Taxonomic revision of Blumeria based on multi-gene DNA sequences, host preferences and morphology

Abstract

A taxonomic revision of the hitherto monotypic genus Blumeria was conducted incorporating multi-gene sequence analyses, host preference data and morphological criteria. The sequenced loci included rDNA ITS, partial chitin synthase gene (CHS1), as well as fragments of two unnamed orthologous genes (Bgt-1929, Bgt-4572). The combined evidence led to a reassessment and a new neotypification of B. graminiss. str. (emend.), and the description of seven additional species, viz. B. americana sp. nov. (mainly on hosts of the Triticeae), B. avenae sp. nov. (on Avena spp.), B. bromi-cathartici sp. nov. (on Bromus catharticus), B. bulbigera comb. nov. (on Bromus spp.), B. dactylidis sp. nov. (on Dactylis glomerata as the main host, but also on various other hosts), B. graminicola sp. nov. (on Poa spp. as principal hosts, but also on various other hosts), and B. hordei sp. nov. (on Hordeum spp.). Synonyms were assessed, some were lectotypified, and questionable names previously associated with powdery mildew on monocots were discussed although their identities remained unresolved. Keys to the described species were developed.

1. Introduction

Powdery mildew fungi are responsible for a variety of common and important diseases of cereals and grasses (Poaceae), many have world-wide distributions and some cause significant yield losses and quality reduction in the cereal production (Everts, Leath, & Finney, 2001). The causal pathogen of the powdery mildew disease on wheat (Triticum aestivum L.) was first described as Erysiphe graminis DC. (Candolle, 1815). Thereafter, the species has been redescribed many times under various names legitimately or illegitimately, as reviewed by Braun and Cook (2012). According to Speer (1975 [1973–1974]), Golovin erected Blumeria to separate wheat powdery mildew from other Erysiphe spp. based on Blumer’s observations (Blumer, 1933; Golovin, 1958). However, the erection of a new genus failed to comply with the rules of nomenclature in force at the time due to the absence of a Latin description, therefore the generic name, Blumeria Golovin, 1958, was considered invalid. Speer redescribed the genus Blumeria properly, made the combination B. graminis (DC.) Speer and identified seven host species in addition to T. aestivum (Speer, 1975 [1973–1974]). Intra-specific variation was noted very early by Saccardo who distinguished the form E. graminis f. dactylis-glomeratae (Exs: Sacc. Mycoth. Ven. 606, 1876) from others.Jaczewski (1927) introduced 26 formae (host/substrate forms), corresponding to each host genus. Marchal (1902) introduced the first formae speciales (f. spp.) for E. graminis based on inoculation experiments. Several subsequent authors added additional formae and formae speciales introduced the first formae speciales (f. spp.) for E. graminis based on inoculation experiments. Several subsequent authors added additional formae and formae speciales (Marchal, 1902; Jaczewski, 1927; Mains, 1933; Cherewick, 1944; Bunkina, 1967, 1973, 1974). In the latest classification of powdery mildew, B. graminis is the only species in this genus, which is classified in the monotypic tribe Blumerieae within Erysiphaceae (Braun & Cook, 2012). The host species have been recorded in 107 genera of Poaceae (Braun & Cook, 2012, also see Taxonomy section). Some are common hosts with a wide distribution, such as Avena, Bromus, Dactylis, Elymus, Hordeum, Poa, Secale, and Triticum. Some are rarely recorded, suggesting occasional host jumps by the pathogen.

Multi-locus sequences analyses (MLSA) conducted by Inuma, Khodaparast and Takamatsu (2007) recovered nine lineages correlated with host specialization, and demonstrated reproductive isolation between the lineages. Among these lineages, some had a restricted host range of a single genus or species, i.e. lineages on Avena, Bromus, Diarrhena and Hordeumf, while others infected host species in 2 or 3 genera, i.e. lineages on Dactylis, Poa andTriticum. Bromus hosted three lineages. Validation of these phylogenetic species (lineages) within the B. graminis complex required comprehensive morphological analyses. The purpose of this study was to describe and distinguish eight of them by a combination of molecular characters, morphology and putative host preference, and provide formal taxonomic names. The ninth lineage on Diarrhena (Inuma et al., 2007) was not included because only one specimen was available.

2. Materials and methods

2.1. Fungal specimens and gDNA extraction

For DNA MLSA and morphological examination, 163 specimens of B. graminis s. lat. on cereal crops and various grasses were borrowed from five international herbaria: Canadian National Mycological Herbarium (DAOM), U.S. National Fungus Collections (BPI), Martin-Luther-Universität, Institut für Biologie, Bereich Geobotanik und Botanischer Garten Herbarium (HAL), the National Museum of Nature and Science (Tokyo, TNS), and Conservatoire et Jardin botaniques de la Ville de Genève (G). Previously developed DNA sequences, i.e. 49 rDNA-ITS, and 25 CHS1, were downloaded from GenBank as references. An additional 126 specimens from Senckenberg Museum für Naturkunde Görlitz (GLM) were examined for morphology (by UB).

For genomic DNA extraction, three or four 2 mm discs of infected leaf tissues were excised from specimens using Disposable Biopsy Punches Integra™ Miltex® (VWR, Mississauga Ontario, Canada). Alternatively, similar amounts of mycelia and/or chasmothecia were removed from leaf surfaces using tweezers. Macherey-Nagel Nucleomag® 96 Trace kit (Macherey-Nagel, GmbH & Co. KG, Düren, Germany) on a KingFisher Flex magnetic particle processor (Thermo Fisher Scientific Oy, Vantaa, Finland), or an E.Z.N.A. Forensic DNA Kit (Omega Biotek, Inc., Norcross, Georgia, United States) were used for DNA extraction according to the manufacture’s protocols with minor modifications. The modifications were as follows: prior to extraction, samples were frozen using liquid nitrogen and ground using sterile disposable micro-centrifuge tube pestles (PES-15-B-SI Axygen, Corning, New York, USA), and DNA was suspended in 70 µL of elution buffer. Extracted DNA aliquots were stored at -20 °C and the stock DNA was stored at -80 °C.

2.2. PCR, sequencing and analyses

The rDNA-18S (~100 bps)-ITS-28S (~50 bps, abbreviated as ITS region in the following text) region was amplified with two forward and two reverse primers in various combinations, P7 (Mori, Sato, & Takamatsu, 2000), PMITS1, PMITS2 (Cunnington, Takamatsu, Lawrie, & Pascoe, 2003) and ITS4 (White, Bruns, Lee, & Taylor, 1990), and a portion of the chitin synthase gene (CHS1) with primers CHS1-E1f (Seko, Heluta, Grigaliunaite, & Takamatsu, 2011) and CHS1-B3r or CHS1-2r (Inuma et al., 2007). Fragments of two unnamed orthologous genes were amplified using primers designed using the published alignment of 93 phylogenetic informative genes from 31 whole genome sequences of B. graminis (Menardo, Wicker, & Keller, 2017). Geneious R10 primer design module (Biomatters, Aukland, New Zealand) was used to search for candidate oligos with all parameters set as default. The final primer sequences were: locus 1 (Bgt-1929 hypothetic protein exon 1–2 on chromosome 4): FM_27522F 5’-TGTGACGATGGAGATTGTGA-3’ and FM_27868R 5’-CCCATTCGCTGATTGCATAA-3’; and locus 2 (Bgt-4572 exon 3 on chromosome 9, 81% match with E3 ubiquitin-protein ligase inVenustampulla echinocandica Unter., Réblová & Bills): FM_110716F 5’-ATGGAAGGAGTTGATGCAGA-3’ and FM_110946R 5’-GAACTGCTCATCAATTCGCT-3’ (Etymology of primers: FM stands for the initial of Fabrizio Menardo, first author of the 31 B. graminis genome sequences; numbers = location on the concatenated alignment of 93 orthologous loci; F = forward, R = reverse).

Polymerase chain reaction (PCR) was performed in 10 μL reactions containing 1 μL of gDNA, 1× Titanium Taq buffer (with 3.5 mM MgCl₂), 0.1 mM dNTPs, 0.08 µM each of forward and reverse primer, 0.5× Titanium Taq DNA Polymerase (BD Biosciences, San Jose, California, USA), and 0.01 mg bovine serum albumin (BSA) on a TProfessional thermocycler (Biometra, Göttingen, Germany). Touchdown thermocycling protocols were used for rDNA-ITS and two anonymous loci: initial denaturation at 95 °C for 3 min, followed by 10 cycles of 95 °C for 30 s, annealing at 63 °C (decrease 0.5 ºC per cycle) for 45 s, and extension at 72 °C for 2 min, then 20 cycles of 95 °C for 30 s, annealing at 58 °C for 30 s, and extension at 72 °C for 2 min, with a final extension at 72 °C for 10 min. For CHS1, an initial denaturation at 95 °C for 3 min, followed by 6 cycles of 95 °C for 1 min, annealing at 58 °C (decrease 0.5 ºC per cycle) for 45 s, and extension at 72 °C for 1 min 30 s, then 33 cycles of 95 °C for 30 s, annealing at 55 °C for 30 s, and extension at 72 °C for 2 min, with a final extension at 72 °C for 10 min. The PCR products were visualized by using Ethidium Bromide on 1% agarose gels in 1× TBE buffer.

PCR products were amplified with the same primers for Sanger sequencing using ABI BigDye™ Terminator v3.1 Cycling Sequencing Kits with BigDye® Seq Mix diluted 1:8. In 10 µL reaction, volumes of each reagent were 1.75 µL of 5× Sequencing buffer, 2.5 µL of 20% trehalose, 0.5 µL of BigDye® Seq Mix, 0.5 µL of 3.2 µM primer, 3.75 µL sterile HPLC water and 1 µL of PCR product without purification. Thermocycler profiles for the sequencing reactions had an initial denaturation at 95 °C for 3 min, followed by 40 cycles at 95 °C for 30 s, annealing at 55 °C for 15 s and extension at 60 °C for 2 min. An Applied Biosciences Prism® 3130xl Genetic Analyzer (Life Technologies™, California, USA) was used to generate DNA sequences from the sequencing amplification reactions. Sequences were edited using Sequencher 5.4.6 (Gene Codes Corporation, Michigan, USA) or Geneious 10.2.3 (https://www.geneious.com, Biomatters, Aukland, New Zealand).

DNA sequences were aligned with MAFFT online version (Katoh, Rozewicki, & Yamada, 2017). Parsimony and Bayesian inference analyses were conducted on the matrices of each individual locus and a concatenated alignment of four loci for a subset of samples. The most parsimonious trees were searched for using heuristic branch-swapping algorithm, tree-bisection-reconnection (TBR), 200 replicates, number of rearrangements per replicate limit 25000 in PAUP* 4.0b10 (Swofford, 2002), 2000 bootstrap replicates. The best-fit models selected by Modeltest 3.7 (Nylander, 2004) were TrNef+G for ITS,CHS1 and Bgt-1929, and Trn+Gfor Bgt-4572. For Bayesian inference, using MrBayes V3.2.6 (Ronquist et al., 2012), models were set as Nst = 6, rates = invgamma; mcmc were 100 000 000 generations, sampling per 2000 generations; 2 parallel runs each with 4 chains were simultaneously implemented. The runs were terminated when the standard deviation of the average split frequency was lower than 0.03.

Outgroups were selected for analyses of each gene by BLAST searching for available sequences for the closest relatives in NCBI GenBank. These were Podosphaera macularis (Wallr.) U. Braun & S. Takam.MH687414 for ITS, V. echinocandica chitin synthase 1 (XM 032013628.1 range 2317–2600) for CHS1, Rhynchosporium graminicola Heinsen(= R. commune Zaffarano, B.A. McDonald & A. Linde) hypothetic protein KJ410022.1 range 2906–3301 for Bgt-1929; V. echinocandica E3 ubiquitin-protein ligase XM 032013958.1 range 1811–2040 for locus Bgt-4572.

2.3. Microscopy

The infection signs and symptoms were recorded by examining all material in the specimen packets by naked eye, and by a Leica M165C stereo microscope (Leica Microsystems (Canada) Inc., Ontario, Canada). Colors were recorded using standardized color codes according to Kornerup and Wanscher (1978). Photographs were taken with a Leica DFC425 camera and processed using Leica Application Suite software (LAS v4.12.0, Leica Microsystems (Switzerland), Ltd., Heerbrugg, Switzerland). For examining microscopic features of primary and secondary hyphae, hyphal appressoria, conidia, conidiophores, chasmothecia, and asci, the infected leaves were rehydrated using the lactic acid method described by Shin and La (1993) with a minor modification. A Microscope Slide Warmer (VWR, Mississauga Ontario, Canada) was used instead of the alcohol burner for more gentle heating to avoid extreme morphological changes of organ shapes and sizes. The rehydrated mycelia and fruiting structures were scraped from the leaf surface using forceps and mounted in a drop of lactic acid, with or without cotton blue, on microscope slides. Observations were made using a Zeiss Imager M2 (Carl Zeiss Canada Limited, Ontario, Canada). Microphotographs were taken with an Axiocam 503 Color camera and analysed by ZEN (blue edition) 2.6 pro (Carl Zeiss Microscopy GmbH, Jena, Germany); or a Zeiss Axio Scope.A1 (Germany) and Axiocam ERc 5s. At least thirty measurements were made for the sizes whenever possible.

For scanning electron microscopy (SEM), infected leaf material from dried herbarium specimens was rehydrated in a moist chamber (a petri-dish with layers of moist filter paper inserted with the lid on) for 1–2 h. Small pieces of leaf tissue with spores were mounted on carbon double sticky tape on aluminum stubs and coated with an 8 nm thick layer of gold in an Emitech K550V sputter coater (EM Technologies Ltd., Ashford, Kent, England). The samples were imaged on a Quanta 600 SEM operating at 20 kV (FEI Company TM, Brno, Czech Republic).

3. Results

3.1. Sequencing and molecular phylogeny

For the total of 163 samples, varied numbers of sequences were obtained for each locus attempted: 48 rDNA-ITS, 61 CHS1, 103 Bgt-1929 and 144 Bgt-4572. Adding reference sequences downloaded from GenBank (49 ITS and 25 CHS1) resulted in matrices of 97 taxa with 778 characters for ITS, and 87 taxa with 320 characters for CHS1. For the two hypothetical protein loci, only one reference sequence was downloaded from GenBank as outgroup (also see a paragraph in 2.2), resulting in matrices of 104 taxa with 391 characters for Bgt-1929, and 145 taxa with 226 characters for Bgt-4572. Comparisons of the four loci showed that ITS had the poorest performance in amplification and lowest number of informative characters (86/778 = 11%). CHS1 had the highest percentage of informative characters, i.e. 100/320 = 31% (Supplementary Figs. S1,2,3,S4 legends), however, the number of lineages amplified by CHS1 was not as high as two hypothetical protein loci (Supplementary Figs. S1,2,3,S4 ). For instance, none of the B. americana samples was amplified, therefore B. americana was not present in the CHS1 tree (Supplementary Fig. S2). A large number of B. graminicola were only amplified by Bgt-1929 and Bgt-4572. However, B. bromi-cathartici and B. bulbigera were not represented on Bgt-1929 and Bgt-4572 trees because only a few samples for each species were available and PCR amplification was not successful. All trees agreed on the separation of the lineages that were included, in general. A specimen on Anthoxanthum nitens (Weber) Y. Schouten & Veldkamp (= Hierochloe odorata (L.) P. Beauv., synonym recorded on specimen packets) from Saskatchewan (DAOM 4071) grouped in B. graminiss. str. on the ITS tree, however in B. graminicola on all other trees. The identification was determined based on majority rule, but the possibility of a mixed infection or DNA processing error cannot be ruled out. The identities of several orphan lineages cannot be determined, and these are labelled as Blumeria sp. (Table 1; Fig. 1; Supplementary Figs. S1, S2, S4). The species relationships were not strongly supported in any of the trees based on individual genes, indicating limited phylogenetic signals were present for the deeper relationships, therefore a holistic approach was applied as follows.

