Phylogenetic studies uncover a predominantly African lineage in a widely distributed lichen-forming fungal species

A number of lichen-forming fungal species are widely distributed. Here, we investigate biogeographic patterns in a widely distributed isidiate taxon – Parmelinella wallichiana – using molecular sequence data. Our results revealed that Parmelinella wallichina, as currently circumscribed, is not monophyletic but falls into four clades, two of them represented by a sample only. A third clade, occurring in Africa and southern India is described as a new species, Parmelinella schimperiana Kirika & Divakar, sp. nov. Our study adds a further example of previously overlooked, geographically distinct, lineages that were discovered using molecular data.

Parmelinella is a small genus (ca. 10 species) and belongs to the parmelioid clade in the family Parmeliaceae (Divakar et al. 2015). The species included in this genus are characterized by a pored epicortex, isolichenan in the cell walls, subirregular lobes, cylindrical or bifusiform conidia, simple cilia and rhizines, and a yellow-grey upper cortex -containing secalonic acid derivatives and atranorin (Elix 1993;Crespo et al. 2010;Thell et al. 2012). Species in the genus are mainly distributed in subtropical to tropical regions of Africa, Asia, Australasia and South America. Parmelinella chozoubae, P. manipurensis and P. nimandairana are restricted to Asia; P. salacinifera, is reported from Southeast USA, central and south America, and Thailand; P. simplicior occurs in Asia and East Africa; and P. cinerascens, P. lindmanii, P. mutata and P. versiformis are endemic to South America (Elisaro et al. 2010;Benatti 2014). For a long time only four additional Parmelinella species were known from India (Divakar and Upreti 2005), but recent studies added six species to the genus, most of which had previously been known to occur only in South America (Elisaro et al. 2010;Benatti 2014). Of the ten species, only two, P. simplicior and P. wallichiana, have previously been reported from East Africa (Swinscow and Krog 1988; Alstrup et al. 2010).
Parmelinella wallichiana is the only widely distributed species in this genus and is known from Africa, Asia, Australia and South America. While it is widespread in East Africa and Asia, the species is known from a few localities in Australia and South America. Parmelinella wallichiana normally reproduces asexually by isidia and grows in wide range of ecological environments. The species is most frequently epiphytic but also found rarely on rocks. Studies have demonstrated broad, intercontinental distributions of a number of lichen-forming fungi that reproduce via asexual propagules (see e.g. Divakar et al. 2005;Molina et al. 2011a and b;Leavitt et al. 2013a;Roca-Valiente et al. 2013;. This study aims to assess biogeographic patterns in the widely distributed, isidiate, lichen-forming fungal species Parmelinella wallichiana. To this end, we generated DNA sequences of nuclear ribosomal internal transcribed spacer region (ITS1, 5.8S and ITS2), large subunit (nuLSU) and mitochondrial small subunit (mtSSU). Phenotypical features were re-evaluated and compared in light of the relationships inferred from the phylogenetic reconstructions.

Taxon sampling
A DNA data matrix was assembled using sequences of nuclear ITS, nuLSU and mitochondrial SSU rDNA of 21 samples, representing 18 specimens of P. wallichiana s. lat. from Africa, Asia and S. America assembled together with DNA sequences of P. aff. wallichiana and P. lindmanii (Elisaro et al. 2010) downloaded from GenBank. GenBank accession numbers and information of studied materials are shown in Table  1. The data sets include 12 sequences from previous publications (Blanco et al. 2004;Divakar et al. 2004;Divakar et al. 2006;Divakar et al. 2010b: Eliasaro et al. 2010: Kirika et al. 2015, and 25 were newly generated for this study. Three specimens of Bulbothrix isidiza were used as an out-group since it has been shown to belong to a sister group in a previous study (Kirika et. al.2015).

DNA extraction and PCR amplification
Total genomic DNA was extracted from small pieces of thallus devoid of any visible damage or contamination using the USB PrepEase Genomic DNA Isolation Kit (USB, Cleveland, OH) in accordance with the manufacturer's instructions. We generated sequence data from nuclear ribosomal markers, the ITS region and a fragment of the nuLSU, in addition to a fragment of the mtSSU. Polymerase-chain-reaction (PCR) amplifications were performed using Ready-To-Go PCR Beads (GE Healthcare, Pittsburgh, PA, USA) using the dilutions of total DNA. Fungal ITS rDNA was amplified using ITS1F primers (Gardes and Bruns 1993), ITS4 and ITS4A Larena et al. 1999); mtSSU rDNA was amplified using the primers mrSSU1, mrSSU3R and mrSSU2R (Zoller et al. 1999); nuLSU rDNA was amplified using LR0R and LR5 (Vilgalys and Hester 1990). PCR products were visualized on 1% agarose gel and cleaned using ExoSAP-IT (USB, Cleveland, OH, USA). Cycle sequencing of complementary strands was performed using BigDye v3.1 (Applied Biosystems,

Sequence editing and alignment
New sequences were assembled and edited using GENEIOUS v8.1.7 (Biomatters Ltd. 2005. Multiple sequence alignments for each locus were performed using the program MAFFT v7 Katoh and Toh 2008). For the ITS and nuLSU sequences, we used the G-INS-i alignment algorithm and '20PAM / K=2' scoring matrix, with an offset value of 0.3, and the remaining parameters were set to default values. We used the E-INS-i alignment algorithm and '20PAM / K=2' scoring matrix, with the remaining parameters were set to default values for the mtSSU sequences. The program Gblocks v0.91b (Talavera and Castresana 2007) was used to delimit and remove ambiguous alignment nucleotide positions from the final alignments using the online web server (http://molevol.cmima.csic.es/castresana/Gblocks_server. html), implementing the options for a less stringent selection of ambiguous nucleotide positions, including the 'Allow smaller final blocks', 'Allow gap positions within the final blocks', and 'Allow less strict flanking positions' options.

