generic boundaries in the Ceratocystidaceae will simplify future treatments and work with an important group of fungi including distantly related species illogically aggregated under a single name.

Introduction

Ceratocystis was established in 1890 to accommodate C. fimbriata, a pathogen causing black rot of sweet potatoes in the USA (Halsted 1890). The genus now includes many important fungi including important pathogens of plants and the causal agents of sap stain in timber that are symbiotic associates of insects (Fig. 1). These fungi have ascomata with round usually dark bases that are sometimes ornamented. These bases give rise to long necks terminating in ostiolar hyphae and from which ascospores exude in slimy masses (Fig. 2). All species have ascospores surrounded by sheaths, which can be hat-shaped, ellipsoidal or obovoid and that are either evenly or unevenly distributed around the spores (Fig. 3). The asexual states of most species in Ceratocystis are morphologically “chalara”- or “thielaviopsis”-like forms and characterised by simple, tubular conidiogenous cells. These cells, which are phialides, typically taper towards their apices and produce chains of rectangular conidia or in some cases dark barrel-shaped secondary conidia (Fig. 3). Some species produce simple, single-celled or more complex chlamydospores (Fig. 3) that facilitate a soil-borne life-style.

Fig. 1

Disease symptoms of plants infected with species of Ceratocystis s.l. A. Eucalyptus wilt in Uruguay caused by C. fimbriata s.l. B. Dying clove trees infected with C. polychroma in Sulawesi. C. Wilting shoots of Acacia mearnsii in South Africa infected with C. albifundus. D. Ceratocystis wilt of Acacia sp. caused by C. manginecans. E, F. Wilted shoots and damaged stems of Protea cynaroides in South Africa caused by C. albifundus. G. Resin exudation from the stem of A. mearnsii in South Africa caused by C. albifundus. H. Fungal mats of C. albifundus on Acacia exuvialis. I. Vascular streaking caused by C. manginecans after wounding. J. Fungal mats of C. albifundus on A. exuvialis. K. Staining of the wood of Acacia caused by C. albifundus. L. Streaking and stain of mango trees from infections by C. manginecans in Oman. M. Cross section through a Eucalyptus grandis stump showing streaking caused by C. fimbriata s.l. N. Sweet potato with black rot caused by C. fimbriata s. str. O. Rotted cacao pod infected with C. ethacetica (now T. ethacetica). P. Ascomata of C. polonica (now E. polonica) in the gallery of the bark beetle Ips typographus.

Fig. 2

Morphological features of the ascomata of species of Ceratocystis s.l. A, B. Ascomata of C. albifundus and C. fimbriata respectively, on woody substrates with masses of ascospores emerging from their necks. C–E. Ascomata showing different morphological features such as light-coloured bases of C. albifundus (CMW4059), pear-shaped ascomatal bases characteristic of C. pirilliformis (CMW6579), ornamented bases and divergent necks of C. cerberus (now T. cerberus) (CMW 36668). F, G. Apices of ascomata showing a range of forms of ostiolar hyphae such as long, divergent ostiolar hyphae of C. ethacetica (CMW 36671) (now T. ethacetica) and short, convergent ostiolar hyphae of C. inquinans (now H. inquinans) (CMW 21106). H, I. Hat-shaped ascospores being released from ostiolar hyphae in C. sumatrana (now H. sumatrana) (CMW 21113) and C. pirilliformis (CMW 6670). J. Bases of ascomata in the C. moniliformis s.l. complex (now Huntiella) with distinct plates at the bases of the ascomatal necks, and (K, L) spine-like ornamentations of H. microbasis (CMW 21117) and H. oblonga (CMW 23803) respectively. M. Digitate ornamentations on the ascomatal bases in species residing in C. paradoxa s.l. (now Thielaviopsis) (CMW 36642).

Fig. 3

Sexual and asexual spores in Ceratocystis s.l. A–D. A range of ascospore shapes all with hyaline sheaths and including those that are fusoid [e.g. C. eucalypti (now D. eucalypti), photo from Kile ], hat-shaped (e.g. C. fimbriata, CMW 15049), oblong (e.g. C. paradoxa, now T. paradoxa, CMW 36642) and obovoid (e.g. C. laricicola, now E. laricicola, CMW 20928). E–H. Simple tubular conidiophores commonly tapering to their apicies, and found in most species of Ceratocystis s.l. E. Flasked-shaped phialidic conidiophores of T. paradoxa (CMW 36642) releasing obovoid secondary conidia. F. Phialide releasing cylindrical conidia of C. pirilliformis (CMW 6670). G. Chlamydospore of T. basicola (CMW 7068) and H. C. pirilliformis (CMW 6670). I–L. Darkly pigmented, thick-walled aleurioconidia of (I) T. paradoxa (CMW 36642), (J) T. euricoi (CMW 28537), (K) T. punctulata (CMW 26389) and (L) T. ethacetica (CMW 36671). M, N. Cylindrical and barrel-shaped conidia of C. pirilliformis (CMW 6670). O. Oblong secondary conidia of T. ethacetica (CMW 36671). P. Secondary conidia of T. punctulata (CMW 26389).

Since the time of its first discovery, Ceratocystis has been beset by taxonomic complications and controversy. The first of these emerged with the description of Ophiostoma in 1919 (Sydow & Sydow 1919). It was set up to accommodate several Ceratostomella spp., with O. piliferum as type species and including Ceratostomella moniliformis. Not long thereafter, Melin & Nannfeldt (1934) disposed additional species in the genus, including the type species of Ceratocystis, C. fimbriata. These studies and others (Bakshi 1951, Moreau 1952) resulted in a long-standing confusion between the two genera. This is largely because the genera have morphologically similar ascomata featuring globose bases and generally long necks from which ascospores exude in slimy masses (Upadhyay 1981). According to Malloch & Blackwell (1993) the basic construction of the ascomata may be the result of an adaptation to insect-associated niches and shows the convergent evolution of fruiting structures that facilitate insect-borne transport of spores to new environments (Malloch & Blackwell 1993). Interestingly, but adding to the confusion between them, species of both Ceratocystis and Ophiostoma have evanescent asci that are seldom seen. Ascospores were confused with conidia when the genera were first discovered. The fact that both genera include species with hat-shaped ascospores re-inforced debate over their relationships for many years (Van Wyk ).

The taxonomic confusion between Ceratocystis and Ophiostoma was finally resolved once DNA sequence data became available to provide phylogenetic insights into their relatedness. Hausner and Spatafora & Blackwell (1994) provided the first phylogenetic trees showing that these genera are unrelated. A considerable body of evidence has contributed to the current understanding that Ophiostoma resides in the Ophiostomatales in the Sordariomycetidae and that Ceratocystis is accommodated in the Ceratocystidaceae (Microascales) in the Hypocreomycetidae (Réblová ). Importantly, resolution of the taxonomic confusion regarding these genera has made it possible to study them independently and thus to better understand their similarities, but also their many very different ecologies (Seifert ).

Once Ceratocystis was clearly recognised as unrelated to Ophiostoma, an increasingly clear picture emerged of a genus that included species that were morphologically and ecologically very distinct from one another. These differences have been substantially amplified by the discovery of many new and often cryptic species, revealed through DNA-sequence comparisons (Wingfield ). For example, perhaps the two best-known species names within Ceratocystis, C. fimbriata and C. moniliformis, are now known to represent complexes of many different species (Van Wyk ). Recognition of these complexes has made it possible to interpret their very clear differences.

Wingfield provided the first intensive, phylogenetically based reconsideration of the taxonomy of Ceratocystis. This study included all available sequence data up to 2006 when the study was completed, and it clearly exposed five very different taxonomic groups. These included the species of the C. fimbriata complex, the C. moniliformis complex, and the C. coerulescens complex, as well the Thielaviopsis and Ambrosiella complexes, known only by their asexual states. Importantly, species in these complexes could easily be separated by their morphological and ecological differences. The DNA sequence data used merely reaffirmed the circumscription of the groups. Wingfield provided substantial evidence that species in Ceratocystis s. l. should be assigned to discrete genera. They argued that this would substantially reduce taxonomic confusion among these very different groups of fungi and importantly, also enhance understanding of their different ecologies.

Wingfield were not able to place all species of Ceratocystis s. l. in discrete complexes. Some, such as C. paradoxa, C. adiposa and C. fagacearum fell away from all clearly defined species groups. In retrospect, it appears that this problem stemmed from a lack of sampling and was resolved by the discovery of additional species that could define complexes based on these isolated phylogenetic branches. Such a pattern has become clearly evident from a recent study of a large collection of isolates that would previously have been identified as C. paradoxa (Mbenoun ). These isolates have now been shown to represent a number of very different but related species that are now recognised as comprising the C. paradoxa complex. It is, therefore, very likely that other complexes will emerge in Ceratocystis s. l., as new species are collected and treated in the future.

Ceratocystis s. l., as it is currently defined includes many ecologically important fungi (Fig. 1). For example, most species in the C. fimbriata complex are important and in some cases devastating plant pathogens (Kile 1993, Wingfield ). These include C. albifundus, a virulent pathogen of Acacia mearnsii in Africa (Roux & Wingfield 2013), C. cacaofunesta, a pathogen of cacao in South America (Engelbrecht ), C. platani, an invasive alien pathogen of Platanus trees in Europe (Gibbs 1981, Ocasio-Morales ), and C. manginecans that has devasted mango (Mangifera indica) and Acacia mangium trees in the Middle East and south-east Asia respectively (Van Wyk ).

Species in the C. coerulescens complex include associates of bark beetles (Coleoptera: Scolytinae) as well as important causal agents of sap-stain in timber (Seifert 1993, Wingfield ). The Thielaviopsis complex includes plant pathogens, while the Ambrosiella complex comprise obligate associates of ambrosia beetles (Coleoptera: Scolytinae) (Batra 1967, Kile 1993). Species in the C. moniliformis complex are mostly wound-inhabiting saprobes or mild pathogens, often causing sap stain in timber (Hedgcock 1906, Seifert 1993). The members of the C. paradoxa complex are all pathogens of monocotyledonous plants, including pineapples and palms (Mitchell 1937, Alvarez ).

All available evidence shows that Ceratocystis s. l. represents a suite of morphologically, phylogenetically and ecologically different fungi. There is no reasonable argument for retaining them in a unitary genus, and indeed, doing so would result only in confusion arising from a diminished lack of appreciation of their dramatic differences. Placing them in discrete genera will enhance the perception of opportunities to understand these organisms and, where applicable, to manage or conserve them. It will provide an improved interpretive framework for analysing the ecological differences among the species, such as differences in pathogenicity and insect associations, particularly when complete genome sequences become available for these fungi, as they have recently done for C. fimbriata s. str., C. moniliformis s. str. and C. manginecans (Wilken ).

Revising Ceratocystis s. l. and providing genera to accommodate the well-defined groups in this aggregate genus must be done in conformity with the principles of the new International Code for algae, fungi and plants (Melbourne Code) adopted at the 18th International Botanical Congress (McNeill ). Importantly, this must reflect the One Fungus One Name (1F1N) principles that originally emerged from the Amsterdam Declaration (Hawksworth ) and subsequent discussions (Hawksworth 2011, Norvell 2011, Wingfield ). In this regard, De Beer listed six genus names as possible synonyms of Ceratocystis s. l. One of these names belongs to a sexual genus Endoconidiophora, originally described for E. coerulescens (Münch 1907). The five other names were all considered to denote asexual genera under the dual nomenclature system: they included Thielaviopsis (Went 1893, type species T. ethacetica), Chalaropsis (Peyronel 1916, type species Ch. thielavioides), Hughesiella (Batista & Vital 1956, type species Hu. euricoi), Ambrosiella (Von Arx & Hennebert 1965, type species A. xylebori), and Phialophoropsis (Batra 1967, type species Ph. trypodendri). These names are available for new generic circumscriptions accommodating groups currently residing in Ceratocystis s. l.

The major aim of this study was to revise the generic boundaries for species currently accommodated in Ceratocystis s. l. This task involved obtaining material from as many species as possible and applying 1F1N principles. Generating the full genome sequences for 19 species including representatives of all the phylogenetic groups in Ceratocystis s. l. provided the opportunity to screen multiple gene regions to address genus-level questions. In addition, gene regions from the AFTOL project (Lutzoni ), the ITS barcoding initiative (Schoch ), as well as additional barcoding genes from an ongoing project at CBS (Stielow ) were used to design Microascales-specific primers and to select the most appropriate gene regions to clearly resolve generic boundaries for Ceratocystis s. l.

Materials and methods

Cultures

All cultures used in this study were obtained from the Culture Collection of the Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria, South Africa (CMW) and Centraalbureau voor Schimmelcultures, Utrecht, the Netherlands (CBS). Single spore or single hyphal-tip cultures were prepared and maintained on 2 % Malt Extract Agar (MEA). A list of isolates used in this study is presented in Table 1.

Table 1

Isolates used in the phylogenetic analyses in this study.