Fig. 1 - Phylogeny of Blumeria graminis s.lat. inferred from concatenated sequences of rDNA-ITS, CHS1, and two hypothetic protein loci (Bgt-1929, Bgt-4572). Taxon labels include voucher numbers, host name, and country codes (ARG Argentina, CAN Canada, CHE Switzerland, CZE Czech Republic, UK England, FIN Finland, DEU Germany, IRN Iran, ISR Israel, JPN Japan, MEX Mexico, RUS Russia, UKR Ukraine, USA United States); superscript EET stands for ex-epitype, EHT ex-holotype, ENT ex-neotype; values on branches are parsimony bootstrapping value (bp)/Bayesian posterior probability (pp), ~ indicates either bp <70 or pp <0.9. Sequences of taxa in grey font were downloaded from GenBank. Tribes (and subtribes for B. avenae and B. dactylidis) of principal hosts were noted under the species names in parentheses.

Table 1 Fungal specimens for multi-locus sequence analyses

ID	Vouchera	Hostb	Country, province	Year	ITS	CHS1	Bgt1929	Bgt4572
Blumeria americana
	HAL 000028F	Apera spica-venti	DEU, Sachsen-Anhalt	1981	n.a.	n.a.	MT633883	n.a.
	DAOM 96986	Elymus canadensis	USA, Wisconsin	1963	MT622285	n.a.	MT633813	MT650031
	BPI 562942	Elymus canadensis	USA, Wisconsin	1963	MT622266	n.a.	MT633895	MT649952
	BPI 562946	Elymus canadensis	USA, Washington	1934	n.a.	n.a.	n.a.	MT649953
	BPI 563259	Elymus elymoides (Sitanion hystrix)	USA, Arizona	1945	n.a.	n.a.	MT633897	MT650005
	BPI 563260	Elymus elymoides (Sitanion hystrix)	USA, California	1939	n.a.	n.a.	MT633850	MT650006
	BPI 562963	Elymus glaucus	USA, Washington	1938	MT622267	n.a.	MT633860	MT649955
	DAOM 155345	Elymus lanceolatus (Agropyron dasystachyum)	CAN, Northwest Territories	1940	n.a.	n.a.	n.a.	MT650044
	DAOM 137541	Elymus lanceolatus (Agropyron dasystachyum)	USA, Wyoming	1961	MT622287	n.a.	MT633811	MT650033
	BPI 562806	Elymus lanceolatus (Agropyron dasystachyum)	USA, Wyoming	1961	n.a.	n.a.	MT633896	MT649926
	BPI 562805	Elymus lanceolatus (Agropyron dasystachyum)	USA, Oregon	1935	n.a.	n.a.	MT633862	MT649925
	DAOM 186037HT	Elymus repens (Agropyron repens)	CAN, Alberta	1980	MT622296	n.a.	MT633817	MT650055
	BPI 562811	Elymus violaceus (Agropyron latiglume)	USA, Alaska	1948	n.a.	n.a.	n.a.	MT649927
	BPI 562998	Hordeum brachyantherum	USA, Alaska	1948	n.a.	n.a.	n.a.	MT649965
	DAOM 217858	Hordeum jubatum	CAN, Saskatchewan	1931	n.a.	n.a.	MT633831	MT650057
	DAOM 156886	Hordeum jugatum	CAN, Manitoba	1935	n.a.	n.a.	n.a.	MT650046
	DAOM 155342	Hordeum jugatum	CAN, Northwest Territories	1940	MT622292	n.a.	MT633821	n.a.
	DAOM 145059	Leymus cinereus (Elymus cinereus)	CAN, British Columbia	1953	n.a.	n.a.	n.a.	MT650034
	BPI 562828	Pascopyrum smithii (Agropyron smithii)	USA, North Dakota	1942	n.a.	n.a.	n.a.	MT649930
	DAOM 217856	Psathyrostachys juncea (Elymus junceus)	CAN, Saskatchewan	n.a.	MT622297	n.a.	n.a.	n.a.
	BPI 562967	Psathyrostachys juncea (Elymus junceus)	USA, North Dakota	1942	MT622268	n.a.	MT633905	MT649957
B. avenae
	DAOM 147852	Avena barbata	CAN, Ontario	1965	MT622288	n.a.	MT633886	MT650036
	BPI 562860	Avena fatua	USA, California	1942	n.a.	n.a.	n.a.	MT649935
	DAOM 236220HT	Avena sativa	GBR, Exeter	1948	MT622301	n.a.	MT633815	MT650063
	BPI 562866	Avena sativa	GBR, Exeter	1948	n.a.	n.a.	n.a.	MT649936
	BPI 562873	Avena sterilis	ISR, Jerusalem	1951	n.a.	MT633953	MT633846	MT649937
B. bulbigera
	DAOM 82541ET	Bromus hordeaceus subsp. hordeaceus (Bromus mollis)	FIN, Regio aboënsis	1957	MT622280	n.a.	n.a.	n.a.
B. dactylidis
	DAOM 91654	Anthoxanthum odoratum	FIN, Regio aboënsis	1961	MT622284	MT633920	MT633814	MT650030
	HAL 000018F	Anthoxanthum odoratum	DEU, Sachsen-Anhalt	1978	n.a.	n.a.	MT633885	n.a.
	BPI 562894	Bromus catharticus	USA, Georgia	1938	n.a.	n.a.	n.a.	MT649939
	HAL 000027F	Bromus ramosus subsp. benekenii (Bromus benekenii)	DEU, Sachsen-Anhalt	1977	MT622307	n.a.	MT633805	n.a.
	DAOM 118220HT	Dactylis glomerata	CAN, British Columbia	1934	MT622286	MT633919	MT633812	MT650032
	HAL 000029F	Dactylis glomerata	DEU, Sachsen-Anhalt	1978	MT622308	n.a.	MT633903	n.a.
	BPI 562935	Dactylis glomerata	DEU, Oberbayern	1950	n.a.	MT633935	MT633861	MT649950
	BPI 562975	Festuca pratensis	CZE, Kromeriz	1962	n.a.	MT633946	MT633859	MT649959
	HAL 000025F	Festuca gigantea	DEU, Sachsen-Anhalt	1977	n.a.	MT633944	MT633806	MT650066
	BPI 563038	Hordeum vulgare	ETH, Jimma	1954	n.a.	n.a.	n.a.	MT649973
	BPI 563046	Hordeum vulgare	JPN, Kyoto	1895	n.a.	n.a.	n.a.	MT649975
	HAL 000006F	Phleum phleoides	RUS, Baskortostan	1977	n.a.	n.a.	MT633809	n.a.
	BPI 563065	Phleum pratense	LVA, Vidzeme	1935	n.a.	n.a.	n.a.	MT649979
B. graminicola
	DAOM 150568	Agrostis sp.	CAN, Manitoba	1925	MT622290	MT633915	n.a.	MT650040
	HAL 000022F	Alopecurus geniculatus	DEU, Sachsen-Anhalt	1980	n.a.	MT633967	MT633884	n.a.
	DAOM 4071	Anthoxanthus nitens (Hierochloe odorata)	CAN, Saskatchewan	1936	MT622275	MT633955	MT633826	MT650016
	HAL 000016F	Apera spica-venti	DEU, Sachsen-Anhalt	1975	MT622305	MT633961	MT633882	n.a.
	BPI 562847	Apera spica-venti (Agrostis spica-venti)	LVA, Vidzeme	1931	n.a.	n.a.	MT633843	MT649933
	DAOM 152932	Beckmannia eruciformis	CAN, Saskatchewan	1926	MT622291	n.a.	n.a.	MT650043
	DAOM 217878	Beckmannia syzigachne	CAN, Saskatchewan	1959	MT622298	MT633911	n.a.	MT650059
	BPI 562902	Bromus japonicus	CHN, Nanking	1931	n.a.	n.a.	n.a.	MT649943
	BPI 562917	Bromus catharticus var. elatus (Bromus unioloides)	USA, Texas	1932	n.a.	n.a.	n.a.	MT649948
	BPI 562911	Bromus diandrus var. rigidus (Bromus rigidus)	USA, Washington	1935	n.a.	MT633962	MT633844	MT649945
	BPI 562916	Bromus tectorum	USA, Washington	1935	n.a.	MT633936	MT633842	MT649947
	BPI 562924	Dactylis glomerata	ROU, Lapusna-Cornesti	1934	n.a.	n.a.	n.a.	MT649949
	DAOM 155348	Elymus lanceolatus (Agropyron dasystachyum)	CAN, Northwest Territories	1940	n.a.	n.a.	n.a.	MT650045
	BPI 562974	Festuca gigantea	DEU, Mitterfranken	1946	n.a.	MT633934	MT633899	MT649958
	BPI 562965	Festuca idahoensis	USA, Washington	1935	n.a.	MT633960	MT633901	MT649956
	BPI 562984	Holcus lanatus	DEU, Mittelfranken	1946	n.a.	n.a.	n.a.	MT649961
	DAOM 165199	Melica sp.	USA, California	1975	n.a.	n.a.	MT633872	MT650051
	DAOM 55075	Melica subulata	CAN, British Columbia	1956	n.a.	MT633939	MT633835	MT650021
	DAOM 179493	Milium effusum	FIN, Perä-Pohjanmaa	1979	MT622295	MT633950	MT633818	MT650053
	BPI 563056	Milium effusum	DEU, Mittelfranken	1947	n.a.	n.a.	n.a.	MT649978
	BPI 562826	n.a. (Agropyron sericeum)	USA, Alaska	1948	n.a.	n.a.	n.a.	MT649929
	DAOM 89817	Poa annua	CAN, Quebec	1959	n.a.	n.a.	n.a.	MT650026
	DAOM 231497	Poa arctica	CAN, Northwest Territories	1967	MT622300	MT633909	MT633830	MT650062
	DAOM 91194	Poa arctica	CAN, Nunavut	1962	MT622283	MT633921	MT633823	MT650029
	BPI 563089	Poa arida	USA, North Dakota	1941	n.a.	n.a.	n.a.	MT649981
	BPI 563094	Poa bulbosa	RUS, Turkmenskaya	1978	n.a.	MT633932	MT633898	MT649982
	DAOM 217863	Poa compressa	CAN, Saskatchewan	1927	n.a.	n.a.	MT633870	MT650058
	DAOM 152605	Poa compressa	CAN, Manitoba	1933	n.a.	MT633965	MT633874	MT650042
	BPI 563101	Poa compressa	USA, Kansas	1953	n.a.	MT633940	n.a.	MT649985
	DAOM 207718	Poa crocata	CAN, Manitoba	1926	n.a.	n.a.	MT633871	MT650056
	DAOM 38753	Poa glauca	CAN, Quebec	1948	n.a.	n.a.	n.a.	MT650019
	DAOM 148500	Poa glauca	CAN, Ontario	1973	n.a.	n.a.	MT633876	MT650039
	BPI 563103	Poa glaucifolia	USA, North Dakota	1941	n.a.	MT633931	MT633892	MT649986
	BPI 563108	Poa nemoralis	DEU, Oberpfalz	1946	MT622270	MT633945	MT633841	MT649987
	DAOM 148498	Poa nemoralis var. montana	CAN, Ontario	1973	n.a.	MT633917	MT633834	MT650038
	DAOM 181460	Poa palustris	CAN, Manitoba	1979	n.a.	MT633912	MT633832	MT650054
	DAOM 89819	Poa palustris	CAN, Quebec	1959	n.a.	n.a.	MT633877	n.a.
	DAOM 149529	Poa palustris	CAN, Ontario	1971	n.a.	MT633916	n.a.	n.a.
	BPI 563115	Poa palustris	USA, Wisconsin	1948	n.a.	n.a.	n.a.	MT649988
	DAOM 145060	Poa pratensis	CAN, Northwest Territories	1955	n.a.	n.a.	n.a.	MT650035
	DAOM 159510HT	Poa pratensis	CAN, Ontario	1974	n.a.	MT633954	MT633820	MT650048
	DAOM 90074	Poa pratensis	USA, Wisconsin	1962	n.a.	MT633966	MT633887	MT650027
	BPI 563132	Poa pratensis	USA, Alaska	1951	MT622271	MT633948	MT633891	MT649989
	BPI 563135	Poa pratensis	USA, Virginia	1940	n.a.	n.a.	n.a.	MT649990
	BPI 563138	Poa pratensis	USA, Wisconsin	1962	MT622272	MT633949	MT633890	MT649991
	BPI 563148	Poa pratensis	USA, Alaska	1951	n.a.	MT633947	MT633865	MT649992
	BPI 563158	Poa pratensis	USA, Oregon	1937	n.a.	MT633930	MT633900	MT649993
	BPI 563162	Poa pratensis	USA, California	1946	n.a.	MT633964	MT633889	MT649994
	DAOM 159686	Poa pratensis subsp. alpigena	CAN, Northwest Territories	1974	n.a.	MT633938	MT633810	MT650049
	BPI 563191	Poa secunda	USA, Washington	1935	n.a.	n.a.	n.a.	MT649996
	BPI 563096	Poa secunda (Poa canbyi)	USA, North Dakota	1941	n.a.	n.a.	n.a.	MT649983
	BPI 563098	Poa secunda (Poa canbyi)	USA, Washington	1935	n.a.	n.a.	n.a.	MT649984
	BPI 563183	Poa secunda (Poa scabrella)	USA, Oregon	1935	n.a.	n.a.	n.a.	MT649995
	BPI 563076	Poa sp.	IND, Srinagar	1960	n.a.	n.a.	n.a.	MT649980
	HAL 000019F	Poa trivialis	DEU, Sachsen-Anhalt	1977	MT622306	n.a.	MT633867	n.a.
	BPI 563200	Poa trivialis	NLD, Warmond	1958	MT622273	MT633929	MT633840	MT649997
	BPI 563205	Poa vaseyochloa	USA, Oregon	1938	n.a.	n.a.	n.a.	MT649998
	BPI 563212	Polypogon monspeliensis	USA, California	1945	n.a.	MT633928	MT633839	MT649999
	BPI 563216	Polypogon monspeliensis	USA, Washington	1935	n.a.	MT633959	MT633838	MT650000
	DAOM 91193	Puccinellia angusta	CAN, Nunavut	1962	MT622282	n.a.	MT633824	MT650028
	BPI 563219	Puccinellia angusta	CAN, Northwest Territories	1962	n.a.	n.a.	MT633888	MT650001
	BPI 563220	Puccinellia borealis	USA, Alaska	1948	n.a.	n.a.	n.a.	MT650002
	BPI 562833	Thinopyrum intermedium (Agropyron trichophorum)	USA, North Dakota	1942	n.a.	n.a.	n.a.	MT649932
	BPI 563289	Triticum aestivum	AUS, New South Wales	1964	n.a.	n.a.	n.a.	MT650011
B. graminis s.str.
	HAL 000007F	Brachypodium sylvaticum	RUS, Bashkortostan	1977	MT622302	n.a.	MT633868	n.a.
	BPI 562937	Dasypyrum villosum	ROU, Dolj	1979	n.a.	MT633957	MT633902	MT649951
	DAOM 984745	Elymus cf trachycaulus	CAN, New Brunswick	2016	MT622310	MT633937	MT633829	n.a.
	BPI 562803	Elymus ciliaris (Agropyron ciliare)	CHN, Nanking	1931	n.a.	n.a.	MT633863	MT649924
	DAOM 148234	Elymus hystrix (Hystrix patula)	CAN, Quebec	1955	MT622289	MT633918	MT633822	MT650037
	DAOM 161299	Elymus hystrix (Hystrix patula)	CAN, Ontario	1977	MT622293	MT633914	MT633819	MT650050
	HAL 000009F	Elymus repens	UKR, Zaporizhia	1984	MT622303	n.a.	MT633808	MT650064
	DAOM 156887	Elymus repens (Agropyron repens)	CAN, Manitoba	1934	n.a.	n.a.	MT633873	MT650047
	DAOM 152080	Elymus repens (Agropyron repens)	CAN, Manitoba	1945	n.a.	n.a.	MT633875	MT650041
	DAOM 217883	Elymus repens (Agropyron repens)	CAN, Saskatchewan	1939	n.a.	n.a.	MT633869	MT650060
	DAOM 40453	Elymus repens (Agropyron repens)	CAN, Prince Edward Island	1953	MT622278	MT633924	MT633802	MT650020
	DAOM 225780	Elymus repens (Agropyron repens)	CAN, British Columbia	1998	MT622299	MT633910	MT633816	MT650061
	DAOM 27352	Elymus repens (Agropyron repens)	CAN, Quebec	1951	MT622277	MT633968	MT633880	n.a.
	DAOM 82544	Elymus repens (Agropyron repens)	FIN, Ostrobotnia	1951	n.a.	n.a.	MT633878	MT650025
	DAOM 82543	Elymus repens (Agropyron repens)	FIN, Lapp. Enontekiensis	1960	n.a.	MT633956	MT633879	MT650024
	BPI 562819	Elymus repens (Agropyron repens)	ROU, Muntenia	1933	n.a.	n.a.	n.a.	MT649928
	DAOM 74558	Elymus repens (Agropyron repens)	USA, Wisconsin	1960	MT622279	MT633923	n.a.	MT650022
	BPI 562802	Elymus_caninus (Agropyron caninum)	DEU, Mittelfranken	1947	n.a.	n.a.	n.a.	MT649923
	BPI 562989	Hordeum sp.	DEU, Bergisches Land	1931	n.a.	n.a.	n.a.	MT649963
	BPI 563055	Milium effusum	DEU, Thueringen	1945	MT622269	MT633933	MT633853	MT649977
	DAOM 169330	Phleum pratense	CAN, New Brunswick	1978	MT622294	MT633913	MT633833	MT650052
	DAOM 38670	Secale cereale	CAN, Nova Scotia	1952	n.a.	MT633952	MT633836	n.a.
	BPI 563247	Secale cereale	DEU, Mittelfranken	1947	n.a.	MT633927	MT633851	MT650004
	HAL 000026F	Secale cereale	DEU, Sachsen-Anhalt	1977	n.a.	n.a.	MT633866	n.a.
	BPI 563245	Secale cereale	USA, Wisconsin	1957	n.a.	MT633958	MT633852	MT650003
	BPI 563290	Triticum aestivum	AUS, New South Wales	1969	n.a.	n.a.	n.a.	MT650012
	DAOM 984746	Triticum aestivum	CAN, Ontario	2016	MT622311	MT633951	MT633803	MT650067
	DAOM 984747	Triticum aestivum	CAN, Ontario	2016	MT622312	MT633908	MT633828	MT650068
	DAOM 984748	Triticum aestivum	CAN, Ontario	2016	MT622313	n.a.	MT633827	MT650069
	DAOM 7738	Triticum aestivum	CAN, Nova Scotia	1936	n.a.	MT633925	MT633837	MT650017
	HAL 000036F	Triticum aestivum	CHN, Xinjian Uyg. Aut. Reg	1959	MT622309	n.a.	MT633804	n.a.
	BPI 563287	Triticum aestivum	CHN, Nanking	1932	n.a.	n.a.	n.a.	MT650010
	BPI 553981	Triticum aestivum (Triticum vulgare)	CHN, Kwangsi	1938	n.a.	n.a.	MT633864	MT649922
	BPI 563264	Triticum sp.	DEU, n.a.	1974	n.a.	MT633963	MT633848	MT650008
	BPI 563263	Triticum sp.	IND, Hebbal	1968	n.a.	n.a.	MT633849	MT650007
B. hordei
	BPI 859594	Agrostis exarata	USA, California	1939	n.a.	n.a.	n.a.	MT650013
	BPI 562851	Alopecurus aequalis	DEU, Bavaria	1946	n.a.	n.a.	n.a.	MT649934
	BPI 562908	Bromus hordeaceus subsp. hordeaceus (Bromus mollis)	IRL, Boyne Valley	1933	n.a.	n.a.	n.a.	MT649944
	BPI 562915	Bromus tectorum	USA, Nebraska	1941	n.a.	n.a.	n.a.	MT649946
	BPI 562987	Hordeum sp.	CHN, Chekiang	1931	n.a.	n.a.	n.a.	MT649962
	BPI 562996	Hordeum sp.	EGY, Gizo	1978	n.a.	n.a.	MT633858	MT649964
	BPI 563004	Hordeum murinum	ISR, Jerusalem	1935	n.a.	MT633943	MT633857	MT649966
	HAL 000010F	Hordeum murinum	UKR, Zaporizhia	1984	MT622304	MT633942	MT633807	MT650065
	BPI 563022	Hordeum vulgare	AUS, New South Wales	1977	n.a.	n.a.	MT633855	MT649969
	BPI 563021	Hordeum vulgare	AUS, Narrabri	1966	n.a.	n.a.	MT633894	MT649968
	DAOM 18154HT	Hordeum vulgare	CAN, Quebec	1940	MT622276	n.a.	MT633825	MT650018
	BPI 563037	Hordeum vulgare	IND, Panakpur	1956	n.a.	n.a.	n.a.	MT649972
	BPI 563033	Hordeum vulgare	IND, Katgodam	1948	n.a.	n.a.	MT633854	MT649970
	BPI 563039	Hordeum vulgare	MEX, Mixquiahuala	1947	n.a.	n.a.	MT633893	MT649974
	BPI 563035	Hordeum vulgare	USA, Texas	1944	n.a.	n.a.	MT633881	MT649971
	BPI 870274	Hordeum vulgare	USA, Pennsylvania	1944	n.a.	MT633926	MT633847	MT650014
	BPI 563019	Hordeum vulgare	USA, West Virgia	1954	n.a.	n.a.	MT633856	MT649967
	BPI 563048	Hordeum vulgare subsp. vulgare (Hordeum vulgare var.tetrastichon)	ROU, Ilfov-Bucuresti	1933	n.a.	n.a.	n.a.	MT649976
	BPI 562951	Leymus condensatus (Elymus condensatus)	USA, Wyoming	1941	n.a.	n.a.	n.a.	MT649954
	BPI 562830	Pseudoroegneria spicata (Agropyron spicatum)	USA, Washington	1935	n.a.	n.a.	n.a.	MT649931
	BPI 563286	Triticum aestivum	MEX, n.a.	1966	MT622274	n.a.	MT633904	MT650009
Blumeria sp.
	DAOM 82542	Deschampsia cespitosa	FIN, Lapponia enontekiensis	1958	MT622281	MT633922	n.a.	MT650023
	BPI 562901	Bromus japonicus	ROU, Lapusna-Cornesti	1931	n.a.	n.a.	n.a.	MT649942
	BPI 562983	Holcus lanatus	ROU, Vlasca-Comana	1930	n.a.	n.a.	n.a.	MT649960