Phylogenetic analyses
Phylogenetic relationships were inferred using maximum likelihood (ML), and Bayesian inference (BI). Exploratory phylogenetic analyses of individual gene topologies showed no evidence of well-supported (≥ 70% bootstrap values) topological conflict, thus relationships were estimated from a concatenated, three-locus (ITS, nuLSU, mtSSU) data matrix using a total-evidence approach (Wiens 1998). We used the program RAxML v8.1.11 (Stamatakis 2006;Stamatakis et al. 2008) to reconstruct the concatenated ML gene-tree using the CIPRES Science Gateway server (http://www. phylo.org/portal2/). We implemented the 'GTRGAMMA' model, with locus-specific model partitions treating all loci as separate partitions, and evaluated nodal support using 1000 bootstrap pseudoreplicates. Exploratory analyses using alternative partitioning schemes resulted in identical topologies and highly similar bootstrap support values. We also reconstructed phylogenetic relationships from the concatenated multilocus data matrix under BI using the program BEAST v1.8.2 (Drummond and Rambaut 2007). We ran two independent Markov Chain Monte Carlo (MCMC) chains for 20 million generations, implementing a relaxed lognormal clock, a birth-death speciation process prior. The most appropriate model of DNA sequence evolution was selected for each marker using the program PartitionFinder v1.1.1 (Lanfear et al. 2012), treating the ITS1, 5.8S, ITS2, nuLSU, and mtSSU as separate partitions. The first 2 million generations were discarded as burn-in. Chain mixing and convergence were evaluated in Tracer v1.5 (Rambaut and Drummond 2009), considering ESS values >200 as a good indicator. Posterior trees from the two independent runs were combined using the program LogCombiner v1.8.0 (Drummond et al. 2012), and the final maximum clade credibility (MCC) tree was estimated from the combined posterior distribution of trees.

Morphological and chemical studies
Morphological characters, including lobe shape, size and width, cilia and rhizines were studied using a Leica Wild M 8 dissecting microscope. All the specimens of P. wallichiana included in the molecular analysis were evaluated (see Table 1). In the case of the new species, additional herbarium specimens were also studied.
Observations and measurements of ascospores were made in water, at 40× (objective) and 10× (eye piece) magnification with a Leica Leitz DM RB microscope. For each species at least 20 spores from different specimens were measured. Mean value (M) and standard deviation (SD) were calculated. In the description of the new species, the results of the measurements are given as (minimum value observed) M ± SD (maximum value observed). M, SD and n (number of spores measured) are expressed within parentheses. Chemical constituents were identified by thin layer chromatography using standard methods (Orange et al. 2010). Extraction of secondary metabolites for TLC analysis was done by pacing small pieces of the thallus in Eppendorf tubes and then adding a few drops of acetone in the tube. The resulting extract was then spotted on glass plates coated with Silica gel using capillary tubes. Plates were developed in Camag horizontal developing chamber (Oleico Lab Stockholm) using solvent system A (Toluene:Dioxane:acetic acid, 45:15:2), plates were then air dried, sprayed with 10% sulphuric acid and then heated in an oven at 110 degrees Celsius to visualize the spots. Substances were identified by comparing the spots with controls (Orange et al. 2010).

Results and discussion
A total of 28 new DNA sequences of Parmelinella wallichiana were generated for this study (Table 1). These were deposited in GenBank under accession numbers KX341978-KX342008. The dataset included samples from wide geographic regions as Asia, East Africa and South America. The final alignment of the combined data set was 2174 positions in length and was comprised of 458 unambiguously aligned nucleotide position characters in ITS, 844 in the nuLSU, and 872 in the mtSSU. As the topologies of the single locus phylogenies did not show any conflicts they were analyzed in a concatenated data matix (data not shown). The ML and BI analyses were identical in their topology and hence only the ML tree with support values of both analyses is depicted in Figure 1. Specimens representing Parmelinella wallichiana did not form a monophyletic lineage (Fig. 1). This is inconsistent with currently hypothesized species boundaries based on phenotypical features (Divakar and Upreti 2015;Benatti 2014). Species-level polyphylies are commonly found in Parmeliaceae and other groups of lichen-forming fungi (see reviews by Crespo and Lumbsch 2010;Lumbsch and Leavitt 2011).
Specimens representing P. wallichiana s. lat. fell into four distinct well-supported clades. Clade 'A' included samples from Kenya, Cameroon, and a single sample from South India. Clade 'B' included a single sample from coastal region (Coast Province) of Kenya. Clade 'C' included most samples from Asia; and clade 'D' was represented by a single sample from South America (Brazil). Specimens in clade 'A' are characterized in having smaller ascospores (5-10 × 5-7.5 µm), whereas they are larger (15-20 × 9-14 µm) in clade 'C'. Further, the same strongly supported monophyletic clades -'A' and 'C' -were recovered in reciprocally monophyletic clades in the independent gene trees (data not shown) (Hudson and Coyne 2002). Presence of the same clades in different single-locus genealogies can be taken as strong evidence that the clades are reproductively and evolutionarily isolated lineages representing distinct species-level lineages (Dettman et al. 2003;Pringle et al. 2005;de Quieroz 2007). The relationships among the clades were well supported ( Fig. 1). Clade 'B' formed sister-group relationship with clade 'A', whereas clade 'D' was sister to P. lindmanii, and clade 'C' sister to a clade including clade 'D' and P. lindmanii. The type material of Parmelinella wallichiana is from Nepal in the Himalayas (Hale 1976a;Divakar and Upreti 2005). Since all samples sequenced by us from the Himalayan regions (China and India) clustered in clade 'C', we consider this clade as P. wallichiana s. str.
For clade 'A' there are a few potential names available that we studied. For example, Parmelia junodi was described from the Cape Province in South Africa (Steiner 1907) and Parmelia tiliacea var. eximia has been described from Tanzania (Steiner 1888). These taxa have previously been considered synonyms of P. wallichiana (Hale 1976a). However, according to a recent study by Benatti (2014), Parmelia tiliacea var. eximia is a synonym of Parmelinella cinerascens and the type material of Parmelia junodi contained mixture of different species, such as Parmelinopsis minarum or P. horrescens and a fragment to belonged Parmelinella cinerascens. Thus we conclude that those two names are synonyms of Parmelinella cinerascens. The latter is a rare species occurring in South America and until recently was classified in the genus Canoparmelia (Elix et al. 1986). Recently, based on morphological data, Canoparmelia cinerascens was transferred to the genus Parmelinella (see Benatti 2014). Unfortunately, we were unable to sequence this species and hence cannot confirm the phylogenetic position of C. cinerascens. Samples clustered in clade 'A' collected from Africa and South India are morphologically similar to Parmelinella wallichiana s. lat. Since there is no name available for this clade, a new species is described below to accommodate samples from Africa and South India (clade 'A'). Further, the segregation of this new taxon from P. wallichiana s.str. is corroborated by morphological data, discussed below. The new species has a disjunct distribution occurring in Africa and South India. There are abundant examples of this disjunct distribution pattern in flowering plants (see e.g. Mani 1974;Kadereit 2004).
Clades 'B' and 'D' were each represented by a single specimen from Kenya and Brazil, respectively. The sample from the coastal region of Kenya (clade 'B') has a deviating morphology, i.e. very narrow, sublinear and dichotomous lobes, although the specimen from coastal Brazil (clade 'D') was more similar to P. wallichiana s. lat. In both cases, study of additional samples will be required before a formal description of these putative species.