Previous name	New name	Country	Host/substrate	Collector; collection year	Herbarium Specimen1	Culture collection number(s)1	Strain status	GenBank accession numbers2
								60S	LSU	MCM7
Ambrosiella beaveri	Ambrosiella beaveri	USA	Vitus rotundifolia	D. Six; 2005	–	CMW 26179; CBS 121753; DLS 1624	ex-paratype	KM495492	KM495315	KM495405
A. ferruginea	A. ferruginea	Germany	Fagus sylvatica	G. Zimmerman; 1971	–	CMW 25522; CBS 460.82	not type	KM495493	KM495316	KM495406
A. hartigii	A. hartigii	Germany	Acer sp.	– ; 1970	–	CMW 25525; CBS 403.82	not type	KM495494	KM495317	–
A. xylebori	A. xylebori	Ivory Coast	Coffea canephora	L. Brader; 1961	–	CMW 25531; CBS 110.61	ex-isotype	KM495495	KM495318	KM495407
Ceratocystis acaciivora	Ceratocystis acaciivora	Indonesia	Acacia mangium	M. Tarigan; 2005	PREM 59884	CMW 22563	ex-holotype	KM495496	KM495319	KM495408
C. adiposa	C. adiposa	Japan	Saccharum officiarum	T. Miyake; 1934	–	CMW 2573; CBS 136.34	not type	KM495497	KM495320	KM495409
C. albifundus	C. albifundus	South Africa	Acacia mearnsii	J. Roux; 1997	–	CMW 4068; CBS 128992	not type	KM495498	KM495321	KM495410
C. atrox	C. atrox	Australia	Eucalyptus grandis	M.J. Wingfield; 2005	PREM 59012	CMW 19385; CBS 120518	ex-holotype	KM495499	KM495322	KM495411
C. bhutanensis	Huntiella bhutanensis	Bhutan	Picea spinulosa	T. Kirisits & D.B. Chhetri; 2001	PREM 57804	CMW 8217; CBS 114289	ex-holotype	KM495500	KM495323	KM495412
C. cacaofunesta	C. cacaofunesta	Ecuador	Theobromae cacao	T.C. Harrington; 2000	BPI 843731	CMW 14803; CBS 115163; C 1695	original collection	KM495501	KM495324	KM495413
C. caryae	C. caryae	USA	Carya ovata	J.A. Johnson; 2001	–	CMW 14808; CBS 115168; C 1827	original collection	KM495502	KM495325	KM495414
C. cerberus	Thielaviopsis cerberus	Cameroon	Elaeis guineensis	M. Mbenoun & J. Roux; 2010	PREM 60770	CMW 36668; CBS 130765	ex-holotype	KM495503	KM495326	KM495415
C. chinaeucensis	H. chinaeucensis	China	Eucalyptus grandis x E. urophylla	M.J. Wingfield & S.F. Chen; 2006	PREM 60735	CMW 24658; CBS 127185	ex-holotype	KM495504	KM495327	KM495416
C. coerulescens	Endoconidiophora coerulescens	Germany	Picea abies	T. Rohde; 1937	–	CMW 26365; CBS 140.37; MUCL 9511; C 313; C 695	not type	KM495506	KM495329	KM495418
C. colombiana	C. colombiana	Colombia	Coffea arabica	M. Marin; 2000	PREM 59434	CMW 5751; CBS 121792	ex-holotype	KM495507	KM495330	KM495419
C. corymbiicola	C. corymbiicola	Australia	Eucalyptus pilularis	G. Kamgan Nkuekam; 2008	PREM 60433	CMW 29349; CBS 127216	ex-paratype	KM495508	KM495331	KM495420
C. curvata	C. curvata	Ecuador	Eucalyptus deglupta	M.J. Wingfield; 2004	PREM 60154	CMW 22432	ex-paratype	KM495509	KM495332	KM495421
C. decipiens	H. decipiens	South Africa	Eucalyptus saligna	G. Kamgan Nkuekam & J. Roux; 2008	PREM 60560	CMW 30855; CBS 129736	ex-holotype	KM495510	KM495333	KM495422
C. diversiconidia	C. diversiconidia	Ecuador	Terminalia ivorensis	M.J. Wingfield; 2004	PREM 60160	CMW 22445; CBS 123013	ex-holotype	KM495511	KM495334	KM495423
C. douglasii	E. douglasii	USA	Pseudotsuga taxifolia	R.W. Davidson; 1951	BPI 595613 = FP 70703	CMW 26367; CBS 556.97	ex-holotype	KM495512	KM495335	KM495424
C. ecuadoriana	C. ecuadoriana	Ecuador	Eucalyptus deglupta	M.J. Wingfield; 2004	PREM 60155	CMW 22092; CBS 124020	ex-holotype	KM495513	KM495336	KM495425
C. ethacetica	T. ethacetica	Malaysia	Ananas comosus	A. Johnson; 1952	PREM 60961	CMW 37775; IMI 50560; MUCL 2170	ex-epitype	KM495514	KM495337	KM495426
C. eucalypti	Davidsoniella eucalypti	Australia	Eucalyptus sieberi	M.J. Dudzinski; 1989	DAR 70205	CMW 3254; C 639	ex-holotype	KM495515	KM495338	KM495427
C. eucalypticola	C. eucalypticola	South Africa	Eucalyptus sp.	M. van Wyk & J. Roux; 2002	PREM 60168	CMW 11536; CBS 124016	ex-holotype	KM495516	KM495339	KM495428
C. fagacearum	C. fagacearum	USA	Quercus rubra	S. Seegmuller; 1991	–	CMW 2656; C463	not type	KM495518	KM495341	KM495430
C. ficicola	C. ficicola	Japan	Ficus carica	Y. Kajitani; 1990	NIAES 20600	CMW 38543; MAFF 625119	ex-holotype	KM495519	KM495342	KM495431
C. fimbriata	C. fimbriata	USA	Ipomoea batatas	C.F. Andrus; 1937	–	CMW 15049; CBS 141.37	not type	KM495520	KM495343	KM495432
C. fimbriatomima	C. fimbriatomima	Venezuela	Eucalyptus hybrid	M.J. Wingfield; 2006	PREM 59439	CMW 24174; CBS 121786	ex-holotype	KM495521	KM495344	KM495433
C. fujiensis	E. fujiensis	Japan	Larix kaempferi	M.J. Wingfield & Y. Yamaoka; 1997	PREM 57513	CMW 1955; CBS 100208; JCM 9810	ex-holotype	KM495522	KM495345	KM495434
C. harringtonii (= C. populicola)	C. harringtonii	Netherlands	Populus hybrid	J. Gremmen; 1978	–	CMW 14789; CBS 119.78; C 995	original collection	KM495523	KM495346	KM495435
C. inquinans	H. inquinans	Indonesia	Acacia mangium	M. Tarigan; 2005	PREM 59866	CMW 21106; CBS 124388	ex-holotype	KM495524	KM495347	KM495436
C. laricicola	E. laricicola	UK	Larix decidua	D. Redfern; 1983	–	CMW 20928; CBS 100207; C 181; Redfern 56-10	ex-paratype	KM495525	KM495348	KM495437
C. larium	C. larium	Indonesia	Styrax benzoin	M.J. Wingfield; 2007	PREM 60193	CMW 25434; CBS 122512	ex-holotype	KM495526	KM495349	–
C. major	C. adiposa	Netherlands	Air	F.H. van Beyma; 1934	–	CMW 3189; CBS 138.34; ATCC 11932; MUCL 9518	ex-holotype	KM495527	KM495350	KM495438
C. mangicola	C. mangicola	Brazil	Mangifera indica	C.J. Rosetto; 2008	PREM 60185	CMW 28908; CBS 127210	ex-paratype	KM495528	KM495351	KM495439
C. manginecans	C. manginecans	Oman	Prosopis cineraria	A. Al Adawi; 2005	–	CMW 17570; CBS 138185	not type	KM495529	KM495352	KM495440
C. mangivora	C. mangivora	Brazil	Mangifera indica	C.J. Rosetto; 2001	PREM 60570	CMW 27305; CBS 128702	ex-holotype	KM495530	KM495353	KM495441
C. microbasis	H. microbasis	Indonesia	Acacia mangium	M. Tarigan; 2005	PREM 59872	CMW 21117	ex-holotype	KM495531	KM495354	KM495442
C. moniliformis	H. moniliformis	South Africa	Eucalyptus grandis	M. van Wyk; 2002	–	CMW 10134; CBS 118127	not type	KM495532	KM495355	KM495443
C. moniliformopsis	H. moniliformopsis	Australia	Eucalyptus obliqua	Z.Q. Yuan; 2001	DAR 74608	CMW 9986; CBS 109441	ex-holotype	KM495533	KM495356	KM495444
C. musarum	T. musarum	New Zealand	Musa sp.	T.W. Canter-Visscher; –	PREM 60962	CMW 1546; C 907	ex-epitype	KM495534	KM495357	KM495445
C. neglecta	C. neglecta	Colombia	Eucalyptus grandis	C. Rodas & J. Roux; 2004	PREM 59616	CMW 17808; CBS 121789	ex-holotype	KM495535	KM495358	KM495446
C. oblonga	H. oblonga	South Africa	Acacia mearnsii	R.N. Heath; 2006	PREM 59792	CMW 23803; CBS 122291	ex-holotype	KM495536	KM495359	KM495447
C. obpyriformis	C. obpyriformis	South Africa	Acacia mearnsii	R.N. Heath; 2006	PREM 59796	CMW 23808; CBS 122511	ex-holotype	KM495537	KM495360	KM495448
C. omanensis	H. omanensis	Oman	Mangifera indica	A. Al Adawi & M. Deadman; 2003	–	CMW 11056; CBS 118113	original collection	KM495538	KM495361	KM495449
C. papillata	C. papillata	Colombia	Citrus x Tangelo hybrid	B. Castro; 2001	PREM 59438	CMW 8856; CBS 121793	ex-holotype	KM495539	KM495362	KM495450
C. paradoxa	T. paradoxa	Cameroon	Theobromae cacao	M. Mbenoun & J. Roux; 2010	PREM 60766	CMW 36689; CBS 130761	ex-epitype	KM495540	KM495363	KM495451
C. pinicola	E. pinicola	UK	Pinus sylvestris	J. Gibbs; 1988	DAOM 225447	CMW 29499; CBS 100199; C 488; DAOM 225447	ex-holotype	KM495541	KM495364	KM495452
C. pirilliformis	C. pirilliformis	Australia	Eucalyptus nitens	M.J. Wingfield; 2000	PREM 57323	CMW 6579; CBS 118128	ex-holotype	KM495542	KM495365	KM495453
C. platani	C. platani	USA	Platanus occidentalis	T.C. Harrington; 1998	–	CMW 14802; CBS 115162; C 1317	original collection	KM495543	KM495366	KM495454
C. polonica	E. polonica	Norway	Picea abies	H. Solheim; 1990	DAOM 225451	CMW 20930; CBS 100205; C791	ex-neotype	KM495544	KM495367	KM495455
C. polychroma	C. polychroma	Indonesia	Syzygium aromaticum	E.C.Y. Liew; 2002	PREM 57818	CMW 11424; CBS 115778	ex-holotype	KM495545	KM495368	KM495456
C. polyconidia	C. polyconidia	South Africa	Acacia mearnsii	R.N. Heath; 2006	PREM 59788	CMW 23809; CBS 122289	ex-holotype	KM495546	KM495369	KM495457
C. radicicola	T. punctulata	USA	Phoenix dactylifera	D.E. Bliss; –	BPI 596268	CMW 1032; CBS 114.47; MUCL 9526	ex-holotype	KM495548	KM495371	KM495459
C. resinifera	E. resinifera	Norway	Picea abies	H. Solheim; 1986	DAOM 225449	CMW 20931; CBS 100202; C 662	ex-holotype	KM495549	KM495372	KM495460
C. rufipennis	E. rufipennis	Canada	Picea engelmannii	H. Solheim; 1992	–	CMW 11661	original collection	KM495550	KM495373	–
C. salinaria	H. salinaria	South Africa	Eucalyptus maculata	G. Kamgan Nkuekam; 2007	PREM 60557	CMW 25911; CBS 129733	ex-holotype	KM495551	KM495374	KM495461
C. savannae	H. savannae	South Africa	Acacia nigrescens	G. Kamgan Nkuekam & J. Roux; 2005	PREM 59423	CMW 17300; CBS 121151	ex-holotype	KM495552	KM495375	KM495462
C. smalleyi	C. smalleyi	USA	Carya cordiformis	E. Smalley; 1993	BPI 843722	CMW 14800; CBS 114724; C 684	ex-holotype	KM495553	KM495376	KM495463
C. sublaevis	H. sublaevis	Ecuador	Terminalia ivorensis	M.J. Wingfield; 2004	PREM 60163	CMW 22449; CBS 122517	ex-paratype	KM495554	KM495377	KM495464
C. sumatrana	H. sumatrana	Indonesia	Acacia mangium	M. Tarigan; 2005	PREM 59868	CMW 21109; CBS 124011	ex-paratype	KM495555	KM495378	KM495465
C. tanganyicensis	C. tanganyicensis	Tanzania	Acacia mearnsii	R.N. Heath & J. Roux; 2004	–	CMW 15999; CBS 122294	ex-paratype	KM495556	KM495379	KM495466
C. thulamelensis	C. thulamelensis	South Africa	Colophospermum mopane	M. Mbenoun & J. Roux; 2010	PREM 60828	CMW 35972; CBS 131284	ex-holotype	KM495557	KM495380	KM495467
C. tribiliformis	H. tribiliformis	Indonesia	Pinus merkusii	M.J. Wingfield; 1996	PREM 57827	CMW 13013; CBS 115866	ex-holotype	KM495558	KM495381	KM495468
C. tsitsikammensis	C. tsitsikammensis	South Africa	Rapanea melanophloeos	G. Kamgan Nkuekam; 2005	PREM 59424	CMW 14276; CBS 121018	ex-holotype	KM495559	KM495382	KM495469
C. tyalla	H. tyalla	Australia	Eucalyptus dunnii	G. Kamgan Nkuekam & A.J. Carnegie; 2008	–	CMW 28932; CBS 128703	ex-holotype	KM495560	KM495383	KM495470
C. variospora	C. variospora	USA	Quercus alba	J.A. Johnson; 2001	BPI 843737	CMW 20935; CBS 114715; C 1843	ex-paratype	KM495561	KM495384	KM495471
C. virescens	D. virescens	USA	Acer saccharum	D. Houston; 1987	–	CMW 17339; CBS 130772; C 261	not type	KM495562	KM495385	KM495472
C. zambeziensis	C. zambeziensis	South Africa	Acacia nigrescens	M. Mbenoun & J. Roux; 2010	PREM 60826	CMW 35963; CBS 131282	ex-paratype	KM495563	KM495386	KM495473
Chalaropsis sp. 1	Chalaropsis sp. 1	Belgium	Populus sp.	R. Veldeman; 1975	–	CMW 22737; CBS 180.75	not type	KM495580	KM495403	KM495490
Chalaropsis sp. 1	Chalaropsis sp. 1	USA	Ulmus sp.	R.W. Davidson; 1939	–	CMW 22738; CBS 130.39; C 1378; MUCL 9540; RWD E-1	not type	KM495581	KM495404	KM495491
Graphium fabiforme	Graphium fabiforme	Madagascar	Dead Adansonia rubrostipa	J. Roux & M.J. Wingfield; 2007	PREM 60310	CMW 30626; CBS 124921	ex-holotype	KM495564	KM495387	KM495474
G. fimbriisporum	G. fimbriisporum	France	Ips typographus gallery, in stump of Picea abies	M. Morelet; 1992	PFN 1494	CMW 5605; CBS 870.95; MPFN 281-8	ex-holotype	KM495565	KM495388	KM495475
G. laricis	G. laricis	Austria	Synnemata occuring in galleries of the bark beetle Ips cembrae	T. Kirisits & P. Baier; 1995	DAOM 229757	CMW 5601; CBS 116194; DAOM 229757; IFFF ICL/MEA/13	ex-holotype	KM495566	KM495389	KM495476
G. pseudormiticum	G. pseudormiticum	South Africa	Pinus sp.	M.J. Wingfield; 1984	PREM 51539	CMW 503	ex-holotype	KM495567	KM495390	KM495477
Huntiella chlamydoformis nom. prov.	H. chlamydoformis nom. prov.	Cameroon	Theobromae cacao	M. Mbenoun & J. Roux; 2009	PREM 60837	CMW 36932; CBS 131674	ex-holotype	KM495505	KM495328	KM495417
H. pycnanthi nom. prov.	H. pycnanthi nom. prov.	Cameroon	Theobromae cacao	M. Mbenoun; 2009	PREM 60835	CMW 36916; CBS 131672	ex-holotype	KM495547	KM495370	KM495458
Knoxdaviesia capensis	Knoxdaviesia capensis	South Africa	Protea longifolia	M.J. Wingfield; 1984	–	CMW 997; CBS 120015	not type	KM495568	KM495391	KM495478
K. cecropiae	K. cecropiae	Costa Rica	Cecropia angustifolia	L. Kirkendall & J. Hulcr; 2005	PRM 858080	CMW 22991; CCF 3565	ex-holotype	KM495569	KM495392	KM495479
K. proteae	K. proteae	South Africa	Protea repens flower infested with insects	L.J. Strauss; 1985	PREM 48924	CMW 738; CBS 486.88	ex-holotype	KM495570	KM495393	KM495480
K. serotectus	K. serotectus	South Africa	Grow on insect (Cossonus sp.) found in Euphorbia ingens	J.A. van der Linde & J. Roux; 2009	PREM 60566	CMW 36767; CBS 129738	ex-holotype	KM495571	KM495394	KM495481
K. ubusi	K. ubusi	South Africa	Insect tunnels in Euphorbia tetragona	J. Roux; 2010	PREM 60568	CMW 36769; CBS 129742	ex-holotype	KM495572	KM495395	KM495482
Thielaviopsis australis	D. australis	Australia	Nothofagus cunninghamii	M. Hall; 2001	–	CMW 2333	not type	KM495573	KM495396	KM495483
T. basicola	T. basicola	Netherlands	Lathyrus odoratus	G.A. van Arkel; –	–	CMW 7068; CBS 413.52	not type	KM495574	KM495397	KM495484
T. ceramica	H. ceramica	Malawi	Eucalyptus grandis	R.N. Heath & J. Roux; 2004	PREM 59808	CMW 15245; CBS 122299; CMW 15251	ex-holotype	KM495575	KM495398	KM495485
T. euricoi	T. euricoi	Brazil	Air	E.A.F. da Matta; 1956	URM 640	CMW 28537; CBS 893.70; MUCL 1887; UAMH 1382	ex-holotype	KM495517	KM495340	KM495429
T. neocaledoniae	D. neocaledoniae	New Caledonia	Coffea robusta	R. Dadant; 1948	–	CMW 3270; CBS 149.83; C 694	ex-holotype	KM495576	KM495399	KM495486
T. ovoidea	Chalaropsis ovoidea	Netherlands	Firewood	W. Gams; 1976	–	CMW 22733; CBS 354.76; C 1375	not type	KM495577	KM495400	KM495487
T. ovoidea	Ch. ovoidea	Germany	Quercus petraea	H. Kleinhempel; 1987	–	CMW 22732; CBS 136.88	not type	KM495578	KM495401	KM495488
T. Thielavioides	Ch. thielavioides	Italy	Lupinus albus	R. Ciferri; 1937	–	CMW 22736; CBS 148.37; MUCL 6235	not type	KM495579	KM495402	KM495489