a. Specimens in bold were included in the concatenated sequence analyses; superscript HT = holotype, ET = epitype.

b. Names in parentheses were records on specimen packets.

c. Three-letter country codes used: AUS Australia, CAN Canada, CHN China, CZE Czech Republic, DEU Germany, EGY Egypt, ETH Ethiopia, FIN Finland, GBR United Kingdom, DEU Germany, IND India, IRL Ireland, ISR Israel, JPN Japan, LVA Latvia, MEX Mexico, NLD Netherlands, ROU Romania, RUS Russia, UKR Ukraine, GBR United Kingdom, USA United States

Sequences of the 63 samples which were successfully sequenced for at least three loci were concatenated. With missing loci coded as gaps, the matrix resulted in 1639 characters. Added to this matrix were ten reference sequences of ITS and CHS from GenBank. The outgroup sequences used for individual locus analyses were concatenated as a composite outgroup taxon. The phylogeny generated from the concatenated alignment(Fig. 1)was congruent with each individual locus (Supplementary Figs. S1,2,3,4) in terms of the separation of the eight lineages, however had much higher statistical supports for those lineages and their internal branches. This is particularly evident for the B. americana and B. dactylidis clades, showing as paraphyletic on ITS and Bgt-1929 respectively due to low statistical supports. On all the phylogenetic trees, seven lineages corresponding to the described species here appear coherent, while B. graminicola included several sub-lineages that may be indicative of further cryptic speciation. This hypothesis could be evaluated by further studies of morphological or other biological characters.

3.2. Morphological observations

Variations in the following morphological characters were found between species, but also within species. A few features are diagnostic in some cases although many are shared by several species. It is recommended to use the key and DNA analyses for identification. For the lists of additional specimens examined for each described species in the taxonomy section, see Supplementary Appendix 1.

3.2.1. Primary mycelium

The development of all Blumeria spp. is initiated by the formation of the primary hyphae/mycelia (PH/PM), which are mostly flexuous, branched, septate, thin-walled, and smooth. Differences occur in the color of mycelia. In some species the mycelia remain white (whitish to greyish white) through the growing season, e.g. B. graminicola, while in other species become pigmented with age, showing as powdery patches in pale yellow (B. bromi-cathartici and B. bulbigera), orange to light brown (B. graminiss. str., B.americana, B. avenae and B. dactylidis ) or purplish brown ( B. hordei ). Another variation is the duration of primary mycelium development. In almost all species, the formation of the primary mycelium along with the asexual morph starts in spring (based on observations in temperate regions), and lasts until summer in parallel with secondary hyphae and chasmothecia. One exception is B. graminiss. str. in that the development of primary mycelia is hindered by the formation of the sexual morph. For B. graminicola, on the other hand, the development of the primary mycelia along with the asexual morph lasts until late autumn or early winter.

3.2.2. Hyphal appressoria

The hyphal appressoria are mostly nipple-shaped, occasionally lobe- or fork-shaped, 3–6 µm diam, singly or in opposite pairs on hyphal cells. The lobe-shaped morphology was observed in B. americana, B. dactylidis, B. graminiss. str.; fork-shaped in B. americana.

3.2.3. Secondary mycelium

The secondary mycelia (SM) usually set in around late spring to early summer, mostly along with the emergence of chasmothecia. The secondary hyphae (SH) are bristle-like, usually curved-falcate shaped, sometimes filiform, up to about 500 µm long and 3–7 µm wide, attenuated towards the tip or subcylindrical, soon becoming thick-walled; walls 1–2.5 µm wide, colorless and smooth; lumen narrow, 1–2 µm, later often “closed” (almost without any lumen). The shape and size of the SH are very characteristic and diagnostic for the genus Blumeria. However, among species, the variations are subtle including in pigmentation, septation and branching that are often shared by several species. For instance, the wall of the SH in B. graminiss. str., B. americana and B. hordei becomes pigmented with age, ranging from orange, ochraceous to brown; however, in other species (B. bulbigera, B. dactylidis, B. graminicola), remains whitish to dingy greyish white, or at most somewhat yellowish. Branched SH was observed in B. graminis s. str., more than two septa were observed in B. bulbigera and B. graminicola, but unbranched and aseptate (only one basal septum) otherwise.

3.2.4. Conidiophores and conidia

The conidiophores arise from the upper surface of hyphal mother cells, usually towards one end (septum), but occasionally in the middle, composed of a foot-cell with bulbous swelling, (2–)3–5(–7) shorter cells and a catenescent conidium chain with a terminal conidium (primary conidium, only with a basal hilum) and several catenate conidia (secondary conidia, with hila at the apex and base). Foot-cells 20–55 × 5–7 µm, bulbous swellings (8–)9–15 µm diam, basal septum at the junction with the supporting hypha or slightly elevated to 5(–10) µm; shorter cells, 10–30 µm long; conidia 20–35(–40) × (9–)10–16(–18) µm (dried herbarium material) or 24–40 × (10–)11–18(–20.5) µm (fresh material), length/width ratio 1.5–2.6(–3.1), hila (3–)4–6 µm wide, primary conidia broad ellipsoid-ovoid, secondary conidiabroad ellipsoid, barrel-shaped, oblong, fusiform, ovoid, ellipsoid to lemon-shaped (turgescent conidia usually broad ellipsoid, dry and older conidia often limoniform). The shapes and sizes of conidiophores and conidia are usually variable within species, and overlap between species. Other variations include the branched foot-cells and double foot-cells arising from one mother cell in parallel, as observed in B. americana, B. avenae and B. graminicola. The formation of pigmentation and the development of conidiophores are coincident with the primary hyphae. Blumeria graminicola is evidently distinct by producing the asexual morph until late fall or early winter, lasting much longer than other species, and remaining white through the growing season (also see primary mycelia section). This observation agrees with Blumer’s note on the powdery mildew on Poa spp. (Blumer, 1967). Different from most species, B. bromi-cathartici produces more uniform conidia, oblong, relatively long and narrow, 28–43 × 12–17.5 µm, with a length/width ratio of 1.7–3.2, and somewhat wider hila, 5–7.7 µm. Limoniform conidia have been observed in all Blumeria species. The conditions favoring this conidial shape are not quite clear, but age and turgescence seem to be factors. Fully turgescent conidia, including those gently heated in lactic acid, are usually less limoniform. SEM of the conidial apical wall shows a thickened patch with or without a dent in the middle for most species, however B. dactylidis does not have this characteristic.

3.2.5. Chasmothecia

Mature chasmothecia are gregarious or more or less scattered, mostly immersed in dense mycelial patches or layers, surrounded by bristle-like secondary hyphae, mostly dark brown to black, semi-globose or short cone-shaped with a flat top surface, with or without depression in the centre, range from 160 to 285 µm diam. Appendages are mycelioid, sparse, shorter than chasmothecium diam. Asci are broad ellipsoid, ellipsoid, obovoid, ovoid, pyriform, 47–73 × 30–58 µm, with or without a stalk. The stalk is up to 18 µm long, branched or unbranched. The asci of B. graminicola look more stout (wider) than in other species (excluding B. avenae and B. bromi-cathartici, not observed). The ascus stalks of B. bulbigera are evidently long and unbranched, B. americana branched and wavy.

3.3. Taxonomy

Blumeria Golovin ex Speer, Sydowia 27: 2, [1973–1974] 1975, nom. cons. [Art. 14.1, see (Braun, 2013)].

≡ Blumeria Golovin, Sborn. Rabot. Inst. Prikl. Zool. Fitopatol. 5: 124, 1958, nom. inval. (Art. 39.1).

≡ Erysiphe sect. Blumeria (Speer) U. Braun, Feddes Repert. 88: 659, 1978.

= Oidium Link, in Willd., Sp. pl. 4, 6(1): 121, 1824, nom. cons. (Art. 14) [type species – Oidium monilioides (Nees) Link, nom. sanct.], nom. rej. (Art. 56.1), see Braun (2013).