Distribution and ecology.
At present the new species is known from Kenya, Cameroon and South India. It occurs in montane regions and in dry woodland areas. It is predominantly corticolous and sometimes saxicolous rarely terricolous, found corticolous on Mangifera indica, Juniperus procera, Podocarpus spp., Lannaea spp. and on Eucalyptus in artificial habitats.

Oomycete-specific its primers for identification and metabarcoding
Key words oomycete, community barcoding, next generation sequencing, ITS, community analysis, soil community introduction Oomycetes are microscopic stramenopiles that are found in both aquatic and terrestrial environments (Sparrow 1960, 1976, Karling 1981, Dick 2001. Many oomycete species are important pathogens, causing serious economic losses by infecting vegetables, berries, trees, arthropods and vertebrate animals (Kamoun 2003, Herrero et al. 2011. Molecular methods enable rapid identification of pathogens in environmental samples and infected tissues by using specific PCR primers and rapidly evolving high-throughput sequencing (HTS) technologies. For oomycetes, the cytochrome c oxidase subunit 1 (cox1), the internal transcribed spacer (ITS) (Robideau et al. 2011, Vettraino et al. 2012) and the cytochrome c oxidase subunit 2 (cox2) (Choi et al. 2015) have been identified as suitable barcodes. The choice of metabarcoding primers that cover all known oomycete taxa and discriminate other groups, however, is still limited by the inconsistent performance of some existing oomycete-specific primers. Robideau et al. (2011) evaluated the cox1, the ITS and the large ribosomal subunit (LSU) for use in DNA barcoding of oomycetes and suggested using cox1 and ITS in parallel due to their similar performance in resolving oomycete species and with both having superior performance in certain groups. Choi et al. (2015) compared the perfomance of cox1 and cox2 and found the latter to be more easily amplified across a wide range of oomycetes with existing primer sets. They also determined that the cox2 was more efficiently amplified from historic herbarium specimens and noted that in case of cox2 there is existing sequence data for several historic type specimens. As a result, Choi et al. (2015) suggested using the cox2 in addition to the ITS for oomycete barcoding. Additionally, Choi et al. (2015) proposed that for below species-level resolution the cox2-1 spacer could be used. For the cox2, there are also internal primers that can be used to amplify a 350 bp fragment suitable for barcoding (Hudspeth et al. 2000).
Of oomycete-specific ITS primers, ITS6 and ITS7 (Cooke et al. 2000) have been used for community studies, but with notable difficulties, as Coince et al. (2013) recovered only a small percentage of oomycete sequences using these primers. Sapkota and Nicolaisen (2015) optimized the ITS6/ITS7 assay by raising the annealing temperature and as a result improved the specificity of the primers. Other studies, however, have suggested that taxon recovery could be increased by using lower annealing temperatures (Ishii andFukui 2001, Acinas et al. 2005) or multiple annealing temperatures (Schmidt et al. 2013). It is also advisable to re-optimize the PCR reaction whenever the reaction mixture is altered (Innis et al. 1990).
Another oomycete-specific ITS primer, the ITS-O, has been published by Bachofer (2004). Whereas this primer has so far not found use in oomycete community studies, it has been used successfully to amplify the DNA of a wide range of oomycetes in phylogenetic research (Spring et al. 2006, Thines 2007.
The aim of the current study was to develop new oomycete ITS primers with improved taxon coverage and specificity for use in community-level studies. In order to reduce material costs, we decided to develop two oomycete specific forward primers that can be combined with various universal reverse primers. The new and existing primers were analyzed in silico to evaluate the coverage and specificity of the primers and the primers selected as suitable for oomycete ITS barcoding were tested in vitro on cultures, infected plant tissues and soil samples.

Methods and materials
Pure cultures of oomycetes and fungi DNA extracts from the pure cultures of eleven oomycete species from six genera were used in testing of the primers. Additionally, DNA from the cultures of five fungal species was used to test the specificity of the primers (Table 1).

Sampling and DNA extraction
A total of 20 soil samples were collected from beds of forest nurseries and bordering control areas (Table 2). Each sample consisted of 40 subsamples, which were taken table 1. Cultures of oomycetes and fungi that were used in testing the specificity of the new primers.

Species
Isolation with a 5 cm diameter sterile plastic pipe from the top 5 cm soil layer of a 50x50 m plot. The subsamples were pooled, dried and thoroughly mixed following Tedersoo et al. (2014). In addition, six samples of plant tissues with signs of oomycete infection were collected by excising a part of the symptomatic tissues (Table 3). DNA was isolated from 2 g of soil with the MO BIO PowerMax Soil DNA Isolation Kit (MO BIO Laboratories, Inc., Carlsbad, CA, USA). DNA from symptomatic The new forward primers were optimized for use with the universal reverse primer ITS4 ) using OLIGOANALYZER 3.1 (https://eu.idtdna.com/calc/ analyzer) to compare their calculated melting temperatures and GC content. The stability of possible homo-and heterodimers as well as hairpin structures was evaluated to avoid reduced amplification efficiency. All amplification tests were done at a 55 °C annealing temperature. Validation tests were carried out with DNA from various oomycete pure cultures as well as with DNA extracted from plants supposedly infected with pathogenic oomycetes and from 20 soil samples (Table 1). DNA from the pure cultures of five fungal species was used as negative control to test the specificity of the primers. The quality of all DNA samples used in specificity checks was tested by running PCR amplifications with universal ITS primers ITS1 and ITS4 ).

Sequencing of infected plant samples
PCR products obtained from the six symptomatic plant samples were purified using to the ExoSAP method (Bell 2008) and Sanger-sequenced in Macrogen (Macrogen Europe, Amsterdam, The Netherlands). Sequencing was done with the oomycete-specific ITS1oo or ITS3oo primers as well as with the universal ITS4 primer. The obtained sequences were compared against the INSDc to confirm the identification.