ATCC: American Type Culture Collection, Virginia, U.S.A.; BPI: US National Fungus Collections, Systematic Botany and Mycology Laboratory, Maryland, U.S.A.; C: Culture collection of T.C. Harrington, Iowa State University, U.S.A.; CBS: Culture collection of the CBS-KNAW Fungal Biodiversity Centre, Utrecht, The Netherlands; CCF: Culture Collection of Fungi, Department of Botany, Faculty of Science, Charles University, Prague, Czech Republic; CMW: Culture collection Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria, Pretoria, South Africa; DAOM: Plant Research Institute, Department of Agriculture (Mycology), Ottawa, Canada; DAR: New South Wales, Plant Pathology Herbarium, Australia; DLS: Culture collection of D. Six, University of Montana, U.S.A.; FP: Rocky Mountain Forest & Range Experimental Station Herbarium, Fort Collins, Colorado, U.S.A.; IFFF: Culture collection of the Institute of Forest Entomology, Forest Pathology and Forest Protection (IFFF), University of Natural Resources and Applied Life Sciences, Vienna (BOKU), Vienna, Austria; IMI: International Mycological Institute, CABI-Bioscience, Egham, Bakeham Lane, United Kingdom; JCM: Japan Collection of Microorganism, RIKEN BioResource Center, Japan; MAFF: Ministry of Agriculture, Forestry and Fisheries, Tsukuba, Ibaraki, Japan; MPFN: Culture collection at the Laboratoire de Pathologie Forestière, INRA, Centre de Recherches de Nancy, 54280 Champenoux, France; MUCL: Université Catholique de Louvain, Louvain-la-Neuve, Belgium; NIAES: National Institute for Agro-Environmental Sciences, 3-1-3 Kannondai, Tsukuba, 305-8604, Japan; PREM: National Collection of Fungi, Pretoria, South Africa; PRM: Corda Herbarium, Prague, Czech Republic; Redfern: Culture Collection of D.B. Redfern, Forestry Commission, Northern Research Station, Roslin, Midlothian, UK; RWD: Culture collection of R.W. Davidson, Department of Forest and Wood Sciences, Colorado State University, Fort Collins, Colorado; UAMH: University of Alberta Microfungus Collection and Herbarium, Edmonton, Alberta, Canada; URM: Father Camille Torrend Herbarium-URM (previously University of Recife Herbarium), Department of Mycology, Universidade Federal de Pernambuco, Recife, Brazil.

60S: partial 60S ribosomal protein RPL10 gene; LSU: partial nuclear ribosomal DNA large subunit (28S); MCM7: partial mini-chromosome maintenance complex component 7 gene.

DNA extraction

Single spore/single hyphal-tip cultures were inoculated in YM broth (2 % malt extract, 0.2 % yeast extract) and incubated at 25 °C with shaking for 2–5 d. Mycelium was harvested and freeze-dried in 2 mL Eppendorf tubes. The freeze-dried mycelium was submerged in liquid nitrogen, followed by pulverising the mycelium with a pipette tip. About 10 mg of mycelial “powder” was used for DNA extraction using PrepMan Ultra Sample Preparation reagent (Applied Biosystems, Foster City, California) as described in Duong .

Selection of gene regions and primers

Ten different gene regions [the nuclear ribosomal DNA large subunit (LSU), the nuclear ribosomal DNA small subunit (SSU), nuclear ribosomal DNA internal transcribed spacer regions (ITS), the 60S ribosomal protein RPL10 (60S), beta-tubulin (BT), translation elongation factor 1-alpha (EF1), translation elongation factor 3-alpha (EF3), mini-chromosome maintenance complex component 7 (MCM7), the RNA polymerase II largest subunit (RPB1), and the RNA polymerase II second largest subunit (RPB2)] were extracted from 19 Ceratocystis draft genome sequences that included species from all the major clades. The genome sequences, of which three have been published (Wilken ), are available at the Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria. Phylogenetic analyses were conducted with all ten gene regions (data not shown). LSU, 60S, and MCM7 were selected as candidate genes for further investigation including all the isolates in the study, based on their level of support at the basal nodes, the ease of amplification and sequencing, and the popularity of their use in studies of other fungal lineages.

The ITS region has been widely used in phylogenetic studies to distinguish between species in Ceratocystis. However, due to the recent discovery of multiple ITS forms in certain species of Ceratocystis (Al Adawi ), and the fact that gene regions were chosen that were slightly more conserved to resolve the genus level questions, the ITS was intentionally not used in the present study.

Primers LR0R and LR5 (Vilgalys & Hester, 1990) were used in PCR amplification and sequencing of LSU. Primers Algr52_412-433_f1 and Algr52_1102_1084_r1 (Stielow ) were used for PCR amplification and sequencing of 60S. Based on the sequences obtained from genomes, new primers Cer-MCM7F (ACICGIGTITCIGAYGTNAAGCC) and Cer-MCM7R (TTRGCAACACCAGGRTCACCCAT) were designed and used in PCR amplification and sequencing of MCM7.

PCR and sequencing

All PCR reactions were done in a total volume of 25 μL. The reaction mixture consisted of 2.5 μL of 10X PCR reaction buffer, 2.5 mM MgCl2, 200 μM of each dNTP, 0.2 μM of each of the forward and reverse primers for LSU (1 μM of each primer in case of degenerate primers for 60S and MCM7), 1 U FastStart Taq DNA Polymerase (Roche) and 2 μL of genomic DNA solution. The PCR thermal conditions included an initial denaturation at 96 °C for 5 min, followed by 35 cycles of 95 °C for 30 sec, 55 °C for 30 s, and 72 °C for 60 s, and ended with a final extension at 72 °C for 8 min. The annealing temperature was set at 55 °C for all gene regions and all isolates at first. In some cases where the PCR failed or non-specific amplification was observed, we experimented with different annealing temperatures (between 52 °C and 60 °C) until successful amplification was obtained. Direct sequencing of PCR products was done using BigDye® Terminator v. 3.1 Cycle Sequencing kit (Applied Biosystems) with a 1/16 reaction and at 55 °C annealing temperature for all primers. Sequencing PCR products were precipitated using the sodium acetate and ethanol precipitation protocol, followed by fragment separation using an ABI PRISM® 3100 Genetic Analyzer (Applied Biosystems).

Phylogenetic analyses

Sequences from different gene regions were aligned using an online version of MAFFT v. 7 (Katoh & Standley 2013). The three gene regions (LSU, 60S and MCM7) were combined and analysed as a single dataset. Each of the gene regions was also analysed separately and results were compared with those of the combined analyses. Maximum parsimony (MP) analyses were performed in MEGA6 (Tamura ) with 1000 bootstrap replications. The subtree-pruning-regrafting (SPR) algorithm was selected, and alignment gaps and missing data included. Maximum likelihood (ML) analyses were done using raxmlGUI (Silvestro & Michalak 2012) with the GTR+G+I substitution model selected. Ten parallel runs with four threads and 1000 bootstrap replications were conducted. Bayesian inference (BI) analyses were performed using MrBayes v. 3.2 (Ronquist ) employing the GTR+G+I substitution model. Ten parallel runs, each with four chains, were conducted. Trees were sampled at every 100th generation for 5 M generations. After sampling, 25 % of trees were discarded as a burn-in phase and posterior probabilities were calculated from all the remaining trees.

Morphology

Morphological descriptions from the protologues of all species treated in this study were carefully considered when genera were redefined. Based on these species descriptions, the most common characters of all species in a genus were selected and incorporated in the emended and new genus descriptions. Over time, different authors often used different terminology describing similar characters. We aligned the generic descriptions of the different genera with each other using similar terminology.

Results

Maximum likelihood, BI and MP trees obtained from analyses of the individual gene regions (Figs 4–6) and the combined datasets (Fig. 7) of the LSU, 60S and MCM7 sequences, consistently resulted in nine well-supported major lineages. Although trees derived from individual datasets had different topologies (Figs 4–6), they were not significantly incongruent with the trees obtained from the combined analyses (Fig. 7). This was indicated by the fact that most major lineages found in the combined analyses were present in trees resulting from individual datasets. Only few exceptions were observed in the cases of 60S and LSU datasets. In one exceptional case, the 60S dataset (Fig. 5) showed Lineage 6 as split into two clades. In another case, the LSU tree (Fig. 4) depicted lineage 5 as not being monophyletic, although isolates belonging to this lineage still grouped relatively close to each other. Neither of these placements, however, was supported by phylogenetic statistics. Among the three gene regions used, MCM7 proved to be the most informative and resulted in trees with topologies similar to those obtained from the combined dataset.

Fig. 7

Bayesian phylogram derived from analyses of the concatenated dataset (60S, LSU and MCM7) containing 2 237 characters, of which 735 were parsimony informative. Thick branches represent BI posterior probabilities ≥95 %. Bootstrap support values ≥70 % are indicated at nodes as MP/ML. * = no bootstrap support or bootstrap support values <70 %.

Fig. 5

RAxML phylogram derived from analyses of the aligned 60S dataset containing 711 characters, of which 258 were parsimony informative. Thick branches represent BI posterior probabilities ≥95 %. Bootstrap support values ≥70 % are indicated at nodes as MP/ML. * = no bootstrap support or bootstrap support values <70 %.

Fig. 4

Bayesian phylogram derived from analyses of the aligned LSU dataset containing 898 characters, of which 164 were parsimony informative. Thick branches represent BI posterior probabilities ≥95 %. Bootstrap support values ≥70 % are indicated at nodes as MP/ML. * = no bootstrap support or bootstrap support values <70 %.

The first of the nine lineages (Figs 4–7), representing the largest number of species, included C. fimbriata (type species of Ceratocystis) and 31 other species previously included in the C. fimbriata complex. The second lineage included CMW 22736, representing T. thielavioides (type species for Chalaropsis), T. ovoidea, and two isolates from the USA and Belgium, previously described as T. thielavioides, but clearly distinct from CMW 22736. These two isolates are thus referred to as Chalaropsis sp. 1. The third lineage included C. coerulescens, type species for Endoconidiophora, and seven species previously considered part of the C. coerulescens complex. Isolates representing C. virescens, C. eucalypti, T. australis and T. neocaledoniae represented the fourth lineage, which did not include a type species of a previously described genus. Lineage 5 was previously referred to as the C. paradoxa complex, and included C. ethacetica (type species of Thielaviopsis), C. euricoi, C. musarum, C. radicicola and the recently described species, C. cerberus. The sixth lineage was the second largest and included C. moniliformis s. str. and 17 other species, but contained no type species representing a previously described genus. Two new species that are currently being described (Mbenoun et al., unpubl. data) grouped in this lineage, and were labelled according to provisional species names provided by M. Mbenoun (unpublished), namely Huntiella chlamydospora nom. prov. and H. pycnanthi nom. prov. Isolates of Ambrosiella xylebori (type species for Ambrosiella), A. hartigii and A. beaveri formed a distinct lineage. The last two lineages comprised Knoxdaviesia and Graphium species used as outgroups in all analyses.

Five of the 79 species in Ceratocystis s. l. were not accommodated in any of the nine major lineages discussed above (Figs 4–7). Ceratocystis adiposa and C. major had identical sequences in ITS (data not shown), LSU and 60S, and formed a distinct clade that was most closely related to lineage 7 (representing Ambrosiella). Ceratocystis fagacearum and A. ferruginea, although significantly different from each other, formed a clade of their own separating them from other Ceratocystis and Ambrosiella lineages. The fifth species, T. basicola, formed a unique lineage distinct from, but related to species in lineage 2 as its closest relatives.

Generic descriptions and nomenclator

Phylogenetic data generated in this study revealed seven well-supported lineages in Ceratocystis s. l. The distinction between these lineages is also supported by morphological and ecological data for the species in these groups. These lineages are, therefore, treated as distinct genera. Five of the lineages incorporate the type species of earlier described genera, and we thus emend the descriptions of Ambrosiella, Ceratocystis s. str., Chalaropsis, Endoconidiophora, and Thielaviopsis, based on the types and other species accommodated in the lineages. Two lineages for which existing names are not available are treated as novel genera, described here as Davidsoniella and Huntiella. Where necessary, new combinations are provided for the names of species in these genera. Species previously treated in Ceratocystis, but excluded from the newly defined genera in the Ceratocystidaceae (Tables 2 and 3), invalidly described species (Table 4), and homonyms from kingdoms other than the Fungi (Table 5), are not treated in the nomenclator, but listed in the tables as indicated.

Table 4

Species described invalidly in Ceratocystis, but for which validation is possible. More details on each species are presented by De Beer .

Name in Ceratocystis	Basionym	Reason for invalidity
2C. antennaroidospora Roldan	Ceratocystis antennaroidospora Roldan	Art. 40.1
2C. asteroides Roldan	Ceratocystis asteroides Roldan	Art. 40.1
1C. chinensis G.H. Zhao	Ceratocystis chinensis G.H. Zhao	Art. 40.1, 40.6
2C. heveae G.H. Zhao	Ceratocystis heveae G.H. Zhao	Art. 40.6
2C. jezoensis Aoshima	Ceratocystis jezoensis Aoshima	Art. 29.1 & 36.1
1C. kubanica (Sczerbin-Parfenenko) Potlajchuk	Ophiostoma kubanicum Sczerbin-Parfenenko	Art. 36.1
1C. minor (Hedgc.) J. Hunt var. barrasii J.J. Taylor	Ceratocystis minor (Hedgc.) J. Hunt var. barrasii J.J. Taylor	Art. 40.1
2C. pidoplichikovii Milko	Ceratocystis pidoplichikovii Milko	Art. 40.1
2Thielaviopsis wallemiiformis Dominik & Ihnat.	Thielaviopsis wallemiiformis Dominik & Ihnat.	Art. 40.1

Species that most likely belong in the Ophiostomatales and to be excluded from Ceratocystis s.l. upon validation.