= Erysiphe sect. Bulbigera Sawada, Special Bull. Agric. Exp. Sta. Gov. Formosa 19: 149, 1919.

= Erysiphe sect. Graminis Homma, J. Fac. Agric. Hokkaido Imp. Univ. 38: 320, 1937, nom. inval. (Art. 39.1).

= Erysiphe sect. Monilioides S. Blumer, Echte Mehltaupilze (Erysiphaceae): 173, 1967, nom. inval. (Art. 39.1).

= Erysiphe auct. p.p.

Host and distribution: POACEAE subfam. ARUNDINOIDEAE tribe Molinieae, Molinia, Phragmites; subfam. BAMBUSOIDEAE tribe Arundinarieae, Phyllostachys; subfam. CHLORIDOIDEAE tribes Cynodonteae, Buchloe, Chloris, Cleistogenes, Cynodon, Dinebra, Leptochloa, Lepturus, Muhlenbergia; Eragrostideae, Eragrostis; Zoysieae, Spartina, Sporobolus; subfam. DANTHONIOIDEAE tribe Danthonieae, Danthonia, Schismus; subfam. ORYZOIDEAE tribe Oryzeae, Ehrharta, Leersia, Zizania; subfam. PANICOIDEAE, tribes Andropogoneae, Andropogon, Phacelurus, Saccharum; Paniceae, Cenchrus, Digitaria, Oplismenus, Panicum, Setaria; subfam. POOIDEAE tribes Brachypodieae, Brachypodium; Bromeae, Boissiera, Bromus; Diarrheneae, Diarrhena; Meliceae, Glyceria, Melica; Nardeae, Nardus; Poeae, Agrostis, Alopecurus, Ammochloa, Anthoxanthum, Apera, Arrhenatherum, Avena, Beckmannia, Briza, Calamagrostis, Catapodium, Coleanthus, Corynephorus, Cutandia, Dactylis, Deschampsia, Desmazeria, Dichelachne, Echinaria, Eremopoa, Festuca, Gastridium, Gaudinia, Helictochloa, Helictotrichon, Holcus, Koeleria, Lagurus, Lamarckia, Lolium, Macrochloa, Mibora, Milium, Nardurus, Parapholis, Phalaris, Phippsia, Phleum, Pholiurus, Pilgerochloa, Poa, Polypogon, Psilurus, Puccinellia, Rostraria, Sclerochloa, Sesleria, Sphenopholis, Trisetum, Vulpia; Stipeae, Achnatherum, Orthoraphium, Piptatherum, Stipa; Triticeae, Aegilops, Crithopsis, Dasypyrum, Elymus, Eremopyrum, Hordelymus, Hordeum, Leymus, Pascopyrum, Psathyrostachys, Pseudoroegneria, Secale, Taeniatherum, Thinopyrum, Triticale, Triticum, ×Aegilotriticum; Worldwide distribution.

Notes: Host range and distribution were based on collections examined, Braun and Cook (2012), Amano (1986), supplemented by data from numerous additional publications, including (Ershad, 1995; Sharma & Khare, 1995; Mendes et al., 1998; Crous, Phillips, & Baxter, 2000; Shin, 2000; Ahmad, Agarwal, Bambawal, & Puzari, 2007; Voytyuk, Heluta, Wasser, Nevo, & Takamatsu, 2009; Adhikari, 2017; U.S. National Fungus Collections Fungus-Host Database, https://nt.ars-grin.gov/fungaldatabases/fungushost/fungushost.cfm; accessed on May 25, 2020). The higher classification of host genera was sourced from GRIN Global Web v.1.10.6.2 (https://npgsweb.ars-grin.gov/gringlobal, accessed on 25 May 2020). Some host records only appeared once or twice, suggesting that either an occasional host jump might be involved or that the records were based on misidentifications. Several of such collections from Germany on unusual hosts have been examined, e.g. on Brachypodium pinnatum(L.) P. Beauv.(GLM-F48842), Digitaria sanguinea Weber (GLM-F49123), Echinochloa crus-galli (L.) P. Beauv.(GLM-F79561), Holcus lanatus L. (GLM-F96652, 104923), Melica ciliata L. (GLM-F48416), Molinia caerulea (L.) Moench (GLM-F51666), Sesleria albicans Deyl. (GLM-F47242), Setaria viridis (L.) P. Beauv. (GLM-F48849), Trisetum flavescens (L.) P. Beauv. (GLM-F94266), on which B. graminis s. lat. could not be found. There is a possibility that the host range is overstated.

The genus name Oidium Link (1824), typified with O.monilioides (Nees) Link, was previously conserved against Oidium Link (1809), with O. aureum (Pers.) Link as type (current name: Botryobasidium aureum Parmasto), basically in order to maintain the name Oidium for asexual morphs of powdery mildews.However, since the Melbourne code was in effect in 2012 (ICN, Turland et al., 2018), Oidium, typified by O. monilioides, an anamorph-typified name belonging to Blumeria, had to be considered an older heterotypic synonym of Blumeria. Therefore, Braun (2013) proposed to conserve the teleomorph-typified genus name Blumeria against Oidium. This proposal was approved by the General Committee and Blumeria is listed as a conserved name in the appendix of the ICN (Turland et al., 2018).

Blumeria graminis (DC.) Speer, Sydowia 27(1–6):2, 1975 [1973–1974], s. str. (emend.)

Fig. 2A–I.

≡ Erysiphe graminis DC., Fl. franç. 6: 106, 1815.

≡ Alphitomorpha communis γ graminearum Wallr., Verh. Ges. Naturf. Freunde Berlin 1: 31, 1819.

≡ Erysibe communis var. graminum Link, Sp. pl. 4, 6(1): 106, 1824.

≡ Erysiphe communis a. graminearum (Wallr.) Rabenh., Deutschl. Krypt.-Fl. 1: 232, 1844.

≡ E. communis z. graminis (DC.) Fr., Syst. mycol. 3: 242, 1829.

= Oidium tritici Lib., Pl. Crypt. Arduenna (Liège), Fasc. 4, no. 358, 1830. Lectotype (designated by Braun & Kirk, 2019: 88): on leaves of Triticum repens L. ( ≡Elymus repens (L.) Gould), sine loco et anno,Lib., Pl. Crypt. Arduenna 385 (PRM 685898); isolectotypes: Lib., Pl. Crypt. Arduenna 385 (e.g., BR, FH, G, ILLS 529, K, S-F49308).

≡ Torula tritici (Lib.) Corda, Icon. Fung. 5: 51, 1842, nom. illeg. (Art. 53.1), non Corda, 1837.

= Torula rubella Bonord., in Rabenh., Fungi Eur. Exs. (Klotzschii Herb. Viv. Mycol. Continuatio, Ed. Nova, Ser. Sec.), Cent. 3: no. 281, 1860 [Bot. Zeitung 19: 103, 1861; Flora 44: 158, 1861].

[Erysiphe graminis f. sp. tritici E. Marchal, Compt. Rend. Acad. Sci. (Paris) 135: 211, 1902.]

[Erysiphe graminis f. sp. secalis E. Marchal Compt. Rend. Acad. Sci. (Paris) 135: 211, 1902.]

Diagnosis: Development of the primary mycelium and the asexual morphs reduces with the onset of secondary mycelia and chasmothecia in late spring to early summer. The color of primary and secondary mycelia, as well as the asexual morph become yellowish, ochraceous to brownish later in the season. Branched secondary hyphae and lobed hyphal appressoria present.

Type: SWITZERLAND, Vaud, Gingins sur Nyon, on Triticum aestivum, 27 Jun 1998, A. Bolay (neotype, designated here, MycoBank no.: MBT392812, TNS-F87690).

Gene sequences ex-neotype (as isolate MUMH 1707 in GenBank): AB273542 (18S, ITS, 28S partial), AB273580 (CHS1).

Fig. 2 - Blumeria graminis s.str. A: Infected leaf surface with primary mycelia and conidiophores (DAOM 225780, asexual stage on the left), and secondary mycelia and chasmothecia (DAOM 74558, sexual stage on the right); B: Close-up of chasmothecia and secondary mycelia (DAOM 74558); C: A branched secondary hypha with basal septa (arrows, DAOM 74558); D: Conidiophore singly arising from mother cell (DAOM 225780, DAOM 161299); E: Asci ellipsoid or ovoid, with or without a stalk (DAOM 74558); F: Conidia, arrow shows germ tube (DAOM 225780); G: Hyphal appressoria (DAOM 225780, DAOM 161299); H: A secondary hypha unbranched, with one septum at the base (arrow, DAOM 74558); I: SEM of conidium apical wall thickened (on the left, DAOM 225780) and un-thickened (on the right, DAOM 161299). Bars: A 0.5 mm; B 100 μm; C–H 20 μm; I 2 μm.

Exsiccatae: On Elymus spp. – Constantinescu & Negrean, Herb. Mycol. Rom. 2763; Griff., West Amer. Fungi 164; Kari, Fungi Exs. Fenn. 22, 256, 682; Krieger, Fungi Saxon. Exs. 1669; Krypt. Exs. 1482; Lib., Pl. Crypt. Arduenna 358; Rabenh., Fungi Eur. Exs. 477; Săvul., Herb. Mycol. Rom. 2072; Solh., Mycofl. Saximont. Exs. 605, 606, 1119, 1319; Triebel, Microfungi Exs. 357. Secale cereale L.– Eriksson, Fungi Paras. Scand. Exs. 487; Krieger, Fungi Saxon. Exs. 1668; Syd., Mycoth. Germ. 1088; Thüm., Herb. Mycol. Oecon. 104, 151. Triticum aestivum – Crypt. Exs. 3346; Ellis & Everh., Fungi Columb. 505; Eriksson, Fungi Paras. Scand. Exs. 238; Flora Olteniae Exs. 536; Jack et al., Krypt. Badens 819; Kellerm. & Swingle, Kansas Fungi 1314; Krieger, Fungi Saxon. Exs. 1216; Krieger, Schädl. Pilze Kulturgew. 27, 122; Maire, Mycoth. Boreali-Africana 70; Petrak, Mycoth. Gen. 627; Rabenh., Herb. Viv. Mycol. 759; Rabenh., Fungi Eur. Exs. 671; P. Sacc., Mycoth. Ital. 6147; Săvul., Herb. Mycol. Rom., Fasc. 90, 1659; Seym. & Earle, Econ. Fungi 96; Smards, Fungi Latv. Exs. 573; Thüm., Fungi Austr. Exs. 238.

Mycelia on stalks, inflorescences and leaves, amphigenous, thin to dense, effuse or in patches; primary mycelia at first white, later pigmented (Fig. 2A); primary hyphal cells about 30–55 µm long and 3–7 µm wide; secondary mycelia (onset in late spring to early summer, mostly in Jun to Jul), dense woolly to felt-like, in patches, often around chasmothecia, dingy greyish white to grey, with age turning ochraceous, greyish brown to dingy brownish, sometimes rusty reddish brown (Fig. 2A, B); secondary hyphae about 200–500 × 3–7 µm, aseptate, branched or unbranched (Fig. 2C, H), thick-walled; SH walls1–2.5 µm; lumen later often pigmented, yellowish, ochraceous to brownish. Hyphal appressoria nipple-shaped, occasionally lobe-shaped, 3.5–7 µm wide, single or opposite in pairs (Fig. 2G). Conidiophores 60–170 × 4–7 µm, foot-cells (20–)25–35(–40) × 5–7 µm, bulbous swelling (8–)10–14(–15) µm wide, basal septum at the junction with the supporting hypha or slightly elevated to 8 µm, 4–6 µm wide at the basal septum; 3–5 shorter cells, 12–25 µm long (Fig. 2D); conidia (20–)24–35(–40) × (9–)12–16(–17) µm (herbarium material), (23–)28–40(–45) × (10–)14–18(–20.5) µm (fresh material), length/width ratio 1.6–2.5(–3.1), hila (3–)4–5(–6) µm wide (Fig. 2F); apical walls thickened with depression in centre, or not thickened (SEM, Fig. 2I); germ tubes showing a specific form of germination, with two types, lateral primary germ tubes one or more, narrow, short, about 0.5 times width of conidium lacking an appressorium, appear within 1 h, followed by a broader, lateral or terminal appressorial germ tube (see Braun & Cook, 2012 for instructions), straight or somewhat flexuous, 12–50 × 2.5–4 µm, extending to 1.25–3 times the width of the conidium and with an elongated swollen tip (not shown). Chasmothecia surrounded by bristle-like secondary hyphae (Fig. 2B), immature (100–)110–180 µm diam, mature 175–245(–260) µm diam; peridium cells obscure, irregularly polygonal, 8–20 µm diam; appendages few to numerous, in the lower half of the chasmothecium, mostly sparingly developed, mycelioid, simple, rarely irregularly branched, interlaced with the mycelium, short, usually shorter than the chasmothecial diam, thin-walled, hyaline to pigmented, aseptate to septate. Asci 6–30, subcylindrical to saccate, (50–)80–95(–105) × 20–45 µm, stalked or un-stalked (Fig. 2E), (4–)8-spored (ascospores rarely developed); ascospores ellipsoid-ovoid, 20–24 × 10–14 µm, colorless to faintly pigmented.

Host range and distribution: POACEAE primarily on tribe TRITICEAE, Aegilops, Dasypyrum, Elymus (including Hystrix), Hordeum, Secale, Triticum, also on tribe POEAE, Milium, Phleum, occasionally on tribe BRACHYPODIEAE, Brachypodium. Africa: Angola, Canary Islands, Ethiopia, Kenya, Libya, Malawi, Morocco, South Africa, Sudan, Tanzania, Zambia, Zimbabwe; Asia: Afghanistan, China, India, Iran, Iraq, Israel, Japan, Yemen, Kazakhstan, Korea, Kyrgyzstan, Lebanon, Myanmar, Nepal, Pakistan, Russia (Siberia, Far East), Saudi Arabia, Thailand, Turkey, Turkmenistan, Uzbekistan; Australia; Caucasus: Azerbaijan, Armenia, Georgia; Europe: throughout the continent; New Zealand; North America: Canada, Mexico, USA; Central & South America: Argentina, Brazil, Chile, Colombia, Ecuador, El Salvador, Guatemala, Nicaragua, Peru, Uruguay.

   Notes: De Candolle (1815) introduced Erysiphe graminis without any typification or reference to any host species. Braun (1987) searched for original collections of E. graminis in de Candolle’s herbarium in Genève (G) up to 1815, that could serve for lectotypification purposes, but was unsuccessful. Therefore, a specimen from herbarium G was designated as neotype [Germany, Bonn, Botanical Garden Poppelsdorf, on Triticum aestivum, 25 Jun 1869, F.A. Körnicke [1878] (G 00122110)]. De Candolle’s (1815) work on fungi of France was part of his investigation of Central European fungi, including powdery mildews. He did not cite exact host names for E. graminis, but noted this powdery mildew was on cereal crops. This background information was the motivation for Braun’s (1987) decision to designate a specimen on Triticum aestivum from Central Europe as the neotype. However, sequence analyses of powdery mildew samples in general, and very old specimens from the 19th century in particular, were not yet possible at that time. With our current protocols, we managed to sequence the previous neotype from 1869 (G 00122110). Much to our surprise, the retrieved sequence clustered far distant from the B. graminis (s. str.) clade and instead close to B. bromi-cathartici, together with sequences obtained from Blumeria sp. on Aegilops cylindrica Host (Armenia), Dasypyrum villosum (L.) P. Candargy (Romania), both also belonging to tribe Triticeae, and Piptatherum holciforme (M. Bieb.) Roem. & Schult. (Armenia) of tribe Stipeae (data not shown). The previous neotype designated by Braun (1987) was collected in a botanical garden. It is likely that the infection concerned was caused by an exotic powdery mildew, possibly originating from the Middle East. Results of our sequence analyses demonstrate that Braun’s (1987) neotypification of E. graminis is in conflict with de Candolle’s (1815) original concept, focusing on Central European species, which provides strong justification for rejecting that typification. In accordance with ICN (Turland et al., 2018, Art. 9.19 (c), Braun’s (1987) neotype is superseded here by the designation of a new neotype that is not in conflict with the protologue and stabilizes the application of E.graminis in the intended original sense.