High-throughput sequencing of soil samples
In total, 20 soil samples were sequenced using Illumina Miseq 2x300 PE HTS technology in the Estonian Biocentre (Tartu, Estonia). Amplicons were prepared with the primers ITS1oo and ITS4ngs (Tedersoo et al. 2014), both of which were tagged with one of the MID identifiers (cf. Tedersoo et al. 2014). The ITS1oo was used as the forward primer in order to sequence both ITS1 and ITS2 regions. PCR was performed as described above but in four replicates. PCR products were pooled and 5 µl of each product was resolved on 1% agarose gel to confirm amplification. Negative controls without template and positive controls containing DNA of Aphanomyces astaci were used in the sequencing process. The quantity of the products was normalized with the SequalPrep Normalization Plate Kit (Invitrogen, Carlsbad, CA).

Analysis of Illumina sequencing data
Based on sequencing primers, read 1 and read 2 were shuffled to contain regions of ITS1 and ITS2, respectively (using FQGREP (https://github.com/indraniel/fqgrep)). These paired-end reads were analysed separately, because in most cases the amplified full-length ITS region exceeded 600 bp and could not be merged. Sequencing reads were quality filtered and assigned to samples using MOTHUR (Schloss et al. 2009) (average quality over 15 bases ≥ 30). Potential chimeras were detected and removed using USEARCH 7.0.1090 (Edgar 2010). Sequences shorter than 150 bases were discarded and longer sequences were trimmed to 150 bases for clustering. The quality filtered ITS1 and ITS2 sequences were separately clustered to Operational Taxonomic Units (OTUs) based on 97% sequence similarity using CD-HIT (Li and Godzik 2006). The most abundant sequence was selected as a representative (using mothur) for BLASTn searches against a custom oomycete nucleotide database combined from the reference collections of Hyde et al. (2014) and Robideau et al. (2011) and INSDc. For each OTU, 10 best-matching references were determined for precise annotation. We considered OTUs to belong to oomycetes if they best matched known oomycetes. Oomycete OTUs with e-values < e -20 and identities above 80% were considered reliable enough to assign sequences to an order. OTUs with best matches other than oomycetes were assigned at the class level if e-value was < e -20 and identity above 75%.

Primer selection and in silico analyses
As a result of aligning all oomycete ITS sequences present in the INSDc, it was possible to choose two short regions which are conserved across the majority of oomycetes and allow for the discrimination of other taxonomic groups. The primer ITS1oo overlaps with the primer ITS-O (Bachofer 2004) across 17 positions out of a total of 18 and is therefore not an original primer but a modification of ITS-O. This modification comes from a one bp shift which results in the deletion of a cytosine at the 5' end and the addition of an adenine at 3' end. The position of the added 3' adenine is polymorphic in other groups such as fungi and plants and should therefore make the modified IT-S1oo more specific than the original ITS-O (Bachofer 2004). Primer coverage analysis of ITS1oo and ITS3oo shows that the primer sequences are conserved in nearly all known oomycete taxa. In case of ITS3oo, mismatches can be seen in some accessions of Hyaloperonospora and Perofascia lepidii. Both primers have significant mismatches in comparison to most other stramenopiles, fungi and plants (Figure 1).
The location of the 18 bp long ITS1oo, modified from the ITS-O (Bachofer 2004), covers 13 nucleotides at end of the ribosomal 18S gene and 5 nucleotides in the beginning of ITS1. The similarly 18 bp long ITS3oo is located at the end of the 5.8S gene, ending 7 nucleotides before the beginning of ITS2 (Figure 1). Both of the new primers were used as forward primers in combination with the universal reverse primer ITS4. The ITS4 was chosen due to its position at the beginning of the 28S gene, which allows for the amplification of both ITS1 and/or ITS2 when used together with ITS1oo or ITS3oo.

Analyses of pure culture and infected plant material
The primer pairs ITS1oo/ITS4 and ITS3oo/ITS4 produced a single amplification band of the expected length from all 15 tested oomycete strains, representing six genera (Achlya, Aphanomyces, Phytophthora, Pythium, Saprolegnia, Scoliolegnia) and eleven species. No visible bands were obtained in gel with DNA from five fungal species.
• Four samples, extracted directly from the symptomatic tissues of a grey alder (Alnus incana), a potato (Solanum tuberosum), a tomato (Solanum lycopersicum) and a goutweed (Aegopodium podagraria), produced a single amplification band with both primer pairs ITS1oo/ITS4 and ITS3oo/ITS4 and were sequenced. Sequencing of the grey alder sample was successful with the primer ITS3oo, whereas the other three samples were successfully sequenced with both ITS1oo and ITS3oo. Comparisons against the NCBI GenBank nucleotide database showed that the sequence from the first sample belongs to Phytophthora sp. (99% similarity), the sequences from the second and third samples belong to Phytophthora infestans (100% and 99% similarity) and the sequence from the goutweed sample belongs to Plasmopara nivea (99% similarity). One sample from a zucchini plant (Cucurbita pepo) and one from a grape vine (Vitis vinifera) produced multiple amplification bands of different sizes with both primer pairs and were not sequenced.

Soil sample oomycete diversity
Altogether 67133 quality filtered ITS1 reads were recovered from the 20 soil samples. In all, 281 singletons were discarded from further analyses. Nearly 66% of all reads belonged to unknown taxa, 25% to oomycetes and 9% to other taxonomic groups ( Figure 2). The quality filtered ITS1 sequences were clustered into 1820 OTUs based on 97% similarity threshold, 30% of which were assigned to a known class or order. Out of the 554 assigned OTUs, nearly 73% belonged to oomycetes, 16% to fungi and 9% to plants. Of 404 oomycete OTUs, 307 were assigned to a known order. On average, oomycetes comprised 61 OTUs (range, 13-94) represented by 32% (range, 1-66%) of reads in soil samples (Figure 3). For the ITS2 subregion, 77734 quality filtered reads comprised 1720 OTUs and 241 singletons. Out of all ITS2 reads, 30% were assigned to oomycetes and 8% to fungi, whereas 60% belonged to unknown taxa (Figure 2). Oomycetes comprised 493 of the 672 identified taxa (73%). In total, 333 of these taxa were assigned to a known order. The number of oomycete OTUs averaged 86 (range, 42-148) per soil sample (Figure 3). On average, oomycetes contributed to 36% (range, 12-69%) in soil samples. 1 Oomycete read distribution between orders 2 Read distribution between classes, excluding reads of unknown origin 3 Read distribution between classes, including reads of unknown origin 4 OTU distribution between classes.