Species that have thielaviopsis-like asexual states and probably belong to genera in the Ceratocystidaceae.

Table 5

Species names from the invertebrate fossil genus, Ceratocystis Jaekel (Echinodermata, Stylophora). Although the application of these names to fungal species are permissible because they are dictated by a different nomenclatural Code, their use should preferably be avoided (De Beer ).

Species
Ceratocystis perneri Jaekel
Ceratocystis prosthiakida Rahman, Zamora & Geyer
Ceratocystis spinosa Ubaghs
Ceratocystis vizcainoi Ubaghs

Brader ex Arx & Hennebert, Mycopath. Mycol. Appl. 25: 314. 1965.

?= Phialophoropsis L.R. Batra, Mycologia 59: 1008. 1967. (type species Ph. trypodendri).

Type species: Ambrosiella xylebori Brader ex Arx & Hennebert, Mycopath. Mycol. Appl. 25: 314. 1965.

Sexual state not known. Conidiophores phialidic, single to aggregated in sporodochia, hyaline, unbranched or sparingly branched, one-celled to septate. Conidia formed in chains or as terminal aleurioconidia.

Notes: We followed the emended generic description for Ambrosiella by Harrington , who restricted the genus to those species belonging to the Microascales. DNA sequence data is not available for A. trypodendri, type species of Phialophoropsis, which means the synonymy of the latter genus with Ambrosiella cannot be confirmed for the present. All known Ambrosiella species are associates of ambrosia beetles.

Six, Z.W. de Beer & W.D. Stone, Antonie van Leeuwenhoek 96: 23. 2009.

Note: Sexual state unknown.

L.R. Batra, Mycologia 59: 998. 1967.

Note: Sexual state unknown.

T.C. Harr. & McNew, Mycologia 106: 841. 2014.

Notes: Sexual state unknown. Sequences of this newly described species were not included in our analyses, but Harrington clearly showed that this species groups within Ambrosiella.

(L.R. Batra) T.C. Harr., Mycotaxon 111: 355. 2010

Basionym: Phialophoropsis trypodendri L.R. Batra, Mycologia 59: 1008. 1967.

Notes: Sexual state unknown. Ambrosiella trypodendri is the type species of Phialophoropsis (Batra 1967). No cultures are available for this species. However, Harrington argued that it is morphologically similar to Ambrosiella and provided a new combination for it. Seifert has examined the type, and made a drawing from it that was used to represent this species in The Genera of Hyphomycetes (Seifert ).

Brader ex Arx & Hennebert, Mycopath. Mycol. Appl. 25: 314. 1965.

Descriptions: Von Arx & Hennebert (1965: 312–315, fig. 2); Batra (1967: 990–992, figs 14–19).

Notes: Sexual state unknown. The genus and species were invalidly described by Brader (1964) (Art. 40.1), but Von Arx & Hennebert (1965) redescribed and validated both.

Ellis & Halst., In: Halsted, New Jersey Agric. Coll. Exp. Sta. Bull. 76: 14. 1890.

?= Rostrella Zimm., Meded. Lands Plantentuin, Batavia 37: 24. 1900. (nom. illegit., Art. 53.1, later homonym for Rostrella Fabre, Ann. Sci. Nat., Bot. 6, 9: 66. 1879) (type species Ro. coffeae).

= Ceratocystis Ellis & Halst. section Ceratocystis pro parte, In: Upadhyay, Monogr. Ceratocystis & Ceratocystiopsis: 32. 1981.

Type species: Ceratocystis fimbriata Ellis & Halst., New Jersey Agric. Coll. Exp. Sta. Bull. 76: 14. 1890.

Emended generic diagnosis. Ascomatal bases globose, brown to black, unornamented or with undifferentiated ornamental hyphae. Ascomatal necks long, tapering to apex, straight, dark-brown to black, hyaline at apex. Ostiolar hyphae divergent, non-septate, tapered, light brown to hyaline. Asci dehiscent. Ascospores one-celled, hat-shaped, hyaline, accumulating in cream-coloured masses at tips of necks. Primary conidiophores phialidic, flask-shaped. Secondary conidiophores flaring or wide-mouthed. Primary conidia cylindrical, hyaline. Secondary conidia barrel to subglobose shaped, hyaline to light brown. Aleurioconidia globose, ovoid to pyriform, singly or in chains, pale-brown to brown.

Notes: The most characteristic features of this genus are the ascomatal bases lacking distinct ornamentations and hat-shaped ascospores. The possibility that Ro. coffeae might not be a synonym of C. fimbriata is discussed under the latter species, below. However, even if the two species are distinct, the species from coffee would probably still group in Ceratocystis s. str., which means Rostrella will remain a synonym of Ceratocystis.

Tarigan & M. van Wyk, S. Afr. J. Bot. 77: 301. 2011.

M.J. Wingf., De Beer & M.J. Morris, Syst. Appl. Microbiol. 19: 196. 1996. (as “albofundus”).

M. van Wyk & M.J. Wingf., Australas. Pl. Pathol. 36: 411. 2007.

Engelbr. & T.C. Harr., Mycologia 97: 64. 2005.

Note: In earlier studies this species was treated as residing in the Latin American “cacao” population of C. fimbriata (Baker Engelbrecht ).

J.A. Johnson & T.C. Harr., Mycologia 97: 1086. 2005.

M. van Wyk & M.J. Wingf., Fungal Diveristy 40: 111. 2010.

Kamgan & Jol. Roux, Antonie van Leeuwenhoek 101: 237. 2012.

M. van Wyk & M.J. Wingf., Fungal Diversity 46: 122. 2011.

M. van Wyk & M.J. Wingf., Fungal Diversity 46: 125. 2011.

M. van Wyk & M.J. Wingf., Fungal Diversity 46: 122. 2011.

M. van Wyk & M.J. Wingf., IMA Fungus 3: 54. 2012.

Kajitani & Masuya, Mycoscience 52: 351. 2011.

Ellis & Halst., New Jersey Agric. Coll. Exp. Sta. Bull. 76: 14. 1890.

≡ Sphaeronaema fimbriatum (Ellis & Halst.) Sacc., Syll. Fung. 10: 125. 1892.

≡ Ceratostomella fimbriata (Ellis & Halst.) Elliott, Phytopathology 13: 56. 1923.

≡ Ophiostoma fimbriatum (Ellis & Halst.) Nannf., Svenska Skogsv.-Fören. Tidskr. 32: 408. 1934.

≡ Endoconidiophora fimbriata (Ellis & Halst.) R.W. Davidson, J. Agric. Res. 50: 800. 1935.

?= Rostrella coffeae Zimm., Meded. Lands Plantentuin, Batavia 37: 32. 1900.

≡ Ophiostoma coffeae (Zimm.) Arx, Antonie van Leeuwenhoek 18: 210. 1952.

≡ Ceratocystis moniliformis f. coffeae (Zimm.) C. Moreau, Bull. Sci. Minist. France Outre-Mer 5: 424. 1954.

Descriptions: Davidson (1935: 799–800); Hunt (1956: 11–16); Webster & Butler (1967: 1459–1463, pl. I-VI); Griffin (1968: 703); Morgan-Jones (1967a, figs A–G); Olchowecki & Reid (1974: 1699, pl. XIII, fig. 258); Matsushima (1975: 169, pl. 382, 383); Nag Raj & Kendrick (1975: 118, 141, fig. 45); Upadhyay (1981: 44, figs 69–72); Potlajczuk & Schekunova (1985: 150); Engelbrecht & Harrington (2005: 63–64).

Notes: The original description of Ceratocystis fimbriata was from sweet potato in the USA. Analyses of DNA sequences have shown that the fungus treated as C. fimbriata in various studies and from various countries and host plants, represent a species complex that includes many different cryptic species (Van Wyk ). The name C. fimbriata should be restricted to the fungus from sweet potato and to other isolates belonging to the same phylogenetic species.

Pontis (1951) listed Rostella coffeae as a synonym of C. fimbriata, but mentioned biological differences between isolates from the coffee tree and sweet potato. Several recent studies, based on DNA sequence comparisons for multiple gene regions, have distinguished host-specific and geographically-separated populations, including populations from coffee, in the C. fimbriata species complex (Harrington 2000, Barnes ). Van Wyk described two of these host-specific groups from coffee in Colombia as new species, but did not consider the possibility that one of them might represent R. coffeae, probably because the latter was originally described from coffee in Java (Indonesia). For the present we treat R. coffeae as a synonym of C. fimbriata until future studies with fresh isolates from coffee in Java provide further insights into this question.

M. van Wyk & M.J. Wingf., Fungal Diversity 34: 180. 2009.

Z.W. de Beer & M.J. Wingf., CBS Biodiversity Series 12: 291. 2013.

≡ Ceratocystis populicola J.A. Johnson & T.C. Harr., Mycologia 97: 1084. 2005. (nom. illegit., Art 53.1).

Notes: Johnson described this species validly, but the name was a later homonym for Ceratocystis populicola Olchow. & J. Reid (= Ophiostoma populicola) and thus illegitimate. De Beer provided a new, legitimate name.

M. van Wyk & M.J. Wingf., Persoonia 22: 80. 2009.

M. van Wyk & M.J. Wingf., Mycotaxon 117: 395. 2011.

M. van Wyk, Al Adawi & M.J. Wingf., Fungal Diversity 27: 224. 2007.

M. van Wyk & M.J. Wingf., Mycotaxon 117: 397. 2011.

M. van Wyk, Jol. Roux & C. Rodas, Fungal Diversity 28: 80. 2008.

R.N. Heath & Jol. Roux, Fungal Diversity 34: 57. 2009.

M. van Wyk & M.J. Wingf., Fungal Diveristy 40: 112. 2010.

I. Barnes & M. J. Wingf., Mycologia 95: 867. 2003.

= Ceratocystis zombamontana R.N. Heath & Jol. Roux, Fungal Diversity 34: 53. 2009.

Note: Kamgan Nkuekam showed C. zombamontana to be a synonym of C. pirilliformis based on DNA sequence data.

(Walter) Engelbr. & T.C. Harr., Mycologia 97: 65. 2005.

Basionym: Endoconidiophora fimbriata f. platani Walter, Phytopathology 42: 236. 1952.

Note: This species was considered to represent a population of C. fimbriata from sycamore (Platanus) (Santini & Capretti 2000, Barnes ), until Engelbrecht & Harrington (2005) elevated it to species level.

M. van Wyk, M.J. Wingf. & E.C.Y. Liew, Stud. Mycol. 50, 278. 2004.

R.N. Heath & Jol. Roux, Fungal Diversity 34: 53. 2009.

J.A. Johnson & T.C. Harr., Mycologia 97: 1088. 2005.

R.N. Heath & Jol. Roux, Fungal Diversity 34: 56. 2009.

M. Mbenoun & Jol. Roux, Mycol. Progress 13: 234. 2014.

Kamgan & Jol. Roux, Fungal Diversity 29: 50. 2008.

(R.W. Davidson) C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 22. 1952. emend. J.A. Johnson & T.C. Harr., Mycologia 97: 1083. 2005.

Basionym: Endoconidiophora variospora R.W. Davidson, Mycologia 36: 303. 1944.

≡ Ophiostoma variosporum (R.W. Davidson) Arx, Antonie van Leeuwenhoek 18: 212. 1952.

≡ Ceratocystis moniliformis f. variospora C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 23. 1952. (nom. inval., Art. 39.1).

Descriptions: Hunt (1956: 16–18); Johnson : 1082–1084, figs 8–16).

Notes: Ceratocystis variospora was invalidly reduced to a forma of C. moniliformis by Moreau (1952). Webster & Butler (1967), Upadhyay (1981), and Seifert all treated C. variospora as synonym of C. fimbriata. Johnson re-instated it as a distinct species in the C. fimbriata complex based on phylogenetic analyses.

M. Mbenoun & Jol. Roux, Mycol. Progress 13: 235. 2014.

Peyronel, Le Staz. Sper. agric. 49: 595. 1916.

Type species: Chalaropsis thielavioides Peyronel, Le Staz. Sper. agric. 49: 58. 1916.

Emended generic diagnosis. Sexual state not observed. Conidiophores arise laterally from vegetative hyphae. Conidiogenous cells phialidic, cylindrical, tapering toward apex, hyaline, subhyaline or pale brown. Conidia unicellular, cylindrical with rounded or truncate ends, hyaline to light brown, singly or in chains. Aleurioconidia unicellular, globose, ellipsoidal, ovoid, or pyriform with truncate ends, solitary and terminal on sympodially branching conidiophores, pale brown to brown.

Note: The morphological characters of Chalaropsis species are indistinguishable from those of the asexual states of Ceratocystis s. str.

(Nag Raj & W.B. Kendr.) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810308.

Basionym: Chalara ovoidea Nag Raj & W.B. Kendr., Monogr. Chalara: 127. 1975.

≡ Thielaviopsis ovoidea (Nag Raj & W.B. Kendr.) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 70. 2002.

Description: Nag Raj & Kendrick (1975: 116, 127–128, figs 43B).

Notes: Two isolates of this species were included in our analyses, none of which represented the type. Epitypification of this name is needed.

(Kiffer & Delon) Z.W. de Beer, T.A. Duong & M.J. Wingf.

Basionym: Chalara populi Kiffer & Delon, Mycotaxon 18: 171. 1983. (as “Veldeman ex”).

≡ Thielaviopsis populi (Kiffer & Delon) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 70. 2002.

= Chalaropsis populi Veldeman, Meded. Fac. Landbouwwetensch. Rijksuniv. Gent 36: 1001. 1971. (nom. inval., Art. 39.1, 40.1).

Description: Kiffer & Delon (1983: 171–172, figs 1–2).

Notes: Veldeman (1971) did not provide a formal diagnosis in the description of this species. Kiffer & Delon (1983) studied the original material and validated the name. No isolates representing Chalaropsis populi were available for the present study, but sequence data from previous studies confirm its placement in this genus (Wingfield ).

Note: Two isolates included in this study that had been labelled as T. thielavioides in the CBS collection, emerged as repesenting an undescribed species in this genus, distinct from Chalaropsis thielavioides.

Peyronel, Le Staz. Sper. agric. 49: 58. 1916.

≡ Chalara thielavioides (Peyronel) Nag Raj & W.B. Kendr., Monogr. Chalara: 136. 1975.

≡ Thielaviopsis thielavioides (Peyronel) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 70. 2002.

= Chalaropsis thielavioides Peyronel var. ramosissima Sugiyama, J. Fac. Sci. Univ. Tokyo 10: 33. 1968.

Description: Nag Raj & Kendrick (1975: 117, 136–137, fig. 44).

Note: Nag Raj & Kendrick (1975) considered Ch. thielavioides var. ramosissima a synonym of Ch. thielavioides.

Z.W. de Beer, T.A. Duong & M.J. Wingf., gen. nov. MycoBank MB810235.