Erysiphe graminis f. spp. tritici and secalis (Marchal, 1902) are not formal taxonomic units. Therefore, they are not ruled by the Code (ICN) and are only cited in square brackets at the end of the list of synonyms. As non-taxonomic names, formae speciales do not have type collections, i.e., status and affinity of such names cannot be verified by morphological re-examinations and gene sequencing. However, Marchal’s (1902) examinations were performed in Central Europe, where B. graminis is the principal Blumeria species found on Secale and Triticum species. Therefore, it is probable that Marchal (1902) had dealt with B. graminis, although host species of the Triticeae may also be infested by other Blumeria species.

Blumeria americana M. Liu, sp. nov.                                  Fig. 3A–H.

MycoBank no.: MB 835990.

Diagnosis: Development of primary mycelium from spring to summer, also in parallel with the secondary mycelium and chasmothecia, pigmented with aging, turning yellowish, ochraceous to rusty brown, the secondary mycelium often slightly lighter color than primary mycelia and asexual morph; hyphal appressoria nipple-, lobe- or fork-shaped, 1–2 conidiophores per hyphal mother cell, foot-cells branched or unbranched.

Type: CANADA, Alberta, Waterton Lakes National Park, Cameron Falls, on Elymus repens (as Agropyron repens (L.) P. Beauv.), 27 Aug 1980, J.A. Parmelee 5414 (holoype, DAOM 186037)

Gene sequences ex-holotype: MT622296 (ITS), MT633817 (Bgt-1929), MT650055 (Bgt-4572).

Fig. 3 - Blumeria americana sp. nov. A: Infected leaf adaxial (left) and abaxial surfaces (DAOM 156886) showing asexual stage; B:Infected leaf with sexual stage (DAOM 145059); C: Close-up of chasmothecia imbedded in secondary hyphae (DAOM 145059); D: Asci clavate, ellipsoid, ovoid with a sinuous or branched stalk (DAOM 137541); E: Aseptate secondary hyphae with narrow lumen; F: Young conidiophores arising from mother cells singly (top, DAOM 217856, DAOM 156886) or in pairs (bottom, DAOM 156886); G: Branched foot-cells (DAOM 186037); H: Mature conidiophores and conidia (DAOM 217856); I: Hyphal appressoria of various shapes; J: SEM view of conidia with scale-like artifacts on the surface and a thickened patch at apex (DAOM 217856). Bars: A, B 1 mm; C 100 μm; D 50 μm; E–I 20 μm; J 5 μm (including inset).

   Mycelia amphigenous, effuse or in patches, heavier on adaxial leaf surface; development of primary mycelia from spring to summer, persistent, often occurring in parallel with the secondary mycelium, i.e., neither inhibited nor discontinued with the onset of the development of secondary mycelium and chasmothecia, soon becoming pigmented, greyish orange, brownish orange to rusty brown (Fig. 3A, B); primary hyphae mostly branched in Y-shape, hyphal cells 27–42(–50) × 3–6(–8) µm; secondary mycelia formed from late spring or early summer to August, in dense patches, dingy greyish white to grey; secondary hyphae attenuate to apex with obtuse tips, 4–7 µm wide, wall thick, 1–2.5 µm; SH lumen 1–3 µm, or “closed”, in part yellowish, ochraceous to golden brown (Fig. 3E). Hyphal appressoria nipple-, lobe- or fork-shaped, single or opposite in pairs, 4–5 µm wide, 4–7 µm for lobe or fork-shaped (Fig. 3I). Conidiophores single or in pairs, 220–260 µm long; foot-cells 25–53(–66) µm long, bulbous swelling around middle 10–15 µm wide, branched or unbranched, basal septum 5–7 µm wide, at the junction with the mother cell or elevated up to 10 µm high; 1–3(–4) shorter cells, 13–30 × 4–8(–11) µm; up to 9 conidia per chain; primary conidia broad ellipsoid-ovoid, (18–)20–31(–34) × 10–18 µm, length/width ratio 1.5–2.5; secondary conidia broad ellipsoid, sometimes lemon-shaped, 14–26(–29) × 8.5–14 µm, length/width 1.5–2.1(–2.9), hila 5–7 µm wide; apical walls (SEM) with or without a thickened patch, no depression in the centre. Chasmothecia gregarious or scattered, immersed in woolly, dense secondary mycelia, semi-globose, upper surface depressed, later becoming concave, (110–)140–190(–220) µm diam; appendages sparingly developed, mycelioid; asci 18–28, oblong, ovoid, obovoid, (69–)75–100(–104) ×32–45 µm, stalk wavy or branching, 8–20 × 4.5–9 µm; ascospores not observed.

   Host range and distribution: POACEAE tribe TRITICEAE subtribe Hordinae, Elymus canadensis L., E. elymoides(Raf.) Swezey (= Sitanion hystrix (Nutt.) J. G. Sm.),E. glaucus Buckley, E. lanceolatus (Scribn. & J. G. Sm.) Gould (= Agropyron dasystachyum Ledeb.), E. repens ( ≡Agropyron repens), E. violaceus (Hornem.) Feilberg (= Agropyron latiglume (Scribn. & J. G. Sm.) Rydb.), Hordeum jubatum L., Leymus cinereus (Scribn. & Merr.) Á. Löve (≡ Elymus cinereus Scribn. & Merr.), Pascopyrum smithii (Rydb.) Barkworth & D. R. Dewey (= Agropyron smithii Rydb.) and Psathyrostachys juncea (Fisch.) Nevski (≡ Elymus junceus Fisch.); tribe POEAE subtribe Poinae, Apera spica-venti (L.) P. Beauv. North America: Canada, USA.

   Notes: A number of samples on Elymus grouped in this species, predominantly from North America. However, Elymus can also be infected by B. graminis s.str. and B. graminicola, so it is important to not depend solely on the host for identification. A sequence retrieved from Blumeria on a Deschampsia cespitosa (L.) P. Beauv. sample from Finland appears closely related with B. americana on the ITS tree (Supplementary Fig. S1),but was not confirmed to be a member in other analyses. Previously, Deschampsia belonged to the same subtribe (Holcinae) as Holcus, the host of B. graminicola (more information follows).Examination of additional samples on Deschampsia cespitosa from Germany (GLM-F48875, 49967, 50142, 56429, 58758, 63460, 63463) showed the primary mycelia, conidiophores and conidia turn yellowish, ochraceous to brownish (different from B. graminicola). Furthermore, a recent grass phylogenetic study (Soreng et al., 2017) separated Deschampsia from other members of subtribe Holcinae and erected a new subtribe Aristaveninae. For the time being, Blumeria on Deschampsia cespitosa can only be referred to as Blumeria sp. and is in urgent need of further genetic examination.

Blumeria avenae M. Liu & Hambl., sp. nov.                            Fig. 4A–I.

MycoBank no.: MB 835993.

[Erysiphe graminis f. sp. avenae E. Marchal, Compt. Rend. Acad. Sci. (Paris) 135, 211, 1902.]

Diagnosis: Development of primary mycelium from spring to autumn (Nov), sometimes even in winter (Jan), whitish to greyish white, turning yellowish, orange to brown, 1–2 conidiophores arise from the same mother cell, parallel or in different directions; hyphal appressoria mostly nipple-shaped, single, occasionally in opposite pairs.

Type: UNITED KINGDOM, England, Exeter, on Avena sativa L., 06 Feb 1948, A.S. Boughey (holotype, DAOM 236220; isotype, IMI 24011).

Gene sequences ex-holotype: MT622301 (ITS), MT633815 (Bgt-1929),MT650063 (Bgt-4572)

Exsiccatae: On Avena spp. – Barthol., Fungi Columb. 3151; Briosi & Cavara, Fung. Paras. Piante Colt. Util. Ess. 174.

Fig. 4 - Blumeria avenae sp. nov. A: Various stages of infection on the leaf surface with light orange to brownish mycelia (DAOM 236220, 147852); B: Close-up of conidiophores on leaf surface (DAOM 236220); C: Conidia, note two with germ tubes at or near apices (DAOM 236220, 147852, arrows); D: Appressoria nipple-shaped (DAOM 236220, 147852), single or in opposite pairs; E–F: Single conidiophore arising from mother cell, the septum between mother cell and foot cell elevated severely (E, DAOM 236220, arrow) or slightly (F, DAOM 147852); G: Two conidiophores arising from mother cell in pairs (DAOM 236220); H: A cluster of conidiophores arising from irregular shaped mother cells (DAOM 236220); I: SEM close up apical wall of conidia. Bars: A 0.5 mm; B 100 μm; C–H 20 μm; I 2 µm.

Mycelia on stalks and leaves, amphigenous, effuse, thin or in dense patches; primary myceliaat first whitish, later pigmented (Fig. 4A); primary hyphae with hyphal cells 2–8 µm wide; secondary mycelia develops alone or along with the asexual morph in late spring or early summer, often forming dense woolly to felt-like patches around chasmothecia, dingy greyish white to grey, with age sometimes faintly pigmented, bristle-like secondary hyphae 3–6.5 µm wide, aseptate, wall 1–2 µm thick; SH lumen narrow, 1–2 µm, hyaline, occasionally becoming yellowish, ochraceous to brownish. Hyphal appressoria nipple-shaped, single or in opposite pairs, 3–6 µm wide (Fig. 4D). Conidiophores usually near or next to one septum, erect, single or in pairs (Fig. 4E–H), 60–150 × (4–)5–7(–8) µm, foot-cells (25–)30–45 × 5–6 µm, bulbous swelling 10–14 µm wide, basal septum at the junction with the mother cell or often elevated, 5–13(–25) µm, (4–)5–7(–10) µm wide at the basal septum; (1–)2–4(–5) shorter cells, 12–35 µm long; conidia broad ellipsoid, ellipsoid, ovoid, occasionally lemon-shaped, 20–35(–40) × 10–17 µm (herbarium material), 24–38.5(–44) × 11–19 µm (fresh material), length/width ratio 1.4–2.7(–3.3), hila (4–)5–7 µm wide (Fig. 4C); apical walls severely thickened patch with depressed centre, or not thickened (Fig. 4I). Chasmothecia100–155 µm diam when immature, 160–200 µm diam when mature(observed by UB, not shown in Figs).

Host range and distribution: POACEAE tribe POEAE subtribe Aveninae, Avena. Africa: Canary Islands, Lebanon, Libya, Morocco, South Africa, Zimbabwe; Asia: China, India, Iran, Iraq, Israel, Turkey, Turkmenistan, Uzbekistan; Australia/Oceania: Australia, New Zealand; Caucasus: Azerbaijan, Georgia; Europe: throughout the whole continent; North America: Canada, Mexico, USA; Central & South America: Argentina, Brazil, Chile, Colombia, Guatemala, Peru, Uruguay.

Notes: B. avenae appears only on Avena spp., and is closely related with B. dactylidis (see later description). Briosi & Cavara, Fung. Paras. Piante Colt. Util. Ess. 174, noted “on Avena sativa and Hordeum vulgare L.”. The identity of the leaves in the examined duplicate preserved at HAL has been checked and proved to belong to Avena sativa.

Blumeria sp. on Koeleria pyramidata (Lam.) P. Beauv. [= K. macrantha(Ledeb.) Schult.] (material examined: Germany, GLM-F46754, 48469, 48501, 55994, 56907, 57294, 57300, 57354, 58720) has been examined and is characterized as follows: the primary mycelium quickly becomes brownish; hyphae, conidiophores and conidia turn yellowish, ochraceous to brownish. Koeleria belongs in tribe Poeae subtribe Aveninae (Soreng et al., 2017), suggesting that these infections might be caused by Blumeria avenae. However, a phylogenetic confirmation is still necessary.

Blumeria bromi-cathartici S. Takam. & M. Liu, sp. nov.                      Fig. 5A–H.

MycoBank no.: MB 835994.

Diagnosis: Conidia mostly oblong, 28–43 × 12–17.5 µm, long and narrow with a length/width ratio of 1.7–3.2.

   Type: JAPAN, Mie, Tsu-shi, Mie University, on Bromus catharticus Vahl, 25 Apr 1995, S. Takamatsu (holotype, TNS-F87248).

Gene sequences ex-holotype (TNS-F87248 (recorded as MUMH 0117 in GenBank)): AB000935 (ITS), AB022362 (28S), AB033475 (18S).

Fig. 5 - Blumeria bromi-cathartici sp. nov. (TNS-F87248 = MUMH 0117). A: Infection on leaf surface showing light yellow to orange primary mycelia and conidiophores; B: Close-up ofconidiophores on leaf surface; C: Hypha branching at a narrow angle, and hyphal appressoria nipple-shaped and single; D–E: Conidiophores at different developing stages singly arising from mother cell; F: Primary conidia mostlyoblong; G: Blumeria-type germination of a conidium; H: SEM view of apical walls. Bars: A 0.5 mm; B 100 μm; C–G 20 μm; H 2 µm.

Primary mycelia amphigenous, thick and persistent, white, later yellowish white to greyish yellow (4A2–4B2; Fig. 5A, B); hyphae almost straight to somewhat sinuous, 4–6(–7) um wide, branching at narrow angle, with a septum near the branching point; hyphal appressoria well-developed, nipple shaped, single (Fig. 5C). Secondary hyphae bristle-like, curved-falcate, thick-walled. Conidiophores solitary, erect, 73–119 µm long, simple, straight; foot-cells(27–)30–50(–60) µm long bulbous swelling 9.5–12.5 µm wide, basal septum at the junction with the mother cell, basal septum 4.7–7.7 µm wide; 2–4(–5) straight shorter cells, 50–100 µm long and 7–10 µm wide, 2–8 immature conidia in chains (Fig. 5D, E); conidia oblong, occasionally limoniform, 28–43 × 12–17.5 µm, length/width ratio 1.7–3.2, hila 5.0–7.7 µm wide (Fig. 5F), producing germ tube on the shoulder, germ tubes Blumeria type (Fig. 5G); apical walls(SEM) mostly not thickened, occasionally with a slightly thickened patch (Fig. 5H). Chasmothecia not observed.