Discussion
The ultimate aim of this study was to validate an alternative method for metabarcoding oomycetes in complex substrates such as soil. We developed a novel taxon-specific PCR assay for the ITS region-based identification of oomycetes. When compared with the previously developed ITS-O, ITS6 and ITS7 primers, the ITS1oo, modified from the original ITS-O (Bachofer 2004), and the newly designed ITS3oo exhibit somewhat greater in silico specificity for oomycetes. In comparison to the ITS-O, the modified ITS1oo includes an additional 3' terminal adenine, a position that is polymorphic in fungi and plants and should therefor add to the specificity of the primer. Based on our analyses and in contrast to Sapkota and Nicolaisen (2015), the primer ITS6 has only one mismatching position in comparison to the majority of corresponding plant accessions in the INSDc, whereas the ITS1oo has several mismatches in the 3´ end. This may significantly lower the specificity of the ITS6, as a single internal mismatch does not reduce the amplification efficiency markedly (Kwok et al. 1990). In addition, the ITS6 has no mismatches compared to the majority of non-oomycete stramenopile accessions, while the ITS1oo has 3´ mismatches against several non-oomycete stramenopile groups. Both ITS1oo and ITS6 show complete coverage of all oomycete groups present in the INSDc, whereas the ITS7 has one mismatch in comparison to the accessions of Saprolegnia (Sapkota and Nicolaisen 2015) and Halophytophthora and 2-3 mismatches against four species of the known pathogenic genus Aphanomyces (Sapkota and Nicolaisen 2015). The presence of two or more mismatches can limit the usability of ITS7 in detecting these taxa, especially when using relatively high annealing temperatures (Sipos et al. 2007) as suggested by Sapkota and Nicolaisen (2015). In comparison, the primer ITS3oo has a single mismatch compared to the accessions of genus Hyaloperonospora and two mismatches against the single known species of Perofascia. Furthermore, the modified and newly developed forward primers are located in the very end of the conserved fragments that reduce the size of amplicons by 10-20% compared with the ITS6 forward primer, which is of great importance for HTS platforms producing short fragments such as Illumina and Ion Torrent. When combined with universal reverse primers, these oomycete-specific primers could be used in multiplex with other specific forward primers to address several taxonomic groups of pathogens simultaneously, without adding the cost of multiple barcoded reverse primers (Tedersoo et al. 2015). Previous studies have used oomycete-specific primers ITS6 and ITS7 to amplify the ITS1 region with highly variable success. For example, Vannini et al. (2013) recovered only 23 oomycete OTUs from 10 forest soil samples, where oomycetes contributed to 79% of all reads. More recently, Coince et al. (2013) recovered a total of 10 oomycete OTUs from 20 samples of forest soil that contributed to 15% of all reads. Sapkota and Nicolaisen (2015) improved the ITS6/ITS7 based method by optimizing the annealing temperature and as a result recovered 67 oomycete OTUs (95% of all reads) from 26 agricultural soil samples, but may have missed multiple taxa due to overly strict PCR conditions. Furthermore, it should be noted that fine tuning of PCR conditions is only possible in-house, because PCR buffer including salts (MgCl 2 ) and stabilizers (BSA), the type of polymerase and concentration of primers and templates all affect primer specificity (Innis et al. 1990, Cha andThilly 1993).
In this study, we recovered 404 ITS1-based and 493 ITS2-based oomycete OTUs from 20 soil samples from forest nurseries and bordering control areas. The number of recovered oomycete OTUs is considerably higher than in previous studies, which could be due to higher diversity in the analysed soil samples or a result of some properties of the new assay. Oomycete reads comprised on average 32% and 36% of the total reads of individual soil samples for ITS1 and ITS2, respectively. The assigned oomycete OTUs belonged to the orders of Lagenidiales, Peronosporales, Pythiales and Saprolegniales, confirming the ability of the proposed new assay to detect various oomycete groups from complex samples. Pythiales were found to be dominating in the soil samples, making up nearly 50% of the total oomycete reads, a result that is in line with previous oomycete community studies (Arcate et al. 2006, Sapkota andNicolaisen 2015).
The new primers were also used to identify oomycete pathogens from infected plant samples by using Sanger sequencing. The pathogens were successfully determined in four samples out of six. Sequencing was successful with both ITS1oo and ITS3oo from a goutweed (Aegopodium podagraria), a potato (Solanum tuberosum) and a tomato (Solanum lycopersicum) sample, whereas in the case of a grey alder (Alnus incana) sample only ITS3oo produced an identifiable sequence. This could indicate a somewhat higher specificity of ITS3oo in comparison to ITS1oo in some cases when identifying pathogens from infected plant material. Two samples out of six produced multiple amplification bands, possibly indicating the presence of several oomycete species in the infected sample. This result shows that the new primers can be used to detects oomycete pathogen species directly from infected plant samples in cases where the infected tissue in dominated by one pathogen, without co-amplification of plant and fungal DNA.
Taken together, we provide highly oomycete-specific forward primers that can be used in combination with previously developed oomycete-specific or universal reverse primers. Considering the rapid evolution of high-throughput sequencing, the full ITS sequence is certainly preferable over ITS1 or ITS2 used alone, because these subregions may differ in the taxonomic resolution across genera. Rhizoplaca melanophthalma (DC.) Leuckert & Poelt s.l. represents a group of morphologically distinct and chemically diverse species of lichen-forming fungi with broad ecological and geographical distributions. Species in this group occur all over the world in disjunct populations in continental climates, although species in this complex are notably absent from Australia. Rhizoplaca melanophthalma s.l. commonly grows on siliceous or calcareous rock in arid climates, but can also be found in montane coniferous forests, alpine tundra habitats, and bi-polar populations in the Arctic and Antarctica (McCune 1987). Members of this group are commonly used in air-quality biomonitoring research, making it an important species for conservation (Aslan et al. 2004;Dillman 1996). The species complex belongs to the recently re-circumscribed monophyletic genus Rhizoplaca in Lecanoraceae (Zhao et al. 2016).