Etymology: Named after Ross Wallace Davidson who described at least 41 ophiostomatoid species during his career at the USDA. These included Ceratocystis virescens, the type species for this new genus.

Type species: Davidsoniella virescens (R.W. Davidson) Z.W. de Beer, T.A. Duong & M.J. Wingf.

Ascomatal bases globose, light brown to dark brown to black, ornamental hyphae, simple, septate, stiff. Ascomatal necks long, dark brown at base to light brown at apex. Ostiolar hyphae divergent to straight, non-septate, smooth, light brown to hyaline. Asci dehiscent. Ascospores one-celled, elongate, narrow fusiform to spindle shaped, slightly curved, with thick, hyaline sheath. Conidiophores arise laterally from vegetative hyphae. Conidiogenous cells phialidic, cylindrical, hyaline. Conidia unicellular, cylindrical with flattened ends, barrel-shaped, hyaline, borne in chains of varying length. Aleurioconidia not present.

Note: The most distinctive features of this genus are elongated, spindle-shaped and sheathed ascospores that are substantially longer than those of Endoconidiophora spp.

(J. Walker & Kile) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810241.

Basionym: Chalara australis J. Walker & Kile, Austral. J. Bot. 35: 7. 1987. (non Chalara australis McKenzie, Mycotaxon 46: 291 (1993), nom. illegit.).

≡ Thielaviopsis australis (J. Walker & Kile) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 69. 2002.

Note: Sexual state unknown.

(Z.Q. Yuan & Kile) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810309.

Basionym: Ceratocystis eucalypti Z.Q. Yuan & Kile, Mycol. Res. 100: 573. 1996.

≡ Chalara eucalypti Z.Q. Yuan & Kile, Mycol. Res. 100: 573. 1996.

≡ Thielaviopsis eucalypti (Z.Q. Yuan & Kile) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 69. 2002.

(Kiffer & Delon) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810310.

Basionym: Chalara neocaledoniae Dadant ex Kiffer & Delon, Mycotaxon 18: 166. 1983.

≡ Thielaviopsis neocaledoniae (Kiffer & Delon) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 70. 2002.

= Thielaviopsis neocaledoniae Dadant, Rev. Gén. Bot. 57: 176. 1950. (nom. inval., Art. 39.1).

Description: Kiffer & Delon (1983: 166–170, figs 1–2).

Notes: Sexual state unknown. Dadant (1950) did not provide a Latin diagnosis and also failed to designate a type specimen, making the species name invalid. Kiffer & Delon (1983) obtained the original isolate of Dadant and validated the name.

(R.W. Davidson) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810311.

Basionym: Endoconidiophora virescens R.W. Davidson, Mycologia 36: 301. 1944.

≡ Ceratocystis virescens (R.W. Davidson) C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 22. 1952.

≡ Ophiostoma virescens (R.W. Davidson) Arx, Antonie van Leeuwenhoek 18: 212. 1952.

Description: Samuels (1993: 16, figs 1A–B).

Notes: Hunt (1956), Olchowecki & Reid (1974), and Upadhyay (1981) treated C. virescens as a synonym of C. coerulescens, but Nag Raj & Kendrick (1975), Gibbs (1993), Kile (1993), and Seifert , considered the two species distinct. Witthuhn confirmed the separateness of the species.

Münch, Naturw. Z. Forst- u. Landw. 5: 564. 1907.

= Ceratocystis Ellis & Halst. section Endoconidiophora (Münch) H.P. Upadhyay pro parte, In: Upadhyay, Monogr. Ceratocystis & Ceratocystiopsis: 64. 1981.

Type species: Endoconidiophora coerulescens Münch, Naturw. Z. Forst- u. Landw. 5: 564. 1907.

Emended generic diagnosis. Ascomatal bases globose to ovoid, dark brown, with distinct basal spines. Ascomatal necks long, tapering towards apex, dark brown to black. Ostiolar hyphae divergent, non-septate, hyaline. Asci dehiscent. Ascospores one-celled, elongate to slightly curved with round ends, oblong cylindrical, surrounded by dinstinct translucent sheath. Conidiophores tubular, rectangular, cylindrical, sometimes slightly flared collarette. Conidiogenous cells phialidic, oblong cylindrical. Conidia unicellular, rectangular with two attachment points, hyaline, in chains. Aleurioconidia not present.

Note: The most distinctive features of this genus are the long spines on the ascomatal bases and the sheathed ascospores (see Harrington & Wingfield 1998)

Münch, Naturw. Z. Land. Forstw. 5: 564. 1907.

≡ Ceratocystis coerulescens (Münch) Bakshi, Trans. Br. Mycol. Soc. 33: 114. 1950. emend. T.C. Harr. & M.J. Wingf., Canad. J. Bot. 76: 1448. 1998.

≡ Ophiostoma coerulescens (Münch) Nannf., Svenska Skogsv.-Fören. Tidskr. 32: 408. 1934.

?= Chalara ungeri Sacc., Syll. Fung. 4: 336. 1886.

≡ Thielaviopsis ungeri (Sacc.) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 70. 2002.

Descriptions: Lagerberg : 196–203, figs 22–26); Davidson (1935: 798–799); Siemaszko (1939: 20–22, pl. I, figs 9–13); Bakshi (1951: 2–5); Hunt (1956: 17, 21–23); Griffin (1968: 700–701); Nag Raj & Kendrick (1975: 94, 138–139, fig. 32B); Upadhyay (1981: 65, figs 191–196); Potlajczuk & Schekunova (1985: 149–150); Harrington & Wingfield (1998: 1448–1449).

Notes: Harrington & Wingfield (1998) designated a neotype for C. coerulescens, while Nag Raj & Kendrick (1975) did the same for Ca. ungeri. Nag Raj & Kendrick (1975) and Paulin-Mahady accepted the suggestion by Münch (1907) that Ca. ungeri represented the asexual state of C. coerulescens. In the absence of an ex-type culture representing Ca. ungeri, the synonymy can neither be confirmed nor rejected. Witthuhn showed that isolates identified as C. coerulescens formed three distinct clades based on ITS data. These were later described as C. coerulescens sensu stricto, C. pinicola, and C. resinifera (Harrington & Wingfield 1998).

(R.W. Davidson) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810312.

Basionym: Endoconidiophora coerulescens f. douglasii R.W. Davidson, Mycologia 45: 584. 1953.

≡ Ceratocystis douglasii (R.W. Davidson) M.J. Wingf. & T.C. Harr., Canad. J. Bot. 75: 832. 1997

Notes: Upadhyay (1981) considered Endoconidiophora coerulescens f. douglasii a synonym of C. coerulescens. Wingfield distinguished C. coerulescens from C. douglasii and elevated the latter to species level.

(M.J. Wingf., Yamaoka & Marin) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810313.

Basionym: Ceratocystis fujiensis M.J. Wingf., Yamaoka & Marin, Mycol. Res. 109: 1142. 2005.

(Redfern & Minter) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810314.

Basionym: Ceratocystis laricicola Redfern & Minter, Pl. Pathol. 36: 468. 1987.

Descriptions: Harrington & Wingfield (1998: 1453, 1456); Yamaoka : 369–371, figs 6–10); Marin : 1142, 1144).

Note: Witthuhn and Harrington distinguished C. laricicola from the morphologically similar C. polonica based on differences in bark beetle associates, conifer hosts and molecular data.

(T.C. Harr. & M.J. Wingf.) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810315.

Basionym: Ceratocystis pinicola T.C. Harr. & M.J. Wingf., Canad. J. Bot. 76: 1452. 1998.

(Siemaszko) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810316.

Basionym: Ophiostoma polonicum Siemaszko, Planta Pol. 7: 32. 1939.

≡ Ceratocystis polonica (Siemaszko) C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 22. 1952. emend. T.C. Harr. & M.J. Wingf., Canad. J. Bot. 76(8): 1452. 1998. (as “polonicum”).

Descriptions: Siemaszko (1939: 20, 32–33, pl. IV, figs 1–10); Mathiesen (1951: 208–210); Hunt (1956: 27); Solheim (1986: 205–206); Yamaoka : 1217–1219); Harrington & Wingfield (1998: 1452–1453, 1455); Marin : 1142, 1144).

Notes: Siemaszko (1939) erroneously linked a leptographium-like asexual state to O. polonicum. Upadhyay (1981) therefore treated the species as a synonym of O. penicillatum. However, Solheim (1986) and Harrington (1988) distinguished the two species. Visser confirmed the placement of the species in Ceratocystis based on DNA sequence data. Harrington & Wingfield (1998) designated a neotype. Ceratocystis polonica was distinguished from the morphologically similar C. laricicola by Witthuhn and Harrington , based primarily on differences in conifer hosts and bark beetle associates. Marin showed that European and Japanese populations of C. polonica are genetically isolated and possibly in the process of speciation.

(T.C. Harr. & M.J. Wingf.) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810337.

Basionym: Ceratocystis resinifera T.C. Harr. & M.J. Wingf., Canad. J. Bot. 76: 1449. 1998.

(M.J. Wingf., T.C. Harr. & H. Solheim) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810317.

Basionym: Ceratocystis rufipennis M.J. Wingf., T.C. Harr. & H. Solheim, Canad. J. Bot. 75: 828. 1997. (as “rufipenni”).

Z.W. de Beer, T.A. Duong & M.J. Wingf., gen. nov. MycoBank MB810236.

Etymology: Named after the late John Hunt, author of the monograph of Ceratocystis that was published in 1956 (Hunt 1956) and in honour of the major contribution he made to the taxonomy of this group of fungi during his short career.

Type species: Huntiella moniliformis (Hedgc.) Z.W. de Beer, T.A. Duong & M.J. Wingf.

Ascomatal bases globose to pyriform, black, ornamented with dark brown to black, conical spines, occasionaly septate. Ascomatal necks long, tapering to apex, black, with a disk-like base. Ostiolar hyphae convergent to divergent, hyaline. Asci dehiscent. Ascospores one-celled, hat-shaped, hyaline. Primary conidiophores phialidic, long, septate, tapering to tip. Secondary conidiophores phialidic, short, septate. Primary conidia cylindrical, truncate ends, hyaline, in long chains. Secondary conidia barrel-shaped, hyaline to pale brown. Aleurioconidia not observed.

Note: The most distinctive features of this genus are the conical spines on the ascomatal bases, the disk-like bases of the ascomatal necks, and the hat-shaped ascospores.

(M. van Wyk, M.J. Wingf. & T. Kirisits) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810318.

Basionym: Ceratocystis bhutanensis M. van Wyk, M.J. Wingf. & T. Kirisits, Stud. Mycol. 50: 373. 2004.

Note: This is an unusual taxon because it is the only species associated with a bark beetle, Ips smutzenhoferi, that infests Pinus wallichiana in Bhutan (Kirisits ).

(R.N. Heath & Jol. Roux) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810319.

Basionym: Thielaviopsis ceramica R.N. Heath & Jol. Roux, Fungal Diversity 34: 60.

Note: Sexual state unknown.

(S.F. Chen, M. van Wyk, M.J. Wingf. & X.D. Zhou) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810320.

Basionym: Ceratocystis chinaeucensis S.F. Chen, M. van Wyk, M.J. Wingf. & X.D. Zhou, Fungal Diversity 58: 274. 2013.

(Mbenoun & Jol. Roux) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810321.

Basionym: Ceratocystis cryptoformis Mbenoun & Jol. Roux, Mycol. Progress 13: 232. 2014.

Note: Although an isolate of this species was not included in the present study, DNA sequences generated by Mbenoun undoubtedly place this species in Huntiella.

(Kamgan & Jol. Roux) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810322.

Basionym: Ceratocystis decipiens Kamgan & Jol. Roux, Austral. Pl. Pathol. 42: 299. 2013.

(Tarigan, M. van Wyk & M.J. Wingf.) Z.W. de Beer, T.A. Duong & M.J. Wingf. comb. nov. MycoBank MB810323.

Basionym: Ceratocystis inquinans Tarigan, M. van Wyk & M.J. Wingf., Mycoscience 51: 58. 2010.

(Tarigan, M. van Wyk & M.J. Wingf) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810324.

Basionym: Ceratocystis microbasis Tarigan, M. van Wyk & M.J. Wingf., Mycoscience 51: 61. 2010.

(Hedgc.) Z.W. de Beer, T.A. Duong & M.J. Wingf. comb. nov. MycoBank MB810325.

Basionym: Ceratostomella moniliformis Hedgc., Annual Rep. Missouri Bot. Gard.17: 78. 1906.

≡ Ophiostoma moniliforme (Hedgc.) Syd., In Sydow & Sydow, Ann. Mycol. 17: 43. 1919.

≡ Endoconidiophora moniliformis (Hedgc.) R.W. Davidson, J. Agric. Res. 50: 800. 1935.

≡ Ceratocystis moniliformis (Hedgc.) M. Moreau & Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 141. 1952. (nom. illegit., Art. 52.1).

≡ Ceratocystis moniliformis (Hedgc.) C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 22. 1952.

= Endoconidiophora bunae Kitajima, Bull. Imp. Forest Exp. Sta. 35: 126. 1936.

≡ Ophiostoma bunae (Kitajima) Arx, Antonie van Leeuwenhoek 18: 211. 1952. (as “lunae”).

≡ Ceratocystis bunae (Kitajima) C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 22. 1952.

= Ceratocystis wilsonii Bakshi, Mycol. Pap. 35: 8. 1951. (as “wilsoni”).

≡ Ceratocystis moniliformis f. wilsonii C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 23. 1952. (as “wilsoni”; nom. inval., Art. 39.1).

= Ophiostoma moniliforme f. davidsonii Luc, Rev. Mycol. (Paris) Suppl. Col. 17: 12. 1952. (nom. inval., Art. 39.1).

= Ophiostoma moniliforme f. pycnanthi Luc, Rev. Mycol. (Paris) Suppl. Col. 17: 12. 1952. (nom. inval., Art. 39.1).

= Ophiostoma moniliforme f. typica Luc, Rev. Mycol. (Paris) Suppl. Col. 17: 12. 1952. (nom. inval., Art. 24.3 & Art. 39.1).

= Ophiostoma moniliforme f. theobromae Luc, Rev. Mycol. (Paris) Suppl. Col. 17: 13. 1952. (nom. inval., Art. 39.1).

= Ceratocystis filiformis Roldan, Philipp. J. Sci. 91: 418. 1962.

Descriptions: Hedgcock (1906: 78–80, pl. 3, fig. 5, pl. 5, figs 3–5); Davidson (1935: 799–800); Moreau & Moreau (1952, figs 1–4); Luc (1952: 12–15, figs 1–2); Hunt (1956: 13, 17–19); Morgan-Jones (1967b, figs A–H); Nag Raj & Kendrick (1975: 116, 141–142, fig. 43A); Upadhyay (1981: 51, figs 109–115); Maekawa : 8–10, figs 7–18); Kowalski & Butin (1989: 238–241).

Notes: Four varieties were described invalidly for C. moniliformis by Luc (1952). Moreau (1952) then reduced two species, C. wilsonii and C. variospora (now considered a distinct species), to formae of C. moniliformis, and treated R. coffeae as a synonym (see notes above under C. fimbriata). Moreau & Moreau (1952) reduced O. moniliforme f. theobromae to synonymy with C. moniliformis, and Hunt (1956) did the same with E. bunae and C. wilsonii. Nag Raj & Kendrick (1975) and Upadhyay (1981) added C. filiformis to the synonyms of C. moniliformis. The accuracy of all these synonymies deserves to be carefully reconsidered based on DNA sequence data and fresh isolates obtained from similar hosts.