Host range and distribution: POACEAE subfam. POOIDEAE tribe BROMEAE, Bromus catharticus. Asia, Japan. North America, United States

   Notes: Inuma et al. (2007) revealed three lineages on Bromus using multi-locus phylogenetic analyses. This species, representing the lineage specific on B. catharticus and including two samples from Japan and one from United States (CA), was clearly separated from a Eurasian species, B. bulbigera (see next species described) which was on various Bromus spp. The third lineage was a sequence retrieved from a specimen collected in Argentina (MUMH2192), had an uncertain affinity to other lineages, and might represent a different species. Morphologically, B. bromi-cathartici is characterized by having longer conidia (mostly oblong), and restricted host ranges. Amano (1986) recorded B. graminis on Bromus catharticus from Europe (Germany), North America (USA), and South America (Argentina, Brazil). In an ITS tree with extended samples (data not shown), several sequences retrieved from Blumeria on several hosts belonging to Triticeae, i.e. Aegilops cylindrica (Armenia), Dasypyrum villosum (Romania), Triticum aestivum (Germany, historic sample from the 19th century, collected in a botanical garden) and a single host species in Stipeae, i.e. Piptatherum holciforme (Romania), cluster close to B. bromi-cathartici. The status of those collections remains unclear. They can currently only be referred to as Blumeria sp. They seem to represent a distinct lineage and undescribed species, but further morphological and phylogenetic analyses are required.

Blumeria bulbigera (Bonord.) M. Liu & U. Braun, comb. nov.                Fig. 6A–J.

MycoBank no.: MB 835998.

Basionym: Torula bulbigera Bonord., in Rabenh., Fungi Eur. Exs. (Klotzschii Herb. Viv. Mycol. Continuatio, Ed. Nova, Ser. Sec.), Cent. 2: no. 175, 1860 [also in Bot. Zeitung 18: 175, 1860; Flora 43: 748, 1860].

≡ Oidium bulbigerum (Bonord.) Sacc. & Voglino, in Sacc., Syll. Fung. 4: 47, 1886.

= Botrytis simplex β monilis Alb. & Schwein., Consp. fung. lusat.: 363, 1805. Type not designated (“in foliis culmisque, … graminum (Bromi mollis etc.). Neotype (designated here, MycoBank no.: MBT392810): Germany, Saxony, Königstein, on Bromus hordeaceus L. (= B. mollis L.), 28 Jun 1905, W. Krieger, Fungi Saxon. Exs. 1921 (HAL, s.n.). Isoneotypes: Krieger, Fungi Saxon. Exs. 1921 (e.g., B, BPI 562907, M-13677, etc.).

= Oidium monilioides var. ochraceum Thüm., Fungi Austr. Exs. 1084, 1874. Lectotype (designated here, MycoBank no.: MBT392811): Czech Republic, Bohemia, Teplice (Tepliz), on Bromus hordeaceus, 1873, F. v. Thümen, Thüm., Fungi Austr. Exs. 1084 (HAL, s.n.). Isolectotypes: Thüm., Fungi Austr. Exs. 1084 (e.g., BPI 409682, ILL 85997).

≡ Oidium ochraceum (Thüm.) Mussat, in Saccardo, Syll. fung. (Abellini) 15: 231, 1901.

[Erysiphe graminis f. sp. bromi E. Marchal Compt. Rend. Acad. Sci. (Paris) 135, 212, 1902.]

Fig.6 – Blumeria bulbigera comb. nov. A: Infection on adaxial leaf surface showing yellow primary mycelia and conidiophores (DAOM 82541); B: White secondary mycelia and chasmothecia produced on adaxial leaf surface (GLM-F54349); C: Exposed disk-shaped chasmothecia with deep depression on upper surfaces (GLM-F54349); D: Asci with cylindrical stalk (GLM-F54349); E: Conidiophore arising singly from irregular mother cell (DAOM 82541); F: Nipple-shaped appressoria arising singly or in opposite pairs, ovoid swelling at the end or middle of hyphae (see arrows, DAOM 82541, GLM-F62360); G: Conidia of subglobose, ovoid, oblong, and lime-shaped, germ tubes produced near the apices (DAOM 82541, GLM-F62360); H: Secondary hypha with several septa (see arrows, GLM-F54349), I: SEM view of conidia; J: Apical wall of conidia (DAOM 82541). Bars: A–C 100 μm; D–H 20 μm; I 5 μm; J 2 µm.

Diagnosis: Development of the primary and secondary mycelium from spring to late summer, or to autumn (Oct); primary mycelia and asexual morph turning pale yellow to ochraceous with age; ovoid swellings may present in the middle or at the end of primary hyphae; secondary hyphae multi-septate, lumen yellowish to ochraceous with age.

Type: GERMANY, Rhine-Westphalia (Guestphalia), Herford, “in foliis graminum” (Bromus hordeaceus), Rabenh., Fungi Eur. Exs., Cent 2, no. 275 (lectotype, designated here, MycoBank no.: MBT392798, B, s.n.; isolectotypes, Rabenh., Fungi Eur. Exs., Cent 2, no. 275 (e.g., L9102521067, HAL s.n). FINLAND, Regio aboënsis, Uusi-Kaupunki, on Bromus hordeaceus subsp. hordeaceus(= B. mollis), 13 Aug. 1957, L. & K. Roivainen (epitype, designated here, MycoBank no.: MBT 392801, DAOM 82541); gene sequences ex-epitype MT622280 (ITS).

Exsiccatae: On Bromus spp. – Bucholtz, Fungi Ross. Exs., Ser. A, 84; Griff., West Amer. Fungi 101; Krieger, Fungi Saxon. Exs. 1921; Rabenh., Fungi Eur. Exs. 81, 175; Săvul., Herb. Mycol. Rom. 1871, 1872, 2273; Solh., Mycofl. Saximont. Exs. 607.

Mycelia on stalks and leaves, amphigenous, thin to dense, effuse or in patches (inconspicuous on B.hordeaceus, only seen in close-up); primary mycelia at first white, often turning pale yellow, ochraceous with age, in patches or effuse thin layers (Fig. 6A); primary hyphae cells 3–7 µm wide, occasionally with swollen portions, up to 10 µm wide (Fig. 6F); secondary hyphae 4–7 µm wide, mostly aseptate, occasionally 2–3 septa (Fig. 6H),wall 1–2.5 µm thick; SH lumen narrow, 1–2 µm, or “closed”, hyaline, sometimes slightly pigmented, yellowish white to pale yellow (3A2–3A3), ochraceous. Conidiophores erect, mostly close to one septum, occasionally in centre, 80–130 × 5–6 µm, foot-cells 25–45 × 5–6 µm, bulbous swelling 9–14 µm wide, basal septum at the junction with the supporting hypha or slightly elevated, to 5 µm, 5–6(–8) µm wide at the septum; 2–4 shorter cells, 10–30 µm long (Fig. 6E); conidia ellipsoid, broad ellipsoid, ovoid, to limoniform, 25–32 × 12–16 µm (herbarium material), 27–35 × 13–18.5 µm, hila 3–5 µm wide, length/width ratio 1.5–2.5 (Fig. 6G); apical walls with a thickened ring, or thickened with depressed centre (SEM; Fig. 6I, J). Development of chasmothecia from spring to summer, loosely surrounded by secondary hyphae, or exposed (Fig. 6B, C), 100–160 µm diam when immature, 150–200 µm when mature; asci ovoid with a short cylindrical stalk 10–16 × 5–7 µm (Fig. 6D); ascospores not observed.

Host range and distribution: POACEAE subfam. POOIDEAE tribe Bromeae,Bromus spp. Africa: Canary Islands, Morocco, West Sahara; Asia: Afghanistan, China, Iran, Iraq, Israel, Jordan, Kazakhstan, Korea, Russia – Siberia, Syria, Turkey, Turkmenistan, Uzbekistan; Australia; Caucasus: Armenia, Azerbaijan, Georgia; Europe: widespread in the whole continent; New Zealand; North America: Canada, USA; South America: Argentina, Chile.

Notes: Blumeria bulbigera seems to be confined to Bromus species and distributed in Eurasia and North America. The broad distribution can probably be explained by a co-distribution with the wide synanthropic occurrence of several Bromus species as neophytes. The records of B. graminis s. lat. on diverse Bromus species (Amano, 1986; Braun, 1995https://nt.ars-grin.gov/fungaldatabases/specimens/Specimens.cfm) need to be confirmed by molecular methods and morphological re-examination.

Blumeria dactylidis M. Liu & Hambl., sp. nov.                            Fig. 7A–K.

MycoBank no.: MB 835995.

= Erysiphe graminis f. dactylidis-glomeratae Sacc., Mycoth. Ven. 606, 1876.

= Erysiphe graminis f. dactylidis Jacz., Karm. Opred. Grib., Vip. 2. Muchn.-rosj. griby (Leningrad): 149, 1927.

= Erysiphe graminis f. alopecuri Jacz., Karm. Opred. Grib., Vip. 2. Muchn.-rosj. griby (Leningrad): 143, 1927.

= Erysiphe graminis f. anthoxanthi Jacz., Karm. Opred. Grib., Vip. 2. Muchn.-rosj. griby (Leningrad): 143, 1927.

[Erysiphe graminis f. sp. dactylidis Okuda et al., Ann. Phytopathol. Soc. Japan 51: 615, 1985.]

Fig. 7 - Blumeria dactylidis sp. nov. A: Infection on adaxial leaf surface showing brown primarymycelia and conidia production (left, DAOM 91654), and on abaxial leaf surfaceshowing greyish white secondary mycelia and chasmothecia (right, GLM-F50119); B:Close-up of conidia chains (DAOM 118220); C: Chasmothecium side view (upper)and overview (lower) showing the deep depression at the upper surface and thesurrounding white secondary mycelia (GLM-F79570); D: Secondary hyphae aseptateand unbranched (GLM-F50119); E: Appressoria in nipple and lobe shapes (DAOM91654, 118220); F–H showing one conidiophore per mother cell (DAOM 91654): F:Conidiophore with elevated septum between mother cell and foot cell, G: Youngfoot cell, H: Conidiophore with slightly elevated septum between the mother celland foot cell; I: Conidia, producing one or two germ tubes (DAOM 91654, DAOM118220, arrows); J: Asci with a slightly protruding stalk (GLM-F79570); K: SEM view of apical wall of conidiaun-thickened. Bars: A 0.5 mm; B, C 100 μm; D–J 20 μm; K 2 μm.

Diagnosis: Primary mycelia and asexual morph turning yellowish, ochraceous to brownish, lasting until autumn (early Oct) in parallel with the secondary mycelium and chasmothecia; secondary mycelia white to greyish white, or orange grey, much lighter than primary mycelia, aseptate; hyphal appressoria nipple- or lobe-shaped, single; conidial apical walls not thickened.

Type: CANADA, British Columbia, North Saanich, on Dactylis glomerataL., 12 Apr 1934, W. Jones, as ‘W.J.’ (holotype, DAOM 118220).

Gene sequences ex-holotype: MT622286 (ITS), MT633919 (CHS1), MT633812 (Bgt-1929), MT650032 (Bgt-4572).

Exsiccatae: On Dactylis spp. – Mäkinen, Fungi Exs. Fenn. 682; Sacc., Mycoth. Venet. 606; Săvul., Herb. Mycol. Rom. 2272; Vill, Fungi Bavar. 962.

Mycelia on stems and leaves, mainly on adaxial leaf surface, or amphigenous, thin to dense, effuse or in patches, primary mycelia along with asexual morph at first white, turning pale yellow (4A2), ochraceous to brown (5A2–5E4; Fig. 7A, B); secondary mycelia development starts mostly in Jun to Jul, lasting until late fall in parallel with primary mycelia and asexual morphs, dingy greyish white to grey, turning ochraceous to pale dingy, orange grey with age (Fig. 7A); primary hyphal cells 3–6 µm wide, occasionally with swollen portions, up to 9 µm wide; hyphal appressoria nipple-, sometimes lobe-shaped, 3–7 µm wide, lobe-shaped up to 10 μm wide (Fig. 7E); secondary hyphae about 500 μm long, in dense woolly to felt-like patches around chasmothecia (Fig. 7C, D), aseptate, wall 1–2 µm thick, central lumen narrow, 0–2 µm, hyaline, lumen usually not pigmented; conidiophores single, erect, 65–135 × 5–6 µm; foot-cells (25–)30–50 × 5–6 µm, bulbous swelling 9–13 µm wide, basal septum at the junction with the supporting hypha or slightly elevated up to 8 µm, 4–7 µm wide at the septum; 3–4 shorter cells, 12–25 µm long (Fig. 7F–H); conidia ellipsoid, broad ellipsoid, ovoid, dolliform, rarely lemon-shaped (Fig. 7I), 21–35 × 11–15 µm (herbarium material), 23–38 × 12–18 µm (fresh material), length/width ratio 1.5–2.6(–2.9), hila 4–7 µm wide; conidial apical walls not thickened (SEM; Fig. 7K). Chasmothecia(100–)140–180 µm diam when immature, 160–245 µm diam when mature; asci obovoid-clavate to saccate, (45–)60–90 × 25–40 µm, stalks very short, inconspicuous (Fig. 7J); ascospores not observed.

Host range and distribution: POACEAE tribe POEAE subtribe Dactylidinae, Dactylis glomerata s. lat. as principal host, less frequently on subtribe Anthoxanthinae, Anthoxanthum odoratum L. and A. aristatum Boiss., occasionally subtribe Loliinae, Festuca pratensis Huds. and Lolium multiflorum Lam., subtribe Poinae,Phleum sp.; tribe BROMEAE, Bromus; tribeTRITICEAE subtribe Hordinae, Hordeum. North Africa: Morocco; Asia: Israel, Japan, Kazakhstan, Korea, Kyrgyzstan, Russia – Siberia, Turkey, Turkmenistan, Uzbekistan; Caucasus: Armenia, Azerbaijan, Georgia; Europe: throughout the whole continent; North America: Canada, USA.

Notes: The principle host genus of this species is Dactylis (tribe Poeae) with Dactylis glomerataas the main host, but it can also be found on other genera in the same tribe, i.e, Anthoxanthum, Festuca, and Phleum. Erysiphe graminis f. sp. dactylidis, introduced by Oku, Yamashita, Doi, and Nishihara (1985) based on inoculation experiments with Dactylis glomerata performed in Japan, undoubtedly refers to Blumeria dactylidis. A wide array of grasses from numerous genera were included in their examinations, but Blumeria on Dactylis glomerata in Japan was strictly confirmed to this host species.

For Blumeria on Festuca gigantea (L.) Vill.(material examined: Germany, GLM-F54303, 63457, 79631, 90974, 99586, 96672, 102932), the primary mycelium quickly becomes ochraceous to pale brownish. Festuca gigantea is a broad-leaved species of Festuca s. lat. This genus belongs in tribe Poeae subtribe Loliinae, which is close to subtribe Dactylidinae (Soreng et al., 2017), suggesting that B. dactylidis might be the causal agent of infections on F. gigantea. However, a phylogenetic confirmation is necessary. BPI 562975 on F. pratensis from the Czech Republic and HAL 000025 F on F. gigantea from Germany belong to B. dactylidis based on the Bgt-1929 tree. The position of BPI 562975 was also confirmed by Bgt-4572 and CHS1 trees. However, Festuca spp. can also be infested by B. graminicola (see notes under this species).

A rarer case is that two samples on Bromus from Germany (HAL 000027 F) and USA (BPI 562894) were also grouped in this clade, the former on Bgt-1929 and ITS tree, the latter on Bgt-4572, indicating the host range potential is across different tribes.

Blumeria graminicola M. Liu & Hambl., sp. nov.                         Fig. 8A–K.

MycoBank no.: MB 835996.

= Erysiphe graminis f. aperae Jacz., Karm. Opred. Grib., Vip. 2. Muchn.-rosj. griby (Leningrad): 143, 1927.

= Erysiphe graminis f. milii Jacz., Karm. Opred. Grib., Vip. 2. Muchn.-rosj. griby (Leningrad): 152, 1927.

= [Erysiphe graminis f. sp. poae E. Marchal, Compt. Rend. Acad. Sci. (Paris) 135, 210-212, 1902.]