Characterization of microsatellite markers in the cosmopolitan lichen-forming fungus
Previous multi-locus and phylogenomic studies support the circumscription of multiple species within R. melanophthalma s.l. (Leavitt et al. 2011(Leavitt et al. , 2013(Leavitt et al. , 2016b, many of which occur in sympatry in Western North America. In Western North America the distribution area of these species extends from the northern boreal zone to Mexico along the Rocky Mountains with a center of diversity in the Great Basin region (Leavitt et al. 2011). Rhizoplaca melanophthalma s. str. has the broadest ecological and geographic distribution of all known species within this complex, with populations occurring in desert, montane and steppe ecosystems in Antarctica, Central Asia, Europe, and North and South America (Leavitt et al. 2013).
The Rhizoplaca melanophthalma group provides an interesting system for assessing genetic diversity, population structure and gene flow in symbiotic fungal species with broad ecological and geographic distributions. To facilitate additional research into population-and landscape-level processes, 10 microsatellite markers were developed for R. melanophthalma s.str.

Materials and methods
A total of 42 specimens representing seven different species in the Rhizoplaca melanophthalma species complex were included in this study. Twenty-one of these represented R. melanophthalma s. str., three R. haydenii, four R. novomexicana, two R. parilis, four R. polymorpha, six R. porteri and two R. shushanii (Table 1). DNA was extracted from these specimens as described previously (Leavitt et al. 2011).
Fragment analysis was performed on an ABI 3730 DNA Analyzer (Applied Biosystems, Foster City, California, USA) using GeneScan-500 LIZ (Life Technologies, Warrington, UK) as an internal size standard. Genotyping was performed utilizing the microsatellite plugin in Geneious 9.1.2 (Biomatters Limited). Polymorphism within the microsatellites was tested in GenAlEx 6.5 (Peakall and Smouse 2012) by calculating Nei's unbiased genetic diversity.

Results and discussion
Of the 25 microsatellites assessed, 18 amplified successfully and 10 were polymorphic in all 21 R. melanophthalma s. str. specimens ( Table 2). The number of alleles per locus ranged from three to 11 with an average of 6.7. Nei's unbiased genetic diversity varied between 0.353 and 0.919 with the average genetic diversity being 0.765 (Table 3). The same 18 microsatellites that amplified successfully with R. melanophthalma s. str. also amplified in R. haydenii, R. novomexicana, R. parilis, R. polymorpha, R. porteri, and R. shushanii, but only three loci were polymorphic in all these species. For these three loci (Rmel1, Rmel, 4, and Rmel8) the number of alleles ranged from 11 to 15 with the average of 13 and Nei's unbiased genetic diversity varied between 0.892 and 0.905 with average of 0.900.
The 10 polymorphic microsatellite markers for the lichen-forming fungus R. melanophthalma will help elucidate population processes that have led to the observed distribution patterns in this widespread species.

the first its phylogeny of the genus Cantharocybe (Agaricales, hygrophoraceae) with a new record of C. virosa from Bangladesh introduction
The genus Cantharocybe was introduced by Bigelow and Smith in 1973 to accommodate Clitocybe gruberi Smith, based on the large yellow basidiomata, oblong to subcylindrical to elongated basidiospores and the presence of lageniform to lecythiform cheilocystidia. However, other taxa in this genus do not have large-sized yellow basidiomata and oblong to elongated basidiospores. Therefore, Lodge et al. (2014) extended the generic circumscription of the genus to include taxa with large, clitocyboid, yellow, dark brown to brownish gray basidiomata with long decurrent or adnate with decurrent tooth lamellae; abundant cheilocystidia which are usually lecythiform, sometimes with a mucronate apex, with or without a rounded capitulum; smooth, inamyloid, oblong, elongate, ellipsoid to broadly ellipsoid or rarely subglobose, basidiospores; a trichoderm or cutis pileipellis; and caulocystidia similar to cheilocystidia.  (Bigelow and Smith 1973, Justo et al. 2010, Esteves-Raventós et al. 2011, Ovrebo et al. 2011, Kumar and Manimohan 2013. Although C. gruberi was reported from China by Bi et al. (1993), the voucher specimen was re-identified as Oudemansiella bii Zhu L. Yang & Li F. Zhang (Yang and Zhang 2003). Recent molecular phylogenetic studies show that Cantharocybe is at the base of the hygrophoroid clade and is sister to Ampulloclitocybe (Pers.) Redhead, Lutzoni, Moncalvo & Vilgalys, but it is not clear if Cantharocybe and Cuphophyllus Donk (Bon) are members of Hygrophoraceae s.s. (Matheny et al. 2006, Binder et al. 2010). The phylogenetic relationships among the taxa of Cantharocybe are well resolved, based on partial nrLSU sequence analyses (Ovrebo et al. 2011, Kumar andManimohan 2013). Except for C. gruberi (non-holotype sequences from Esteves-Raventós et al. 2011), ITS sequences for other taxa of Cantharocybe are unavailable. Since the ITS region is an informative genetic region for species recognition in many groups of fungi (Schoch et al. 2012), we generated ITS sequences from holotype specimens of C. brunneovelutina and C. virosa. This is the first publication with ITS sequences from all holotype collections of Cantharocybe except C. gruberi, which we used to elucidate their phylogenetic relationships.
The first author has recently collected Cantharocybe material from tropical Bangladesh that is morphologically similar to C. virosa to some extent. Careful microscopic observation of the material from Bangladesh indicates that it could be conspecific with the Indian C. virosa, but the nrLSU sequence analysis suggested that it could be a new species or perhaps a variety of C. virosa. Therefore we attempted to obtain the holotype material of C. virosa from TENN in order to generate additional sequences and compare with the collection from Bangladesh. Fortunately, we received cloned ITS sequence of the holotype C. virosa (TENN 63483) from K.W. Hughes (Tennessee, USA) that we included in our further phylogenetic studies. The goal of this study is to elucidate the taxonomic position of the Bangladeshi collection of Cantharocybe, and clarify the confusion with an Indian collection of C. virosa using morphological and molecular evidence.

Collection and deposition
Cantharocybe specimens (Iqbal568 and 693) were collected from Madhupur upazila of Bangladesh on ground near to or associated with the roots of Cocos nucifera, a tree of the plant family Arecaceae during the monsoon (June to August) of 2012-2013. Specimens examined are deposited in the Cryptogamic Herbarium of Kunming Institute of Botany of the Chinese Academy of Sciences (KUN), China; and in the private herbarium of Iqbal (PHI). Cantharocybe brunneovelutina was previously deposited at BRH and CFMR (Ovrebo et al. 2011).