(Yuan & Mohammed) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810326.

Basionym: Ceratocystis moniliformopsis Yuan & Mohammed, Austral. Syst. Bot. 15: 126. 2002.

(R.N. Heath & Jol. Roux) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810328.

Basionym: Ceratocystis oblonga R.N. Heath & Jol. Roux, Fungal Diversity 34: 59. 2009.

(Al-Subhi, M.J. Wingf., M. van Wyk & Deadman) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810329.

Basionym: Ceratocystis omanensis Al-Subhi, M.J. Wingf., M. van Wyk & Deadman, Mycol. Res. 110: 242. 2006.

(Kamgan & Jol. Roux) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810330.

Basionym: Ceratocystis salinaria Kamgan & Jol. Roux, Austral. Pl. Pathol. 42: 298. 2013.

(Kamgan & Jol. Roux) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810331.

Basionym: Ceratocystis savannae Kamgan & Jol. Roux, Fungal Diversity 29: 52. 2008.

(M. van Wyk & M.J. Wingf.) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810332.

Basionym: Ceratocystis sublaevis M. van Wyk & M.J. Wingf., Fungal Diversity 46: 128. 2011.

(Tarigan, M. van Wyk & M.J. Wingf.) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810333.

Basionym: Ceratocystis sumatrana Tarigan, M. van Wyk & M.J. Wingf., Mycoscience 51: 60. 2010.

(M. van Wyk & M.J. Wingf.) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810334.

Basionym: Ceratocystis tribiliformis M. van Wyk & M.J. Wingf., Fungal Diversity 21: 197. 2006.

(Kamgan & Jol. Roux) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810335.

Basionym: Ceratocystis tyalla Kamgan & Jol. Roux, Antonie van Leeuwenhoek 101: 233. 2012.

Went, Meded. Proefstat. Suikerriet W. Java 5: 4. 1893.

= Hughesiella Bat. & A.F. Vital, Anais Soc. Biol. Pernambuco 14: 141. 1956. (type species Hu. euricoi).

Type species: Thielaviopsis ethacetica Went, Meded. Proefstat. Suikerriet W. Java 5: 4. 1893.

Emended generic diagnosis. Ascomatal bases globose, light brown, display dark as result of aleurioconidia and distinctly digitate or stellate appendages. Ascomatal necks long, tapering to apex, dark grey. Ostiolar hyphae divergent, hyaline. Asci dehiscent. Ascospores aseptate, ellipsoidal, hyaline with sheath. Conidiophores lageniform, solitary, occasionaly aggregate in synnemata. Primary conidia aseptate, cylindrical, hyaline. Secondary conidia aseptate, cylindrical to oblong, hyaline becoming grey, thick walled. Aleurioconidia subglobose, oblong or ovoid, thick-walled, forms holoblastically, singly or in chains, grey-brown.

Notes: The most distinctive features of this genus are the distinctly digitate or stellate appendages on the ascomatal bases. This is the only group where some species form synnemata in the asexual state. Thielaviopsis euricoi is the type species of the genus Hughesiella, which is thus a synonym of Thielaviopsis.

(Mbenoun, M.J. Wingf. & Jol. Roux) Z.W. de Beer, T.A. Duong & M.J. Wingf., comb. nov. MycoBank MB810336.

Basionym: Ceratocystis cerberus Mbenoun, M.J. Wingf. & Jol. Roux, Mycologia 106: 778. 2014.

Went, Meded. Proefstat. Suikerriet W. Java 5: 4. 1893. (as “ethaceticus”).

≡ Ceratocystis ethacetica (Went) Mbenoun & Z.W. de Beer, Mycologia 106: 772. 2014.

= Endoconidium fragrans Delacr., Bull. Soc. Mycol. France 9: 184. 1893.

= Catenularia echinata Wakker, De ziekten van het suikerriet op Java, E.J. Brill, Leiden: 196. 1898.

Descriptions: Mbenoun .

Note: The synonymy between T. ethacetica, Cat. echinata and E. fragrans is discussed by Mbenoun .

(Bat. & A.F. Vital) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 70. 2002.

Basionym: Hughesiella euricoi Bat. & A.F. Vital, Anais Soc. Biol. Pernambuco 14: 142. 1956.

≡ Ceratocystis euricoi (Bat. & A.F. Vital) Mbenoun & Z.W. de Beer, Mycologia 106: 774. 2014.

Descriptions: Mbenoun .

Notes: Sexual state unknown. Thielaviopsis euricoi is the type species of the genus Hughesiella, treated above as synonym of Thielaviopsis.

(R.S. Mitchell) Riedl, Sydowia 15: 249. 1962.

Basionym: Thielaviopsis paradoxa (De Seynes) Höhn. var. musarum R.S. Mitchell, J. Council Sci. Industr. Res. Australia, 10: 130. 1937. (nom. inval., Art. 39.1).

≡ Ceratocystis musarum Riedl, Sydowia 15: 248. 1962.

Descriptions: Mbenoun .

Note: The taxonomy of this species is discussed by Mbenoun .

(De Seynes) Höhn., Hedwigia 43: 295. 1904.

Basionym: Sporoschisma paradoxum De Seynes, Rech. Hist. Nat. Veg. Inf. 3: 30. 1886.

≡ Chalara paradoxa (De Seynes) Sacc., Syll. Fung. 10: 595. 1892.

≡ Ceratostomella paradoxa (De Seynes) Dade, Trans. Br. Mycol. Soc. 13: 191. 1928.

≡ Ophiostoma paradoxum (De Seynes) Nannf., Svenska Skogsv.-Fören. Tidskr. 32: 408. 1934.

≡ Endoconidiophora paradoxa (De Seynes) R.W. Davidson, J. Agric. Res. 50: 802. 1935.

≡ Ceratocystis paradoxa (De Seynes) C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 22. 1952.

= Stilbochalara dimorpha Ferd. & Winge, Bot. Tidsskr. 30: 220. 1910.

Descriptions: Davidson (1935: 801–802); Hunt (1956: 13, 19–20); Morgan-Jones (1967c, figs A–G); Nag Raj & Kendrick (1975: 112, 114, 128–129, figs 41–42); Upadhyay (1981: 67, figs 197–204); Mbenoun .

Notes: The synonymy of St. dimorpha with C. paradoxa was suggested by Mbenoun . These authors also discussed and explained the treatment of the names and authorities of the previously considered sexual and asexual states, as suggested by Hawksworth .

(Hennebert) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 70. 2001.

Basionym: Chalaropsis punctulata Hennebert, Antonie van Leeuwenhoek 33: 334. 1967.

= Ceratostomella radicicola Bliss, Mycologia 33: 468. 1941.

≡ Ophiostoma radicicolum (Bliss) Arx, Antonie van Leeuwenhoek 18: 211. 1952.

≡ Ceratocystis radicicola (Bliss) C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 22. 1952.

Descriptions: Hunt (1956: 11, 17, 20); Nag Raj & Kendrick (1975: 106, 142, fig. 38); Upadhyay (1981: 69, figs 205–213); Mbenoun .

Notes: Paulin-Mahady and Mbenoun confirmed the synonymy of T. punctulata and C. radicicola based on similar sequences. Based on the Melbourne Code (McNeill ) the older epithet must take preference, implying that this species will in future be treated as T. punctulata, and not as the better known C. radicicola, unless conservation of the later name against the earlier is proposed and accepted.

Four species could not be consistently accommodated in any of the seven major clades for which genera have been provided. We believe that they represent discrete genera but we have not provided generic names for these lineages. With increased sampling and further study, additional species are likely to be found that will populate these clades. At that time, genera can be provided for them. For the present they have been retained in their existing genera. We also list C. erinaceus and C. norvegica here for which isolates could not be obtained, and for which the generic placements remains uncertain.

(Butler) C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 22. 1952.

Basionym: Sphaeronema adiposum Butler, Mem. Dept. Agric. India, Bot. Ser. 1: 40. 1906.

≡ Ceratostomella adiposa (Butler) Sartoris, J. Agric. Res. 35: 585. 1927.

≡ Ophiostoma adiposum (Butler) Nannf., In Melin & Nannf., Svenska Skogsv.-Fören. Tidskr. 32: 408. 1934.

≡ Endoconidiophora adiposa (Butler) R.W. Davidson, J. Agric. Res. 50: 802. 1935.

= Ceratostomella major J.F.H. Beyma, Zentrabl. Bakteriol., 2. Abt. 91: 348. 1935.

≡ Ophiostoma majus (J.F.H. Beyma) Goid., Boll. Staz. Patol. Veg. Roma 15: 158. 1935.

≡ Ceratocystis major (J.F.H. Beyma) C. Moreau, Rev. Mycol. (Paris) Suppl. Col. 17: 22. 1952.

Descriptions: Sartoris (1927: 578–585, figs 1–4); Davidson (1935: 801–802); Hunt (1956: 10–13); Upadhyay (1981: 35, figs 26–30); Moreau (1952: 17–20, fig. 1); Nag Raj & Kendrick (1975: 104, 140, fig. 37).

Notes: Hunt (1956), Moreau (1952), Griffin (1968), Olchowecki & Reid (1974), and Nag Raj & Kendrick (1975), all treated C. major and C. adiposa as distinct. Upadhyay (1981) suggested the synonymy of C. major with C. adiposa. Identical SSU sequences for the two species by Hausner suggested that the synonymy is sound, and this was confirmed in the present study where the two species had identical sequences in ITS, LSU and 60S.

(Berk. & Broome) Ferraris, Fl. Ital. Crypt., Fungi 1: 233. 1912.

Basionym: Torula basicola Berk. & Broome, Ann. Mag. Nat. Hist. 5: 461. 1850.

≡ Trichocladium basicola (Berk. & Broome) J.W. Carmich., In: Carmichael et al., Genera of Hyphomycetes: 185. 1980.

= Chalara elegans Nag Raj & W.B. Kendr., Monogr. Chalara: 111. 1975.

Description: Nag Raj & Kendrick (1975: 108–113, figs 39–40 as Ca. elegans).

Notes: Sexual state unknown. Delon & Kiffer (1978) synonymised T. basicola with Ca. elegans, at the time treated in Chalara (Nag Raj & Kendrick 1975). Paulin-Mahady showed the species is best treated in Thielaviopsis, thus reversing the synonymy and bringing the name to its current state. Although our data have shown that T. basicola does not form part of Thielaviopsis as defined in the present study, the species is best treated in this genus until an epitype is designated that is linked to the holotype specimen. On that basis a final generic placement can be ascertained.

Bohár, Acta Phytopathol. Entomol. Hung. 31: 215. 1996.

Notes: In the original description of this species from oak in Hungary and the United Kingdom, Bohár (1996) stated that it is closely related, but distinct from C. virescens (now D. virescens). Apart from a similar host, the elongated, sheathed ascospores of D. erinaceus suggest a placement in Davidsoniella. However, no cultures were available and we prefer to consider its generic placement as uncertain until epitypification can be achieved.

(Bretz) J. Hunt, Lloydia 19: 21. 1956.

Basionym: Endoconidiophora fagacearum Bretz, Phytopathology 42: 437. 1952.

= Chalara quercina Henry, Phytopathology 34: 631. 1944.

≡ Thielaviopsis quercina (Henry) A.E. Paulin, T.C. Harr. & McNew, Mycologia 94: 70. 2002.

Descriptions: Hunt (1956: 21); Nag Raj & Kendrick (1975: 94, 131, fig. 32A); Upadhyay (1981: 66); Potlajczuk & Schekunova (1985: 150); Kolařík & Hulcr (2009).

Notes: The asexual state of this causal agent of oak wilt was described first as Ca. quercina (Henry 1944), while the sexual state was later discovered and described as E. fagacearum (Bretz 1952). The isolate used in our study groups outside the major lineages, usually relatively close to, but still very distinct from, A. ferruginea. Because this isolate does not represent the type of either of these species, we have chosen to treat the species in Ceratocystis until typification can be resolved.

L.R. Batra, Mycologia 59: 980. 1967.

≡ Monilia ferruginea Math.-Käärik, Meddel. Statens Skogs-Forskningsinst. 43: 57. 1953. (nom. illegit., Art. 53.1, non M. ferruginea Pers. 1822).

Descriptions: Mathiesen-Käärik (1953: 53–57, figs 5–7); Batra (1967: 1000–1004, figs 30, 31, 40).

Notes: Sexual state unknown. In our analyses, this species did not group in Ambrosiella s. str., but relatively close to, but still very distinct from, C. fagacearum. Because the isolate used in our study does not represent the type for the species, it is best treated in Ambrosiella until typification can be achieved.

J. Reid & Hausner, Botany 88: 977. 2010.

Notes: A culture for this species could not be obtained. The sequences generated by Reid suggest that this species falls outside the C. coerulescens complex (now Endoconidiophora), in which it would otherwise fit based on morphology and its conifer host. An accurate generic placement will only be possible once a culture can be obtained from which the appropriate sequences can be generated. Until such time it is best treated in Ceratocystis s.l.

Discussion

Ceratocystis s. str. as it is defined in the present study is typified by the well-known species C. fimbriata. The genus currently includes 32 species, all of which were included in the analyses making up this study. The genus includes many important plant pathogens of angiosperm trees, but also of root crops (Kile 1993, Engelbrecht & Harrington 2005, Van Wyk ). These fungi all have ascomata with smooth non-ornamented bases and hat-shaped ascospores; two morphological features that distinguish them from species now in the genera Thielaviopsis (previously C. paradoxa s.l.) and Huntiella (previously C. moniliformis s. l.). Both the latter genera have ornamented ascomatal bases, although the morphology of the ornamentations is different in the two genera.

In some cases, species boundaries for Ceratocystis s. str. are very clear, for example in the cases of the tree pathogens C. platani, C. cacaofunesta, and C. albifundus (Wingfield ). In others, distinction at the species level has been debated (Fourie ). Problems have for example arisen where the ITS region has suggested the existence of species boundaries but where it is now recognised that there are often two or more ITS forms within a single isolate (Al Adawi ). Revisions of these species boundaries are likely to emerge when additional tools, especially those taken from whole genome sequences (Wilken ), become available to discriminate more clearly between species. Another problem, already recognised for this group, is that hybridisation has occurred between species (Engelbrecht & Harrington 2005), a factor that will also confuse the recognition of discrete taxa. What is clear, however, is that there are many species already known in this group and many more will likely be found in the future.

The asexual genus Chalaropsis has been emended to accommodate three species that are found on woody substrates. Two of these three were included in the analyses, along with information from a fourth undescribed species discovered in a culture collection. None of these fungi are known to have any economic or critical important ecological significance.