Fig. 8 - Blumeria graminicola sp. nov. A: Chasmothecia covered with white secondary hyphae oninfected leaf surface (DAOM 179493); B: Primary hyphae and conidia (arrow)co-exist with secondary hyphae and chasmothecium (DAOM 159510); C: Filiform appendages of chasmothecium with a septum at the base or near base (arrows,DAOM 159510); D: Asci subglobose, or ovoid with an unbranched short stalk (DAOM159510); E: Secondary hyphae with septa (arrow, DAOM 179493); F: Single conidiophore arising from one mother cell (DAOM 159510); G: Hyphal appressoria(DAOM 181460, DAOM 91194); H: Two conidiophores arising from one mother cell,and branched foot-cell (arrows, DAOM 179493, DAOM 159510); I: Conidia, note oneconidium producing a germ-tube; J: SEM view of scale-like artifacts on surfaceof conidia, thickened apical wall with depressed center (DAOM 24040); K:Close-up of apical wall of conidia (DAOM 91194). Bars: A 0.5 mm; B 100 μm; C–I 20 μm; J 5 μm; K 1 μm.

Diagnosis: Development of primary mycelium and asexual morph until winter (Jan/Feb), not inhibited with the onset of secondary mycelium and chasmothecia (May); primary and secondary mycelia and asexual morph whitish to greyish white, not distinctly pigmented with age; secondary hyphae septate; hyphal appressoria nipple-shaped, lobe-shaped, single; conidial apical wallsseverely thickened depressed in centre.

Type: CANADA, Ontario, Ottawa, Central Experiment Farm, Lawn, on Poa pratensis L., 02 Aug 1974, J. A. Parmelee (holotype, DAOM 159510).

Gene sequences ex-holotype: MT633954 (CHS1), MT633820 (Bgt-1929), MT650048 (Bgt-4572).

Exsiccatae: On Apera spica-venti– Jack et al., Krypt. Badens 829; Krieger, Fungi Saxon. Exs. 1217; Syd., Mycoth. Germ. 1529; Thüm., Fungi Austr. Exs. 1244. On Milium effusum L.: Lundell & Nannf., Fungi Exs. Suec. 1470. On Poa spp. – W.B. Cooke, Mycobiota Mt. Shasta 26; Griff., West Amer. Fungi 102; Liro, Mycoth. Fenn. 623; Ravenel, Fungi Amer. Exs. 308; Ravenel, Fungi Carol., Fasc. II, 85; Săvul., Herb. Mycol. Rom., Fasc. 1777, 1875; Schneider, Herb. Schlesischer Pilze 920; Solh., Mycofl. Saximont. Exs. 1121, 1318; Thüm., Herb. Mycol. Oecon. 123; Thüm., Mycoth. Univ. 257; Vill, Fungi Bavar. 961.

Primary mycelia on leaves, amphigenous, occasionally on stalks, thin to dense, effuse or in patches, development from spring (Apr/May) to autumn (Nov), sometimes even in winter (Jan/Feb), neither inhibited nor discontinued with the onset of secondary mycelium and chasmothecia (May); whitish to greyish white, at most faintly yellowish, not distinctly pigmented with age (Fig. 8A, B); primary hyphal cells 2–6 µm wide; hyphal appressoria nipple-shaped, occasionally lobe-shaped, single or in opposite pairs (Fig. 8G); secondary hyphae 3–7 µm wide, with 1–2 septa (Fig. 8E), thick-walled, wall 1–2 µm thick; central lumen narrow, 0–2 µm, usually hyaline. Conidiophores single or in pairs, close to one septum, erect, 70–150 × 5–7 µm (Fig. 8F, H), mother cells occasionally swollen up to 12 µm wide, foot-cells (20–)25–45(–55) × 5–7 µm, bulbous swelling (8–)9–13(–14) µm wide, basal septum at the junction with the supporting hypha or slightly elevated up to 5(–10) µm, 4–7 µm wide at the septum; (2–)3–5(–7) shorter cells, 12–25 µm long; conidia broad ellipsoid to lemon-shaped, 19–33 × 10–15 µm (herbarium material), 29–35 × 11–17.5 µm (fresh material), hila 4–6 µm wide (Fig. 8I); apical walls severely thickened depressed in centre (Fig. 8J, K). Chasmothecia 100–150 µm diam when immature, 140–200 µm when mature; appendages sparse, mycelioid, with a basal septum (Fig. 8C); asci subglobose to ovoid, stalk inconspicuous, or short, branched or unbranched; ascospores not observed (Fig. 8D).

Host range and distribution: POACEAE tribe POEAE subtribe Poinae, Apera spica-venti, Poa spp. as principle hosts, but also on subtribe Agrostidinae, Agrostis sp., Polypogon monspeliensis (L.) Desf., subtribe Alopecurinae, Alopecurus geniculatus L., Beckmannia spp., subtribe Anthoxanthinae, Anthoxanthum nitens (= Hierochloe odorata), subtribe Coleanthinae, Puccinellia spp, subtribe Dactylidinae, Dactylis glomerata, subtribe Holcinae, Holcus lanatus, subtribe Loliinae, Festuca spp, subtribe Miliinae, Milium effusum; tribe TRITICEAE subtribe Hordeinae,Elymus sp., subtribe Triticinae, Thinopyrum intermedium (Host) Barkworth & D. R. Dewey, Triticum aestivum (only one sample from Australia);tribe BROMEAE, Bromus spp.; tribe MELICEAE, Melica msubulata (Griseb.) Scribn. Africa: Morocco; Asia: Afghanistan, China, India, Iran, Israel, Japan, Kazakhstan, Korea, Kyrgyzstan, Mongolia, Russia (Siberia, Far East), Turkey, Turkmenistan, Uzbekistan; Caucasus: Armenia, Azerbaijan, Georgia, Europe: throughout the continent; New Zealand; North America: Canada, USA; South America: Argentina.

Notes: All the studied samples on Poa spp. grouped in this clade, agreeing with the previous recognition of f. sp. poae by Marchal (1902), based on examinations of central European specimens. A remarkable feature of this species is its persistent white color of mycelia and asexual morphs throughout the growing season.Blumer (1967) already noted the difference between the white mycelium of the powdery mildew on Poaspp.and the pigmented mycelia on other hosts. Poa is the largest grass genus with around 500 species distributed in cool temperate regions (Kellogg, 2015), representing a large pool of genetic diversity. It is very likely that this large genetic diversity set the stage for B. graminicola to also diversify and adapt to different host species in Poa and also further to a wider range of host genera. The DNA sequence analyses demonstrate B. graminicola has the highest nucleotide diversities for all four loci compared with other species (data not shown). Chasmothecia are commonly formed on Apera spica-venti (development beginning in May).

Species of Festuca are hosts of several Blumeria spp., including B. dactylidis and B. graminicola. In an examined specimen on Festuca heterophylla Lam. from Germany (GLM-F59650), the primary mycelium remains white, suggesting an identity of B. graminicola. This observation is supported by results of sequence analyses of collections from Germany on Festuca gigantea (BPI 562974) and USA on F. idahoensis Elmer (BPI 562965) that belong to B. graminicola on Bgt-1929, Bgt-4572, and CHS1 tress. Several collections of B. graminis s. lat. from Germany, Sachsen-Anhalt, on Alopecurus myosuroides Huds. (GLM-F48858, 50092, 54764, 57282, 94380), the primary mycelium and the asexual morph remain whitish as in B. graminicola, agreeing with the grouping of HAL 000022 F (on A. geniculatus) in B. graminicola clade on Bgt-1929 tree. In addition, specimens on Alopecurus could also belong to B. hordei (see BPI 562851 on Bgt-4572 tree).

Blumeria hordei M. Liu & Hambl., sp. nov.                               Fig. 9A–I.

MycoBank no.: MB 835997.

= Erysiphe graminis f. hordei-culti Jacz., Karm. Opred. Grib., Vip. 2. Muchn.-rosj. griby (Leningrad): 150, 1927.

= Erysiphe graminis f. hordei-spontanei Jacz., Karm. Opred. Grib., Vip. 2. Muchn.-rosj. griby (Leningrad): 151, 1927.

[Erysiphe graminis f. sp. hordei E. Marchal Compt. Rend. Acad. Sci. (Paris) 135, 211, 1902.]

Fig. 9 - Blumeria hordei sp. nov. (A–F DAOM 18154, G–I GLM-F99595 and DAOM 18154). A: Chasmothecia embedded in greyish to brownish secondary mycelia, developed simultaneously with deeper colored primary mycelia and conidia (arrow, DAOM 18154); B: Close-upof brownish conidial chains; C: Clavate or ovoid asci with wiggly stalks; D:Two conidiophores arising from single mother cell; E: Conidiophores arising singly from mother cells; F: Secondary hyphae arising from primary hyphae withone septum at the base; G: Conidia, mostly with round ends compared withpointed ends (i.e. B. graminicola Fig. 8 I); H: Nipple shaped appressoria mostly in pairs; I: SEM close-up of apical wall of conidia. Bars: A 0.5 mm; B 100 μm; C–H 20 μm; I 5 µm.

   Diagnosis: Development of primary mycelia and asexual morph from spring to summer, occasionally up to Sep, not inhibited with the onset of the formation of secondary mycelium and chasmothecia; primary mycelia, conidiophores and conidia turning greyish brown with age in summer; secondary hyphae aseptate; hyphal appressoria nipple-shaped, mostly in opposite pairs; one or two conidiophores per mother cells.

Type: CANADA, Quebec, Stat. expér. fédérale, Ste-Anne-de-la-Pocatière, on Hordeum vulgare, 8 Aug 1940, SAP 372 (holotype, DAOM 18154).

Gene sequences ex-holotype: MT622276 (ITS), MT633825 (Bgt-1929), MT650018 (Bgt-4572).

Exsiccatae: On Hordeum spp. – Barthol., Fungi Columb. 3427, 4726; Flora Olteniae Exs. 118, 535; Griff., West Amer. Fungi 165; Kellerm. & Swingle, Kansas Fungi 1133; Linh., Fungi Hung. 80; P. Sacc., Mycoth. Ital. 1473; Săvul., Herb. Mycol. Rom. 89, 1873, 1874, 3205; Thüm., Herb. Mycol. Oecon. 252.

Mycelia on stalks and leaves, amphigenous, thin to dense, effuse or in patches; primary mycelium white, becoming pigmented with age (in summer), greyish yellow (4B4) to greyish brown (5D3), not inhibited by the onset of secondary mycelia in late spring or early summer (mostly in May to Jun); secondary mycelia dingy greyish white to grey, with age sometimes faintly pigmented to greyish orange (5B2–5B3; Fig. 9A, B). Primary hyphal cells 3–6 µm wide; hyphal appressoria nipple-shaped, 3–6 µm wide, mostly in opposite pairs (Fig. 9H); secondary hyphae mostly aseptate (one septum near the base), thick-walled, wall 1–2 µm thick; central lumen narrow, 0–2 µm (Fig. 9F). Conidiophores, single or occasionally in pairs, erect, 60–120 × 5–7 µm, foot-cells 25–45 × 5–7 µm, bulbous swelling 10–15 µm wide, basal septum at the junction with the supporting hypha or slightly elevated up to 5 µm, 5–7 µm wide at the septum; 2–3 shorter cells, 10–25(–30) µm long (Fig. 9D, E);conidia broad ellipsoid, oblong, dolliform, ovoid, rarely limoniform, 20–32 × 11–15 µm (herbarium material), 23–38 × 12–18 µm (fresh material), length/width ratio 1.5–2.6(–2.9), hila 4–6 µm wide (Fig. 9G); apical walls with or without thickened patch, depressed in centre (Fig. 9I). Chasmothecia 125–185 µm diam when immature, 170–285 µm diam when mature;asci broad ellipsoid, ovoid, with short stalks (Fig. 9C); ascospores not observed.

Host range and distribution: POACEAE tribe TRITICEAE subtribe Hordinae, Hordeum spp., including H. murinum L., H. vulgare, occasional samples on Leymus condensatus (J. Presl) Á. Löve, Pseudoroegneria spicata (Pursh) Á. Löve; tribe POEAE subtribe Agrostidinae, Agrostis exarata Trin., Alopecurus aequalis Sobol.; tribe BROMEAE, Bromus spp. Africa: Afghanistan, Angola, Canary Islands, Egypt, Ethiopia, Jordan, Kenya, Lebanon, Libya, Morocco, Mozambique, Saudi Arabia, South Africa, Sudan, West Sahara; Asia: China, India, Iran, Iraq, Israel, Japan, Kazakhstan, Korea, Kyrgyzstan, Nepal, Pakistan, Russia – Siberia, Turkey, Turkmenistan, Uzbekistan, Yemen; Australia; Caucasus: Armenia, Azerbaijan, Georgia; Europe: throughout the whole continent; New Zealand; North America: Canada, Mexico, USA; South America: Argentina, Brazil, Chile, Colombia, Ecuador, Uruguay.

Notes: The clade corresponding to this species included samples mainly on Hordeum murinum L.and H. vulgare, but also sporadic samples on other genera. The identification of the host was as recorded on the specimen label, not confirmed by DNA analyses, therefore, the possibility that the host was misidentified cannot be ruled out. Reversely, not all samples on Hordeum spp. belong to B. hordei, they could belong to B. americana, B. graminis s. str. as well as B. dactylidis. The high level genetic variation supports the recognition of five special forms in barley powdery mildew (B. graminis f. sp. hordei) by Mains and Dietz (1930).

3.4. Key to Blumeria spp. based on morphology and host preference

1a. Primary mycelium and asexual morph remaining whitish to greyish white, at most somewhat yellowish or pale ochraceous with age, but never becoming brown …… 2

1b. Primary mycelium soon or at least with age becoming pigmented, yellowish, ochraceous to finally brown …… 4

2a. Hitherto only known on Bromus catharticus and Bromus sp. from Asia (Japan) and North America (USA); only primary mycelium and asexual morph known; conidia relatively long, 28–43 × 12–17.5 µm, with a length/width ratio of 1.7–3.2 ……Blumeria bromi-cathartici

2b. On other hosts; forming primary and secondary mycelium; conidia shorter, (20–)24–35(–40) µm long, length/width ratio 1.6–2.5(–3.1) …… 3

3a. Formation of the primary mycelium and asexual morph from spring (April/May) to autumn (September to November), sometimes even in winter; on Apera, Milium, and Poa spp., occasionally on additional hosts……Blumeria graminicola

3b. Formation of the primary mycelium from spring to summer, occasionally September, sometimes remaining light colored; on Hordeum spp.……Blumeria hordei

4a. Bristle-like curved-falcate hyphae of the secondary mycelium remaining colorless; on Dactylis spp. (occasionally also on other hosts, such as Alopecurus aequalis, Anthoxanthum odoratum, and Lolium multiflorum) ……Blumeria dactylidis

4b. Bristle-like curved-falcate hyphae of the secondary mycelium usually pigmented with age, yellowish, ochraceous, golden brown to brownish …… Complex of morphologically indistinguishable species, in part with overlapping hosts ranges (identification with certainty only possible by means of sequence analyses):

Mainly on Avena spp. ……Blumeria avenae

Mainly on Bromus spp.……Blumeria bulbigera

Mainly on Hordeum spp., occasionally on Agrostis, Alopecurus, Bromus, Leymus, Pseudoroegneria, Triticum……Blumeria hordei

Mainly on host belonging to Poaceae tribe Triticeae, occasionally on Brachypodium, Milium, Phleum……Blumeria graminis s. str.