Morphological studies
The morphological description of the basidiomata is based on field notes and documented by photographs. Color codes are according to Kornerup and Wanscher (1978). A small fragment of dried specimen was revived in H 2 O, 5% KOH, and Congo red. The notation [n/m/p] is used in the descriptions of basidiospores measurements, which means n basidiospores from m basidiomata of p collections were measured; 20 basidiospores were measured from each voucher specimen. Dimension for basidiospores are given as (a-)bc(-d), in which 'b-c' contains a minimum of 90% of the measured values and extreme values 'a' and 'd' are given in parentheses. Q m = Q ± SD: Q indicates the length/width ratio of a measured basidiospore, Q m indicates to the average of Q basidiospores and SD is the standard deviation. For the pileipellis and stipitipellis observations radial-vertical section were made halfway of the pileus and stipe, respectively. Line drawings were done free hand.

Molecular studies
The protocol for DNA extraction followed that of Doyle and Doyle (1987). ITS1/ ITS4 or ITS1/ITS5 (White et al. 1090) and LROR/LR5 (Vilgalys and Hester 1990) primer pairs were used for the amplification of the internal transcribed spacer region (ITS) and the large subunit nuclear ribosomal RNA (nrLSU), respectively. PCR amplification was carried out following the protocol of Hosen et al. (2013). PCR confirmation was confirmed on 1% agarose electrophoresis gels stained with ethidium bromide. The amplified PCR products were sent to a commercial sequencing provider company (BGI, China) for sequencing.
Three sequences (nrLSU: KF303143 and KX452406, ITS: KX452403) were generated from the Bangladeshi Cantharocybe. Additionally, two ITS sequences were also obtained from the type materials of C. brunneovelutina (BZ-1883: KX452404) and C. virosa (TENN-63483: KX452405). ITS sequences generated for this study were cloned in pMD18-T following manufacturer's instructions. The newly generated sequences were deposited in GenBank. An initial BLASTn search of the nrLSU sequence obtained from the Bangladeshi material against the NCBI database (http:// www.ncbi.nlm.nih.gov/) gave C. virosa (=Megacollybia virosa Manim. & K.B. Vrinda), C. brunneovelutina and C. gruberi as closest hits, with maximum similarities of 96%, 96% and 95%, respectively. The closest nrLSU sequences including Ampulloclitocybe and Cuphophyllus were retrieved from GenBank and additional taxa were chosen after consulting Lodge et al. (2014) and then combined with nrLSU sequence from Bangladesh materials. Two additional datasets were constructed: ITS and ITS+nrLSU to clarify relationships between Indian and Bangladesh collections of Cantharocybe. All datasets were aligned with Mafft v.6.8  and manually adjusted with BioEdit v.7.0.9 (Hall 1999) using default settings. Maximum Likelihood (ML) and Bayesian Inference (BI) methods followed those in Hosen et al. (2013). Phyllotopsis nidulans (Pers.) Singer was served as outgroup for all dataset analyses as inferred from other phylogenetic studies (Ovrebo et al. 2011, Kumar and Manimohan 2013. Both ML and BI analyses were conducted using RAxML v.7.2.6 (Stamatakis 2006) and MrBayes v.3.1.2 (Ronquist and Huelsenbeck 2003) with default settings. For ML analyses, the General Time Reversible Model of evolution with estimated gamma distribution was selected, and statistical support values were obtained using nonparametric bootstrapping (BS) with 1000 replicates. For BI analysis, the substitution model suitable for ITS and nrLSU datasets were determined using the Akaike Information Criterion (AIC) implemented in MrModeltest v.2.3 (Nylander 2004), and the models were SYM+G and GTR+I+G, respectively. Bayesian Inference analyses were conducted using the selected evolutionary model with four chains and generations set to 0.5 million for the ITS and nrLSU datasets, and four million for the combined dataset (ITS+nrLSU). Runs were terminated once the average standard deviation of split frequencies went below 0.01. Trees were sampled every 100 generations, with the first 25% of trees discarded as burn-in. Posterior probabilities (PP) were calculated using the "sump" and "sumt" commands implemented in MrBayes.

Molecular results
Three datasets (ITS, nrLUS and ITS+nrLSU) are constructed and analysed separately. The ITS dataset includes 17 sequences of fungal taxa (Table 1)   disrupted replication unless the DNA was cloned using a vector such as the method we used to obtain our ITS sequences. As there is no notation in GenBank that the AFTOL ITS sequence of C. gruberi was cloned, we infer that forward reads were disrupted by the same intron that was found in C. brunneovelutina and C. virosa. Thus, it is likely that Matheny and Hibbett's AFTOL program only obtained back-reads from the 5' end using the ITS4 primer. The authors of the C. gruberi DQ200927 sequence may or may not have obtained a partial read of an intron, but if so, it would have been of diminishing quality with distance from the ITS4 primer, it would not have matched any known ITS sequences, and it would have been impossible to correct or corroborate without a forward read from the 3' end. We therefore infer that if a partial read of an intron was obtained by Matheny and Hibbett for C. gruberi, that it was trimmed from the GenBank submission because it did not match ITS1 sequences and it could not be corrected or corroborated. Blast searches of GenBank using the ITS sequence of C. virosa, after removing the intron, retrieved C. gruberi sequences with highest similarity.
Tree topologies obtained from both ML and BI methods of phylogenetic analyses are congruent, the ML trees are shown in Fig. 1. The Bangladesh sample of Cantharocybe clusters in a strongly supported clade with the Indian sample of C. virosa in all three datasets, indicating that they are conspecific (Fig. 1). Description. Basidiomata medium-sized to large. Pileus 50-80 mm diam., convex at first then applanate, sometimes uplifted with cracked margin, tawny gray, dark brown (6E4-5) to grayish brown (5E3-5E4, 6E3-6F4), dry, pruinose or with fine appressed scales under lens, margin without striation. Hymenophore lamellate; lamellae adnate to decurrent, subdistant to crowded, white to pallid white (5A1, 6A1); lamellulae numerous, concolorous with lamellae. Stipe 50-80 × 10-15 mm, central, slightly curved, cylindrical, gradually thickening towards the base, at the apex ribbed by the subdeccurent lines of the hymenophore, upper half pale gray or brownish gray (5D2) to grayish brown (5E3) pruina or squamules and the remaining half nearly concolorous with the pileus, with cottony mycelium at the base, interior solid, milky white to white. Context 12 mm thick in the center of the pileus, milky white to white (6A1), firm, solid, unchanging when cut or bruised.
Habitat. Solitary or in clusters, associated with roots of Cocos nucifera (collection Iqbal 568) or along the roadside on ground (collection Iqbal 693) near C. nucifera.