The genus Endoconidiophora was emended to accommodate an important group of eight species that occur mostly on conifers and many of which are symbionts of conifer-infesting bark beetles. These fungi have previously been referred to as “the Gymnosperm section” in the C. coerulescens s. l. group (Harrington 2009, Wingfield ) and they include a number of important pathogenic species such as E. polonica, E. laricicola, E. laricis and E. rufipennis (Redfern ). Other species are mostly agents of sap stain in conifer timber. Unlike species in Ceratocystis s. s., Huntiella and Thielaviopsis as circumscribed here, these fungi have ascospores that are not hat-shaped but rather are obovoid, with distinct sheaths (Fig. 3). Ceratocystis norvegica, a species from conifers in Norway (Reid ) that seems to fit the description of Endoconidiophora, but for which material was not available, should be considered in future treatments of this genus.

Davidsoniella is described as a new genus to accommodate members of what has previously been referred to as “the Angiosperm section” of C. coerulescens s. l. (Harrington 2009, Wingfield ). The group includes four species, of which two, D. virescens and D. eucalypti, have known sexual states. The fusiform ascospores with evenly distributed hyaline sheaths are similar to but distinct from those of species now accommodated in Endoconidiophora. The remaining two species (D. australis and D. neocaledoniae) are known only by their asexual morphs, with the “chalara- or thielaviopsis-like” morphology typical of all species in Ceratocystis s. l. other than Ambrosiella. Interestingly, three of these fungi (D. virescens, D. neocaledoniae and D. australis) are important tree pathogens (Hepting 1944, Dadant 1950, Kile & Walker 1987) while one (D. eucalypti) is not known to be a pathogen (Kile ). Three species (D. eucalypti, D. neocaledoniae and D. australis) are known exclusively from Australasia, while D. virescens occurs in North America. One more species fits the description of Davidsoniella, namely C. erinaceus from oak in Europe (Bohár 1996). No sequence data exist for this species but it should be considered in future treatments of this genus.

The emended asexual genus Thielaviopsis includes species previously placed in C. paradoxa s. l., some of which have known sexual states. Until recently all species in this group were aggregated in the single species, C. paradoxa, but Mbenoun ’s sequencing and mating studies disclosed six species in what they referred to as the C. paradoxa complex. They showed that the type species of Thielaviopsis, T. ethacetica, though previously treated as anamorph of C. paradoxa (Nag Raj & Kendrick 1975), is a distinct species. In one species, T. euricoi, no sexual state has been observed, but the others all produce hat-shaped ascospores. The outstanding characteristic of this genus, however, is the presence of prominent, digitate appendages on the ascomatal bases (Fig. 2). Most of these fungi occur on monocotyledenous plants including palms, pineapple and banana and some are important plant pathogens (Mitchell 1937, Bliss 1941, Abdullah ).

Huntiella was established in this study to accommodate a well-recognised and large group of species that have previously been referred to as residing in C. moniliformis s. l. Nineteen species are recognised in Huntiella of which 18 were included in the analyses. Two of these species are in the process of being described (Mbenoun unpubl. data). Species of Huntiella have hat-shaped ascospore (Fig. 3) similar to those found in Ceratocystis s. str. but they have very distinct ascomata. The latter feature necks with basal plates that easily disconnect from the ascomatal bases, which are also ornamented with spines (Fig. 2). Huntiella spp. are very commonly encountered on tree wounds and they are typically non-pathogenic (Roux ).

The genus Ambrosiella is perhaps the most unusual in Ceratocystis s. l. The five species accommodated in this genus (three of which were included in the analyses) are all symbionts of wood-boring “ambrosia” beetles and they lack a known sexual state. They are the only species in Ceratocystis s. l. that do not have typical “chalara-like” conidiogenous cells. Instead they have tubular tapering conidiophores and rectangular conidia formed in chains.

Four species in Ceratocystis s. l. did not reside in any of the six major phylogenetic clades arising from this study. These species included C. adiposa, C. fagacearum, Thielaviopsis basicola and Ambrosiella ferruginea. These clearly represent discrete genera, which as collections increase in the future, will most likely accommodate additional species. This would be the same situation that has arisen for other genera now recognised in Ceratocystis s. l. and that previously included very few obvious species. For the present, we have chosen not to provide generic descriptions for these species. We believe that they are likely to be more clearly defined in the future, particularly since three of the four require additional work to obtain living material that can be unambiguously reconciled with their typification. Three of these four fungi (C. fagacearum, C. adiposa and T. basicola) are well-recognised plant pathogens (Butler 1906, Yarwood 1981, Juzwik ) and we recognise that name changes could cause some confusion. It will thus be important to make it clear in studies that these fungi are phylogenetically unrelated to the genera in which they are currently treated.

Phylogenetic analyses based on three carefully selected gene regions in this study have provided robust data to be able to distinguish more effectively between a large number of important and very different fungi that have, for many years, been unfortunately lumped in a single genus. The improved resolution has emerged through intensive collecting initiatives in new areas and through the application of new technologies that have improved our ability to recognise cryptic taxa. As global collecting initiatives expand for fungi residing in the Ceratocystidaceae, the taxa accommodated in the genera established in this study will surely increase and the boundaries of the few remaining monotypic lineages will also be elucidated.

The new and rapidly stabilising nomenclatural code for fungi (McNeill ) underpins a natural classification and a single name for all fungal taxa. This is a major and positive change that will ultimately promote a more effective taxonomy for fungi and it will ensure easier relationships with important associated disciplines such as plant pathology (Hawksworth 2011, Wingfield ). A one fungus one name scheme has already been presented for the so-called ophiostomatoid fungi including the Ceratocystidaceae (De Beer ). In the present study, we have followed this approach rigorously. As far as possible, available generic names, in all cases those associated with asexual morphs have been used. In two instances entirely new generic names have been established and these honour two important early pioneers of the taxonomy of the ophiostomatoid fungi. They are John Hunt who produced the first comprehensive monograph of Ceratocystis (Hunt 1956) and Ross W. Davidson who dedicated his career to collecting, identifying and describing species of ophiostomatoid fungi including several Ceratocystis spp.

Table 2

Species previously treated in Ceratocystis, but now excluded from the genus because they were shown to belong to other genera. More details on each species are presented by De Beer .