Mainly on hosts in tribe Triticeae, including Aegilops,Elymus, Hordeum, Pascopyrum,Psathyrostachys, occasionally on Apera……Blumeria americana

3.5. Formae incertae sedis

The following names were introduced as mere host/substrate formae, in almost all cases without description. These formae are nevertheless valid names (the different hosts, which are eponymous, are considered to be sufficient as diagnostic information for formae in plant pathogenic fungi). However, the clarification of the affiliation of these formae requires typifications of these names and corresponding ex-type sequence data. Several of the host genera concerned belong to the host range of more than one Blumeria species.

Erysiphe graminis f. agrostidis Jacz., Karm. Opred. Grib., Vip. 2. Muchn.-rosj. griby (Leningrad): 143, 1927.

Note: Two samples on Agrostis sp. were included in analyses. One from Canada (DAOM 150568) was determined as B. graminicola, another from USA (BPI 859594) as B. hordei.

Erysiphe graminis f. beckmanniae Jacz. (l.c.: 144).

Note: two samples on Beckmannia spp. from Canada (DAOM 152932, DAOM 217878) grouped in B. graminicola.

Erysiphe graminis f. brizae Jacz. (l.c.: 490).

Erysiphe graminis f. bromi-brachypodii Jacz. (l.c.: 145).

Erysiphe graminis f. cynosuri Jacz. (l.c.: 149).

Erysiphe graminis f. deschampsiae Jacz. (l.c.).

Note: The sequences of a sample on Deschampsia from Finland (DAOM 82542) did not belong to any of the described species.

Erysiphe graminis f. gaudiniae Jacz. (l.c.: 490).

Erysiphe graminis f. holci Jacz. (l.c.: 150).

Note:One sample on Holcus lanatus from Germany (BPI 562984) belonged to B. graminicola on Bgt-4572 tree, another from Romania (BPI 562983) had no affinity to any described species.

Erysiphe graminis f. lepturi Jacz. (l.c.).

Erysiphe graminis f. moliniae Jacz. (l.c.).

Erysiphe graminis f. phalaridis Jacz. (l.c.).

Erysiphe graminis f. phlei Jacz. (l.c.).

Note: two samples on Phleum spp. from Russia (BPI 563065, HAL 000006 F) grouped in B. dactylidis on the Bgt-4572 and Bgt-1929 trees, respectively; DAOM 169330 from Canada in B. graminis s.str.

Erysiphe graminis f. sacchari Jacz. (l.c.: 153).

Erysiphe graminis f. sesleriae Jacz. (l.c.: 154).

Erysiphe graminis f. setariae Jacz. (l.c.).

Erysiphe graminis f. atropidis Lavrov, Trudy Tomsk. Gosud. Univ. Kuibysheva, Ser. Biol., 110(4): 193, 1951.

Erysiphe graminis f. cleistogenis Bunkina, Novosti Sist. Nizsh. Rast 10: 81, 1973.

Erysiphe graminis f. stipae Bunkina & Nelen, in Bunkina, Komarovskie Chteniya (Vladivostok) 21: 88, 1974.

Erysiphe graminis f. diarrhenae Bunkina, Novosti Sist. Nizsh. Rast. 1967: 175, 1967.

3.6.   Unresolved names

Acrosporium monilioides Nees, Syst. Pilze: 53, 1817.

≡ Oidium monilioides (Nees) Link, Sp. pl. 4, 6(1): 122, 1824.

≡ Oidium monilioides var. album Link, in Willdenow, Sp. pl. 4, 6(1): 122, 1824.

≡ Torula acrosporium Corda, in Sturm, Deutschl. Flora, III. Abt. Die Pilze Deutschlands, 8. Heft: 75, Nürnberg 1829, nom. illeg. (nom. superfl., Art. 52.1).

Type: on Dinebra retroflexa (Vahl) Panz., on a potted plant (not yet located, probably not preserved).

Notes: Dinebra belongs to Poaceae, tribe Cynodonteae subtribe Elsininae. The description of A. monilioides was based on an infected potted exotic grass. The infection could be by one of the described Blumeria species through host jumping.

Monilia hyalina Fr., Observ. mycol. 1: 210, 1815. Lectotype (designated here, MycoBank no.: MBT392802): Fries, Observ. mycol. 1: Tab. III, fig. 4 (a–d).

≡ Acrosporium hyalinum (Fr.) Sumst. [as ‘hyalina’], Mycologia 5(2): 58, 1913.

Type: “in culmis gramineis” (not preserved).

Note: Fries (l.c.) published a brief description and illustration, and cited Acharius as the collector. Herbarium Acharius housed in the University of Helsinki (H) was searched, but the original material of M. hyalina could not be located (O. Miettinen, pers. comm.). Neither could Uppsala University Museum of Evolution (UPS) locate the type material. Due to the paucity of the information in his original description, the host identity is unknown and it is unclear whether Fries found this fungus on living or dead grass stems. Link (1824) was the first author who reduced M. hyalina to synonymy with O. monilioides, and Sumstine (1913) reallocated the former name to Acrosporium and considered it the asexual morph of E. graminis. Braun and Cook (2012) followed this treatment and cited M. hyalina as synonym of B. graminis. However, this synonymy is doubtful. The ecology of M. hyalina is also unclear. It would also be possible that Fries (l.c.) observed and illustrated a saprobic hyphomycete on dead stems of grasses. His description (“articulis subgloboso-ovatis”) and illustration are not quite in concordance with the characters of the asexual morph of B. graminis, which is characterized by having ellipsoid-ovoid to limoniform conidia (never subglobose ones). The original illustration published by Fries (l.c.) is part of the original material and the only element available for lectotypification. However, as long as type material of M. hyalina is not available and cannot be re-examined, the status and the affinity of this name remain elusive and prevent its use.

Torula papillata Bonord., Bot. Zeitung (Leipzig) 19: 195, 1861.

≡ Oidium papillatum (Bonord.) Sacc. & Voglino, in Sacc., Syll. Fung. 4: 46, 1886. Lectotype (designated here, MycoBank no.: MBT392803): Bonorden, Bot. Zeitung (Leipzig) 19: Taf. VIII, Fig. 10 (a–e), 1861.

Notes: Bonorden (1861) described and illustrated Torula papillata with conidiophores attenuated towards an unswollen base, giving rise to conidia with papilloid apex and base (“utrimque subpapillatis”), formed singly or in chains. These characters are unusual and not in accordance with B. graminis s. lat. Bonorden (l.c.) was a good observer and provided exact drawings, as can be seen in his excellent drawing of the asexual morph of B. graminis s. lat. published as Oidium bulbigerum. Thus, it can be ruled out that he had overlooked existing swellings of the conidiophores in the case of T. papillata. Bonorden (l.c.) did not give any further details as to the host and ecology of this species. It is even possible that he had dealt with a saprobic hyphomycete on dead grass. The generic affinity of this species is quite unclear, although it was reallocated to Oidium by Saccardo and Voglino (in Saccardo, 1886) and accepted as a synonym of B. graminis (≡ Erysiphe graminis) in Braun (1987) and Braun and Cook (2012). Type material of T. papillata is not preserved, so that Bonorden’s original drawing represents the only original material available for lectotypification. Another collection, preferably new material with a culture and ex-culture sequence data, is necessary for epitypification to elucidate the generic affinity of this species.

Oidium monilioides var. flavicans Link, in Willdenow, Sp. pl. 4, 6(1): 123, 1824.

Type: Germany, Berlin (“in foliis Graminum in Germania. Lect. Berolini”), J.H.F. Link (not preserved).

Notes: Type material is not preserved in Berlin (herb. B, Robert Lücking, curator for cryptogams, pers. comm.), and Link (l.c.) did not give any details as to the host range. The mycelium of several Blumeria species turns yellowish, ochraceous to brownish with age.

4.   Discussion

Powdery mildew infections on cereal crops and grasses (Poaceae) have been treated as belonging to a single species since de Candolle’s (1815) first introduction of the name Erysiphe graminis. Subsequent landmark monographs of powdery mildews (Erysiphaceae) maintained this concept (Salmon, 1900; Braun & Cook, 2012) despite the perception of morphological differences between collections on various hosts (Blumer, 1967) and recognition of biological races (formae speciales) within E. graminis (e.g., Marchal, 1902; Oku et al., 1985). Multi-gene phylogenetic analyses of B. graminis (s. lat.) on a relatively broad sampling of hosts (Inuma et al., 2007) revealed nine lineages of B. graminis, and this motivated our present phylogenetic-taxonomic revision. The multi-gene analyses in this expanded study, for an even wider range of host plants and geographic origins, provided additional evidence of the cryptic speciation noted by Inuma et al. (2007) and for the formal recognition of multiple lineages at the species level within B. graminis s. lat.

Supplemented with morphological examinations of numerous specimens deposited in various herbaria under Erysiphe or Blumeria graminis, a summary of host and geographic ranges, and the re-evaluation of synonyms, this study is the first comprehensive taxonomic treatment of Blumeria complex.

MLSA has been commonly used for classifications in many groups of fungi (Taylor et al., 2000; Dettman, Jacobson, & Taylor, 2003). Yet, species delimitation of powdery mildew fungi still heavily depends only on rDNA, partly because the primers developed for other house-keeping genes cannot consistently amplify targeted loci (Braun & Cook, 2012). The challenge becomes more severe with historical herbarium samples, in which genomic DNA might have degraded due to long-term storage or inappropriate handling. During our study, besides the primers listed earlier (see Material and Methods), we tested a number of primers for other house-keeping genes, i.e. elongation factor 1 (TEF1) and beta tubulin (TUB2) to amplify DNA from herbarium samples (Inuma et al., 2007; TEF1 primers were newly designed), but the results were discouraging. Even for the ITS region, only 29% of samples were successfully sequenced, while CHS1 had a slightly higher success rate at 38%, possibly because the shorter amplicons (317 bps) were more easily amplified. To explore more potential loci for species delimitation, we investigated the 31 whole genome sequences available in GenBank, specifically the alignment of 93 phylogenetic informative genes (Menardo et al., 2017). Among our newly designed primers, two sets for two loci worked significantly better than others, i.e. 88% for Bgt-4572 and 63% for Bgt-1929. A BLAST search with the 226 bp amplicon of Bgt-4572 resulted in 81% match with E3 ubiquitin-protein ligase (XM 032013958) in V. echinocandica. E3 ubiquitin-protein ligase is involved in the control of various cellular processes in eukaryotic cells, including plant immune responses (Marino, Peeters, & Rivas, 2012; Duplan & Rivas, 2014). Recent studies have discovered the presence of E3 ubiquitin ligases in many fungal species and their conserved evolution (Marín, 2018). Our data showed that the DNA sequences provided species level resolution albeit a short amplicon (only 226 bp), which can be advantageous for amplifying DNA from herbarium specimens. Given its common presence in many lineages, we see great potential in the application of this gene region for species identification and detection of more fungal groups.

Whether or not cereal powdery mildews have co-evolved with their host plants has been debated in the literature. Some studies demonstrated these fungi might have co-evolved with their hosts (Matsuda & Takamatsu, 2003; Oberhaensli et al., 2011), while other evidence provided no support of this hypothesis (Wyand & Brown, 2003; Inuma et al., 2007). The phylogenetic tree based on concatenated sequences in our study reflected a certain level of co-evolution between Blumeria species and their principal hosts. For instance, the basal location of B.bulbigera (Fig. 1) and followed by the divergence of B. graminicola with B. bromi-cathartici mirrored the derived phylogenetic position of Triticeae and Poeae in relation to Bromeae (Soreng et al., 2017). Similarly, the derived position of B. avenae and B. dactylidis, both having Poeae as principal host, reflected the derived status of Poeae in relation to Triticeae (Fig. 1;Soreng et al., 2017). It is worth noting that the trees based on individual genes did not resolve deep relationships among species with statistical supports (Supplementary Figs. 1,2,3,4). However, besides the principal hosts (see above), almost all species may occur on additional hosts, either of the same or other tribes. A remarkable example is B. graminicola that showed a wide range of hosts including occasional infections on Bromus and Melica (Meliceae), suggesting both host expansion and host jumping may have played roles in increasing host ranges. Collectively, our results suggested it could be the combination of fungus-host co-evolution, host expansion, and host jumping that has shaped the diversity of Blumeria spp. and their host ranges.

In the light of the phylogenetic relationships among Blumeria spp. and the assumption of co-evolution with their principal hosts, the origins of certain species could be inferred. A possible Eurasian origin of B. bulbigera could be implied because most of the common species on Bromus are Eurasian (Kellogg, 2015). The wide distribution outside Eurasia can probably be explained by a co-distribution of B. bulbigera along with the wide synanthropic occurrence of several Bromus species as neophytes. For powdery mildews on cereal crops, i.e B. graminis s.str. on wheat (Triticum)and rye (Secale), B. hordei on barley (Hordeum), and B. avenae on oats (Avena), it was hypothesized previously that the Middle East was the centre of origin, based on investigations of phenotypic and genotypic diversities (Eshed & Wahl, 1970; Troch et al., 2012). Our analyses of the concatenated data matrix showed B. americana in an ancestral position relative to the cereal powdery mildews (Fig. 1), suggesting the divergence of powdery mildew of cereal crops from B. americana. However, the branch grouping B. avenae, B. dactylidis, B. graminis s. str. and B. hordei has low statistical support, indicating a genetic radiation, i.e., each species diverged independently, which could be a Eurasian origin from the more basal lineages, B. bulbigera.

The present phylogenetic-taxonomic revision of Blumeria was based on a broad collection of numerous host genera and species from various regions of the world. Nevertheless, these analyses were just a beginning. Host range and distribution of most Blumeria species are still so far only fragmentarily known. The number of previously recorded hosts and countries was very high (Amano, 1986; Braun & Cook, 2012), but in most cases not confirmed through molecular methods. The individual phylogenetic trees suggest the involvement of several additional, as yet unresolved lineages that probably represent additional species. Sequence analyses based on an even wider range of hosts and geographical origins are urgently needed.

Declaration of conflict interests

none

Declaration of contributions

All authors have participated in the research and article preparation, and agreed on publishing the present work. The project was conceived and designed by ML, SH, UB; and PS, KRB, SH, SK, ML developed molecular data; UB, SK, ML performed morphological examinations and taxonomy; UB handled nomenclature; KH scanning electron-microscopy; ML and UB drafted the manuscript, others contributed to editing.

Acknowledgements

We appreciate the constructive criticisms and suggestions for improvement made by two anonymous reviewers to an earlier version of this manuscript. We thank the Molecular Technologies Laboratory (MTL), and Electron-Imaging Lab at the Ottawa Research & Development Centre for technical assistance; herbaria U. S. National Fungus Collections (BPI), Canadian National Mycological Herbarium (DAOM), Conservatoire et Jardin botaniques de la Ville de Genève (G), Senckenberg Museum für Naturkunde Görlitz (GLM), Martin-Luther-Universität, Institut für Biologie, Bereich Geobotanik und Botanischer Garten Herbarium (HAL), and the National Museum of Nature and Science (Tokyo, TNS) for providing specimens. Special thanks to Dr. Lisa A. Castlebury and Shannon Dominick for facilitating the sampling of specimens in BPI by ML, SH, and PS. The study was funded by Agriculture and Agri-Food Canada STB fungal and bacterial biosystematics J-002272, development of molecular data for DAOM specimens was supported in part by funding from the Genomics Research and Development Initiative (GRDI, Project ID 2679) of the Government of Canada.

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