Morphology and phylogenetic relationships of Cantharocybe
The Bangladeshi C. virosa is characterized by its gray to grayish brown basidiomata, moderately crowded lamellae, fine squamules on stipe surface formed from clusters of lecythiform caulocystidia, ellipsoid to broadly ellipsoid basidiospores, and a trichoderm pileipellis.
Based on molecular analyses, C. virosa, a species recently described from India is conspecific with the Bangladeshi collection (Fig. 1). However, the Indian C. virosa has a pale grayish brown pileus, long cheilocystidia which can be up to 63 µm with a long neck up to 35 µm, a cutis pileipellis or occasionally disrupted with trichodermal patches (Kumar and Manimohan 2013). In comparison, the Bangladeshi collection has a grayish brown to dark brown pileus, cheilocystida with short neck up to 15 µm long, and clearly defined trichoderm pileipellis. Geographically, C. virosa is distributed in the Kerala state (South-West region) of India, while the new collection was collected from Tangail district of the Dhaka division of Bangladesh. Moreover, the nrLSU sequence obtained from the Bangladeshi collection does not perfectly match (96%) with the holotype C. virosa retrieved from GenBank, and the genetic distance between them is 0.96% (8 bases differences and 23 deletions in the Indian collection) of 831 nucleotide sites. These morphological variations, geographic distance and nrLSU sequence inferred suggest that they may have diverged recently from each other due to its allopatric speciation.
Surprisingly, when we blasted the newly generated ITS sequences from the holotypes of C. virosa (TENN 63483) and C. brunneovelutina (BZ-1883) individually against the NCBI database, we did not find C. gruberi as the closest sister species among the first 100 matched species, even 80-88% matched only with some taxa of Tricholoma, Lepista, Macrolepiota, Lepiota, etc. The ITS sequence from the newly collected material from Bangladesh also gave a similar result. These results are caused by the presence of a large intron in C. brunneovelutina and C. virosa, and the truncated sequence of C. gruberi deposited in GenBank (missing the intron and 3' end). Subsequently, we retrieved the closest ITS sequences of those taxa including C. gruberi from GenBank and reconstructed an ITS phylogenetic tree where all Cantharocybe taxa were nested together within the same clade with strong BS value (99% ML BS, 1.0 PP) and apart from Tricholoma and Lepista clades (data not shown). In the ITS sequence analyses, the Indian entity C. virosa showed only 5 base pair difference with 4 deletions out of 742 nucleotides (genetic distance 0.68%) from the Bangladeshi material. Fur-ther extended combined dataset (ITS+nrLSU) also showed little divergence (genetic distance 0.83%) between them (Fig. 1c). This small variation in ITS sequences, with negligible differences in the color of the basidioma, size of basidia and cheilocystidia which were possibly due to environmental variations, do not warrant a new variety or species, suggesting that both south Asian entities are conspecific. Furthermore, both collections were the same ecology and associated with the roots of C. nucifera in a tropical region. Neither coconut trees nor palms in general have been shown to associate with ectomycorrhizal fungi. Halbwachs et al. (2013) however, found hyphae of a Cuphophyllus species (a genus near Cantharocybe) and several species of Hygrocybe as endophytes in plant roots including those of a monocot (Plantago major) so another type of root symbiosis with C. nucifera is possible. The considerable variation in the nrLSU sequence (96% match, 8 bases differences with 23 deletion) of the Indian C. virosa may be explained by the fact that the nrLSU sequence obtained from that collection was not clean, showing evidence of a contaminating sequence or minor indel (K.W. Hughes, pers. comm.).
Molecular phylogenetic analyses indicated that the genus Cantharocybe is monophyletic, with strong bootstrap values (Fig. 1). Likewise, Ovrebo et al. (2011) and Lodge et al. (2014) showed that the monophyletic clade of Cantharocybe has strong BS value comprising C. gruberi and C. brunneovelutina using single locus or multi-locus sequence analyses. Although recent phylogenetic studies (Ovrebo et al. 2011, Kumar and Manimohan 2013 showed the monophyly of Cantharocybe, the sister relationship with other genera remains unclear. Cantharocybe nests at the base of the hygrophoroid clade together with Ampulloclitocybe and Cuphophyllus (Binder et al. 2010, Matheny et al. 2006, Ovrebo et al. 2011), but their relationships were not confidently resolved. In our combined dataset (ITS+nrLSU) analysis, Ampulloclitocybe is only weakly supported (65% ML BS, PP = 0.96) as sister to the Cuphophylloid clade (Fig. 1c). This is accordance with the recent phylogenetic study of Lodge et al. (2014).

Distribution and ecology of Cantharocybe
Cantharocybe is an uncommon genus that only consists of three species. Cantharocybe gruberi has wide distribution from America (New Mexico, western North America and British Columbia) to Europe (Spain). Cantharocybe brunneovelutina is reported from tropical Central America (Belize) whereas C. virosa is from tropical South Asia (Bangladesh and India). Based on the branching order with strong bootstrap support at all nodes in our phylogeny in which C. gruberi is basal, we infer that the genus Cantharocybe may have originated in America or Europe and then migrated independently to Central America and South Asia.
The south Asian species usually occurs with the roots of Cocos nucifera (Manimohan et al. 2010, Kumar andManimohan 2013, this study as well), the North American and European collections were found on needle beds or ground under conifers and Pinus nigra, respectively (Bigelow andSmith 1973, Esteves-Raventós et al. 2011), and the Central American species was found in humus around dead palm trees (Ovrebo et al. 2011). However, their symbiotic association with trees is still unknown. To facilitate identification of Cantharocybe taxa worldwide, a key to the species is given bellow. Basidiomata medium (up to 100 mm broad), pale grayish brown to tawny gray to grayish brown, basidiospores 6.5-11(-12) × 5.5-6.5(-7) µm, ellipsoid to broadly ellipsoid or rarely subglobose, Bangladesh and India (South Asia, tropical