Name in Ceratocystis	Current name	Basionym
C. abiocarpa R.W. Davidson	Grosmannia abiocarpa (R.W. Davidson) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis abiocarpa R.W. Davidson
C. adjuncti R.W. Davidson	Ophiostoma adjuncti (R.W. Davidson) Harrington	Ceratocystis adjuncti R.W. Davidson
C. albida (Math.-Käärik) J. Hunt	synonym of Ophiostoma stenoceras (Robak) Nannf.	Ophiostoma albidum Math.-Käärik
C. allantospora H.D. Griffin	Ophiostoma allantosporum (Griffin) M. Villarreal	Ceratocystis allantospora H.D. Griffin
C. ambrosia Bakshi	Ophiostoma ambrosium (Bakshi) Hausner, J. Reid & Klassen	Ceratocystis ambrosia Bakshi
C. angusticollis Wright & H.D. Griffin	Ophiostoma angusticollis (Wright & Griffin) M. Villarreal	Ceratocystis angusticollis Wright & H.D. Griffin
C. araucariae Butin	Ophiostoma araucariae (Butin) de Hoog & Scheffer	Ceratocystis araucariae Butin
C. arborea Olchow. & J. Reid	Ophiostoma arborea (Olchow. & J. Reid) Yamaoka & M.J. Wingf.	Ceratocystis arborea Olchow. & J. Reid
C. aurea (R.C. Rob. & R.W. Davidson) H.P. Upadhyay	Grosmannia aurea (R.C. Rob. & R.W. Davidson) Zipfel, Z.W. de Beer & M.J. Wingf.	Europhium aureum R.C. Rob. & R.W. Davidson
C. bacillospora Butin & G. Zimm.	Ophiostoma bacillosporum (Butin & G. Zimm.) de Hoog & Scheffer	Ceratocystis bacillospora Butin & G. Zimm.
C. bicolor (R.W. Davidson & Wells) R.W. Davidson	Ophiostoma bicolor R.W. Davidson & D.E. Wells	Ophiostoma bicolor R.W. Davidson & D.E. Wells
C. brunnea R.W. Davidson	Ophiostoma brunneum (R.W. Davidson) Hausner & J. Reid	Ceratocystis brunnea R.W. Davidson
C. brunneo-ciliata (Math.-Käärik) J. Hunt	Ophiostoma brunneo-ciliatum Math.-Käärik	Ophiostoma brunneo-ciliatum Math.-Käärik
C. brunneocrinita E.F. Wright & Cain	Graphilbum brunneocrinitum (E.F. Wright & Cain) Z.W. de Beer & M.J. Wingf.	Ceratocystis brunneocrinita E.F. Wright & Cain
C. cainii Olchow. & J. Reid	Grosmannia cainii (Olchow. & J. Reid) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis cainii Olchow. & J. Reid
C. californica DeVay, R.W. Davidson & Moller	Ophiostoma californicum (DeVay, R.W. Davidson & Moller) Hausner, J. Reid & Klassen	Ceratocystis californica DeVay, R.W. Davidson & Moller
C. cana (Münch) Moreau	Ophiostoma canum (Münch) Syd.	Ceratostomella cana Münch
C. capitata H.D. Griffin	synonym of Ophiostoma tenellum (R.W. Davidson) M. Villarreal	Ceratocystis capitata H.D. Griffin
C. castaneae (Vanin & Solovjev) C. Moreau	Ophiostoma castaneae (Vanin & Solovjev) Nannf.	Ceratostomella castaneae Vanin & Solovjev
C. catoniana (Goid.) C. Moreau	Ophiostoma catonianum (Goid.) Goid.	Ceratostomella catoniana Goid.
C. clavata (Math.) Hunt	Ophiostoma clavatum Math.	Ophiostoma clavatum Math.
C. clavigera (R.C. Rob. & R.W. Davidson) H.P. Upadhyay	Grosmannia clavigera (R.C. Rob. & R.W. Davidson) Zipfel, Z.W. de Beer & M.J. Wingf.	Europhium clavigerum R.C. Rob. & R.W. Davidson
C. columnaris Olchow. & J. Reid	Ophiostoma columnare (Olchow. & J. Reid) Seifert & G. Okada	Ceratocystis columnaris Olchow. & J. Reid
C. concentrica Olchow. & J. Reid	Ceratocystiopsis concentrica (Olchow. & J. Reid) H.P. Upadhyay	Ceratocystis concentrica Olchow. & J. Reid
C. conicicollis Olchow. & J. Reid	Ceratocystiopsis conicicollis (Olchow. & J. Reid) H.P. Upadhyay	Ceratocystis conicicollis Olchow. & J. Reid
C. coronata Olchow. & J. Reid	Ophiostoma coronatum (Olchow. & J. Reid) M. Villarreal	Ceratocystis coronata Olchow. & J. Reid
C. crassivaginata H.D. Griffin	Grosmannia crassivaginata (H.D. Griffin) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis crassivaginata H.D. Griffin
C. crenulata Olchow. & J. Reid	Ophiostoma crenulatum (Olchow. & J. Reid) Hausner & J. Reid	Ceratocystis crenulata Olchow. & J. Reid
C. curvicollis Olchow. & J. Reid	Graphilbum curvicolle (Olchow. & J. Reid) Z.W. de Beer & M.J. Wingf.	Ceratocystis curvicollis Olchow. & J. Reid
C. davidsonii Olchow. & J. Reid	Grosmannia davidsonii (Olchow. & J. Reid) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis davidsonii Olchow. & J. Reid
C. denticulata R.W. Davidson	Ophiostoma denticulatum (R.W. Davidson) Z.W. de Beer & M.J. Wingf.	Ceratocystis denticulata R.W. Davidson
C. distorta R.W. Davidson	Ophiostoma distortum (R.W. Davidson) de Hoog & Scheffer	Ceratocystis distorta R.W. Davidson
C. dolominuta H.D. Griffin	synonym of Ceratocystiopsis minuta (Siemaszko) H.P. Upadhyay & W.B. Kendr.	Ceratocystis dolominuta H.D. Griffin
C. dryocoetidis W.B. Kendr. & Molnar	Grosmannia dryocoetidis (W.B. Kendr. & Molnar) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis dryocoetidis W.B. Kendr. & Molnar
C. epigloea Guerrero	Ophiostoma epigloeum (Guerrero) de Hoog	Ceratocystis epigloea Guerrero
C. eucastaneae R.W. Davidson	synonym of Ophiostoma stenoceras (Robak) Nannf.	Ceratocystis eucastaneae R.W. Davidson
C. europhioides E.F. Wright & Cain	Grosmannia europhioides (E.F. Wright & Cain) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis europhioides E.F. Wright & Cain
C. fagi (W. Loos) C. Moreau	synonym of Ophiostoma quercus (Georgev.) Nannf.	Ceratostomella fagi W. Loos
C. falcata E.F. Wright & Cain	Cornuvesica falcata (E.F. Wright & Cain) C.D. Viljoen, M.J. Wingf. & K. Jacobs	Ceratocystis falcata E.F. Wright & Cain
C. fasciata Olchow. & J. Reid	Ophiostoma fasciatum (Olchow. & J. Reid) Hausner, J. Reid & Klassen	Ceratocystis fasciata Olchow. & J. Reid
C. fimicola (Marchal) H.P. Upadhyay	Sphaeronaemella fimicola Marchal	Sphaeronaemella fimicola Marchal
C. floccosa (Math.) J. Hunt	Ophiostoma floccosum Math.	Ophiostoma floccosum Math.
C. francke-grosmanniae R.W. Davidson	Grosmannia francke-grosmanniae (R.W. Davidson) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis francke-grosmanniae R.W. Davidson
C. fraxinopennsylvanica T.E. Hinds	Togninia fraxinopennsylvanica (T.E. Hinds) Hausner, Eyjólfsdóttir & J. Reid	Ceratocystis fraxinopennsylvanica T.E. Hinds
C. galeiformis Bakshi	Grosmannia galeiformis (B.K. Bakshi) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis galeiformis Bakshi
C. gossypina R.W. Davidson	Ophiostoma gossypinum (R.W. Davidson) J. Taylor	Ceratocystis gossypina R.W. Davidson
C. gossypina var. robusta R.W. Davidson	synonym of Ophiostoma stenoceras (Robak) Nannf.	Ceratocystis gossypina var. robusta R.W. Davidson
C. grandifoliae R.W. Davidson	Grosmannia grandifoliae (R.W. Davidson) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis grandifoliae R.W. Davidson
C. helvellae (P. Karst.) H.P. Upadhyay	Sphaeronaemella helvellae (P. Karst.) P. Karst.	Sphaeria helvellae P. Karst.
C. horanszkyi Tóth	Sphaeronaemella horanszkyi (Tóth) Tóth	Ceratocystis horanszkyi Tóth
C. huntii R.C. Rob.	Grosmannia huntii (R.C. Rob.) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis huntii R.C. Rob.
C. hyalothecium R.W. Davidson	Ophiostoma hyalothecium (R.W. Davidson) Hausner, J. Reid & Klassen	Ceratocystis hyalothecium R.W. Davidson
C. introcitrina Olchow. & J. Reid	Ophiostoma introcitrinum (Olchow. & J. Reid) Hausner, J. Reid & Klassen	Ceratocystis introcitrina Olchow. & J. Reid
C. ips (Rumbold) C. Moreau	Ophiostoma ips (Rumbold) Nannf.	Ceratostomella ips Rumbold
C. leptographioides (R.W. Davidson) J. Hunt	Grosmannia leptographioides (R.W. Davidson) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratostomella leptographioides R.W. Davidson
C. leucocarpa R.W. Davidson	Ophiostoma leucocarpum (R.W. Davidson) Z.W. de Beer & M.J. Wingf.	Ceratocystis leucocarpa R.W. Davidson
C. longirostellata Bakshi	Ophiostoma longirostellatum (Bakshi) Arx & E. Müll.	Ceratocystis longirostellata Bakshi
C. longispora Olchow. & J. Reid	Ceratocystiopsis longispora (Olchow. & J. Reid) H.P. Upadhyay	Ceratocystis longispora Olchow. & J. Reid
C. macrospora Aoshima [nom. inval., Art. 29.1, 36.1]	synonym of Grosmannia laricis (K. van der Westh., Yamaoka & M.J. Wingf.) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis macrospora Aoshima [nom. inval., Art. 29.1, 36.1]
C. megalobrunnea R.W. Davidson & Toole	Ophiostoma megalobrunneum (R.W. Davidson & Toole) de Hoog & Scheffer	Ceratocystis megalobrunnea R.W. Davidson & Toole
C. microspora (Arx) R.W. Davidson	Ophiostoma microsporum Arx	Ophiostoma microsporum Arx
C. minima Olchow. & J. Reid	Ceratocystiopsis minima (Olchow. & J. Reid) H.P. Upadhyay	Ceratocystis minima Olchow. & J. Reid
C. minor (Hedgc.) J. Hunt	Ophiostoma minus (Hedgc.) Syd.	Ceratostomella minor Hedgc.
C. minuta (Siemaszko) J. Hunt	Ceratocystiopsis minuta (Siemaszko) H.P. Upadhyay & W.B. Kendr.	Ophiostoma minutum Siemaszko
C. minuta-bicolor R.W. Davidson	Ceratocystiopsis minuta-bicolor (R.W. Davidson) H.P. Upadhyay & W.B. Kendr.	Ceratocystis minuta-bicolor R.W. Davidson
C. montia (Rumbold) J. Hunt	Ophiostoma montium (Rumbold) Arx	Ceratostomella montium Rumbold
C. multiannulata (Hedgc. & R.W. Davidson) J. Hunt	Ophiostoma multiannulatum (Hedgc. & R.W. Davidson) Hendrix	Ceratostomella multiannulata Hedgc. & R.W. Davidson
C. narcissi (Limber) J. Hunt	Ophiostoma narcissi Limber	Ophiostoma narcissi Limber
C. nigra R.W. Davidson	Graphilbum nigrum (R.W. Davidson), Z.W. de Beer & M.J. Wingf.	Ceratocystis nigra R.W. Davidson
C. nigrocarpa R.W. Davidson	Ophiostoma nigricarpum (R.W. Davidson) de Hoog	Ceratocystis nigrocarpa R.W. Davidson
C. nothofagi Butin	Ophiostoma nothofagi (Butin) Rulamort	Ceratocystis nothofagi Butin
C. novae-zelandiae Hutchison & J. Reid	synonym of Ophiostoma pluriannulatum (Hedgc.) Syd.	Ceratocystis novae-zelandiae Hutchison & J. Reid
C. obscura (R.W. Davidson) J. Hunt	Leptographium obscurum (R.W. Davidson) Z.W. de Beer & M.J. Wingf.	Ceratostomella obscura R.W. Davidson
C. ochracea H.D. Griffin	Ceratocystiopsis ochracea (H.D. Griffin) H.P. Upadhyay	Ceratocystis ochracea H.D. Griffin
C. olivacea (Math.) J. Hunt	Grosmannia olivacea (Math.) Zipfel, Z.W. de Beer & M.J. Wingf.	Ophiostoma olivaceum Math.
C. olivaceapini R.W. Davidson	Grosmannia olivaceapini (R.W. Davidson) Z.W. de Beer, Linnakoski & M.J. Wingf.	Ceratocystis olivaceapini R.W. Davidson
C. ossiformis Olchow. & J. Reid	synonym of Ophiostoma columnare (Olchow. & J. Reid) Seifert & G. Okada	Ceratocystis ossiformis Olchow. & J. Reid
C. pallida H.D. Griffin	synonym of Ceratocystiopsis minuta-bicolor (R.W. Davidson) H.P. Upadhyay & W.B. Kendr.	Ceratocystis pallida H.D. Griffin
C. pallidobrunnea Olchow. & J. Reid	Ceratocystiopsis pallidobrunnea (Olchow. & J. Reid) H.P. Upadhyay	Ceratocystis pallidobrunnea Olchow. & J. Reid
C. parva Olchow. & J. Reid	Ceratocystiopsis parva (Olchow. & J. Reid) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis parva Olchow. & J. Reid
C. penicillata (Grosmann) C. Moreau	Grosmannia penicillata (Grosmann) Goid.	Ceratostomella penicillata Grosmann
C. perfecta R.W. Davidson	Ophiostoma perfectum (R.W. Davidson) de Hoog	Ceratocystis perfecta R.W. Davidson
C. perparvispora J. Hunt	synonym of Ophiostoma microsporum Arx	Ceratocystis perparvispora J. Hunt
C. piceae (Münch) Bakshi	Ophiostoma piceae (Münch) Syd.	Ceratostomella piceae Münch
C. piceiperda (Rumbold) C. Moreau	Grosmannia piceiperda (Rumbold) Goid.	Ceratostomella piceiperda Rumbold
C. pilifera (Fr.) C. Moreau	Ophiostoma piliferum (Fr. : Fr.) Syd.	Sphaeria pilifera Fr.
C. pini (Münch) C. Moreau	synonym of Ophiostoma minus (Hedgc.) Syd.	Ceratostomella pini Münch
C. pluriannulata (Hedgc.) C. Moreau	Ophiostoma pluriannulatum (Hedgc.) Syd.	Ceratostomella pluriannulata Hedgc.
C. polygrapha Aoshima [nom. inval., Art. 29.1, 36.1]	synonym of Grosmannia aoshimae (Ohtaka, Masuya & Yamaoka) Masuya & Yamaoka	Ceratocystis polygrapha Aoshima [nom. inval., Art. 29.1, 36.1]
C. ponderosae T.E. Hinds & R.W. Davidson	synonym of Ophiostoma stenoceras (Robak) Nannf.	Ceratocystis ponderosae T.E. Hinds & R.W. Davidson
C. populicola Olchow. & J. Reid	Ophiostoma populicola (Olchow. & J. Reid) Z.W. de Beer, Seifert, M.J. Wingf.	Ceratocystis populicola Olchow. & J. Reid
C. populina T.E. Hinds & R.W. Davidson	Ophiostoma populinum (T.E. Hinds & R.W. Davidson) de Hoog & Scheffer	Ceratocystis populina T.E. Hinds & R.W. Davidson
C. prolifera Kowalski & Butin	Ophiostoma proliferum (Kowalski & Butin) Rulamort	Ceratocystis prolifera Kowalski & Butin
C. pseudoeurophioides Olchow. & J. Reid	Grosmannia pseudoeurophioides (Olchow. & J. Reid) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis pseudoeurophioides Olchow. & J. Reid
C. pseudominor Olchow. & J. Reid	Ophiostoma pseudominus (Olchow. & J. Reid) Hausner, J. Reid & Klassen	Ceratocystis pseudominor Olchow. & J. Reid
C. pseudonigra Olchow. & J. Reid	Ophiostoma pseudonigrum (Olchow. & J. Reid) Hausner & J. Reid	Ceratocystis pseudonigra Olchow. & J. Reid
C. pseudotsugae (Rumbold) C. Moreau	Ophiostoma pseudotsugae (Rumbold) Arx	Ceratostomella pseudotsugae Rumbold
C. querci (Georgev.) C. Moreau	Ophiostoma quercus (Georgev.) Nannf.	Ceratostomella quercus Georgev.
C. retusi R.W. Davidson & T.E. Hinds	Ophiostoma retusum (R.W. Davidson & T.E. Hinds) Hausner, J. Reid & Klassen	Ceratocystis retusi R.W. Davidson & T.E. Hinds
C. roboris (Georgescu & Teodoru) Potl.	synonym of Ophiostoma quercus (Georgev.) Nannf.	Ophiostoma roboris Georgescu & Teodoru
C. robusta (R.C. Rob. & R.W. Davidson) H.P. Upadhyay	Grosmannia robusta (R.C. Rob. & R.W. Davidson) Zipfel, Z.W. de Beer & M.J. Wingf.	Europhium robustum R.C. Rob. & R.W. Davidson
C. rostrocoronata R.W. Davidson & Eslyn	Ophiostoma rostrocoronatum (R.W. Davidson & Eslyn) de Hoog & Scheffer	Ceratocystis rostrocoronata R.W. Davidson & Eslyn
C. rostrocylindrica (R.W. Davidson) J. Hunt	Leptographium rostrocylindricum (R.W. Davidson) Z.W. de Beer & M.J. Wingf.	Ceratostomella rostrocylindrica R.W. Davidson
C. sagmatospora E.F. Wright & Cain	Grosmannia sagmatospora (E.F. Wright & Cain) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis sagmatospora E.F. Wright & Cain
C. serpens (Goid.) C. Moreau	Grosmannia serpens Goid.	Grosmannia serpens Goid.
C. shikotsuensis Aoshima [nom. inval., Art. 29.1, 36.1]	synonym of Grosmannia europhioides (E.F. Wright & Cain) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis shikotsuensis Aoshima [nom. inval., Art. 29.1, 36.1]
C. sparsa R.W. Davidson	Graphilbum sparsum H.P. Upadhyay & W.B. Kendr.	Ceratocystis sparsa R.W. Davidson
C. spinifera Olchow. & J. Reid	synonym of Ophiostoma fasciatum (Olchow. & J. Reid) Hausner, J. Reid & Klassen	Ceratocystis spinifera Olchow. & J. Reid
C. spinulosa H.D. Griffin	Ceratocystiopsis spinulosa (H.D. Griffin) H.P. Upadhyay	Ceratocystis spinulosa H.D. Griffin
C. stenoceras (Robak) C. Moreau	Ophiostoma stenoceras (Robak) Nannf.	Ceratostomella stenoceras Robak
C. tenella R.W. Davidson	Ophiostoma tenellum (R.W. Davidson) M. Villarreal	Ceratocystis tenella R.W. Davidson
C. tetropii (Math.) J. Hunt	Ophiostoma tetropii Math.	Ceratocystis tetropii (Math.) J. Hunt
C. torticiliata Olchow. & J. Reid	Ophiostoma torticiliata (Olchow. & J. Reid) Seifert & G. Okada	Ceratocystis torticiliata Olchow. & J. Reid
C. torulosa Butin & G. Zimm.	Ophiostoma torulosum (Butin & G. Zimm.) Hausner, J. Reid & Klassen	Ceratocystis torulosa Butin & G. Zimm.
C. tremulo-aurea R.W. Davidson & T.E. Hinds	Ophiostoma tremulo-aureum (R.W. Davidson & T.E. Hinds) de Hoog & Scheffer	Ceratocystis tremulo-aurea R.W. Davidson & T.E. Hinds
C. triangulospora (Butin) H.P. Upadhyay	Ophiostoma triangulosporum Butin	Ophiostoma triangulosporum Butin
C. truncicola (R.W. Davidson) H.D. Griffin	Grosmannia truncicola (R.W. Davidson) Z.W. de Beer & M.J. Wingf.	Ophiostoma truncicola R.W. Davidson
C. tubicollis Olchow. & J. Reid	Graphilbum tubicolle (Olchow. & J. Reid) Z.W. de Beer & M.J. Wingf.	Ceratocystis tubicollis Olchow. & J. Reid
C. ulmi (Buisman) C. Moreau	Ophiostoma ulmi (Buisman) Nannf.	Ceratostomella ulmi Buisman
C. valachicum (Georgescu, Teodoru & Badea) Potl.	Ophiostoma valachicum Georgescu, Teodoru & Badea	Ophiostoma valachicum Georgescu, Teodoru & Badea
C. vesca R.W. Davidson	Grosmannia vesca (R.W. Davidson) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis vesca R.W. Davidson
C. wageneri Goheen & F.W. Cobb	Grosmannia wageneri (Goheen & F.W. Cobb) Zipfel, Z.W. de Beer & M.J. Wingf.	Ceratocystis wageneri Goheen & F.W. Cobb

Table 3

Species previously treated in Ceratocystis s.l. but that can be excluded from the current generic concepts based on morphology. However, the correct generic placement of these species remains uncertain and in need of confirmation with DNA sequences. More details on each species are presented by De Beer .

Name in Ceratocystis	Basionym	Probable ordinal, generic placement
C. acericola H.D. Griffin	Ceratocystis acericola H.D. Griffin	Ophiostomatales, Ophiostoma s.l. or Leptographium s.l.
C. acoma (V.V. Miller & Cernzow) C. Moreau	Ceratostomella acoma V.V. Miller & Cernzow	Ophiostomatales, Ophiostoma s. str.
C. aequivaginata Olchow. & J. Reid	Ceratocystis aequivaginata Olchow. & J. Reid	Ophiostomatales, Leptographium s.l.
C. alba DeVay, R.W. Davidson & W.J. Moller	Ceratocystis alba DeVay, R.W. Davidson & W.J. Moller	Peripheral to Ophiostomatales
C. autographa Bakshi	Ceratocystis autographa Bakshi	Sordariomycetidae, incertae sedis
C. brevicollis R.W. Davidson	Ceratocystis brevicollis R.W. Davidson	Ophiostomatales, Ophiostoma s.l. or Leptographium s.l.
C. buxi (Borissov) C. Moreau	Ceratostomella buxi Borissov	Sordariomycetidae, incertae sedis, Ceratostomella
C. comata (V.V. Miller & Cernzow) C. Moreau	Ceratostomella comata V.V. Miller & Cernzow	Ophiostomatales, Leptographium s.l.
C. deltoideospora Olchow. & J. Reid	Ceratocystis deltoideospora Olchow. & J. Reid	Ophiostomatales, Raffaelea
C. grandicarpa Kowalski & Butin	Ceratocystis grandicarpa Kowalski & Butin	Ophiostomatales, genus uncertain
C. imperfecta (V.V. Miller & Cernzow) C. Moreau	Ceratostomella imperfecta V.V. Miller & Cernzow	Ophiostomatales, Leptographium s.l.
C. magnifica H.D. Griffin	Ceratocystis magnifica H.D. Griffin	Ophiostomatales, Ophiostoma s.l.
C. merolinensis (Georgev.) C. Moreau	Ceratostomella merolinensis Georgev.	Sordariomycetidae, incertae sedis, Ceratostomella
C. microcarpa (P. Karst.) C. Moreau	Ceratostomella microcarpa P. Karst.	Sordariomycetidae, incertae sedis, Ceratostomella
C. seticollis R.W. Davidson	Ceratocystis seticollis R.W. Davidson	Ophiostomatales, genus uncertain
C. stenospora H.D. Griffin	Ceratocystis stenospora H.D. Griffin	Ophiostomatales, Ophiostoma s.l. or Leptographium s.l.
C. trinacriformis (A.K. Parker) H.P. Upadhyay	Europhium trinacriforme A.K. Parker	Ophiostomatales, Ophiostoma s.l. or Leptographium s.l.
C. valdiviana Butin	Ceratocystis valdiviana Butin	Ophiostomatales, Leptographium s.l.