2022 Volume 7
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Molecular phylogeny and morphology reveal a new wood-rotting fungal species, Cyathus wenshanensis sp. nov. from the Yunnan-Guizhou Plateau

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  • A new species of bird's nest fungus, Cyathus wenshanensis is proposed based on a combination of the morphological and molecular evidence. It is characterised by the obconical to cupulate basidiomata covered with hirsute hairs, striations on the outer and inner surface of the peridium, funicular peridioles, a trimitic hyphal system of peridium with generative hyphae having clamp connections, a dimitic hyphal system of peridiole middle, and subglobose, elliptical to ellipsoid-elongate, thick-walled basidiospores. Sequence of the internal transcribed spacers (ITS) gene region was generated, and the phylogenetic analysis was performed with maximum likelihood, maximum parsimony and Bayesian inference methods. The phylogenetic analyses inferred from ITS dataset indicated that C. wenshanensis nested within the genus Cyathus, in which it formed a monophyletic lineage and grouped with C. albinus, C. amazonicus, C. badius, C. parvocinereus, C. pyristriatus and C. uniperidiolus.
  • Aquaporins (AQPs) constitute a large family of transmembrane channel proteins that function as regulators of intracellular and intercellular water flow[1,2]. Since their first discovery in the 1990s, AQPs have been found not only in three domains of life, i.e., bacteria, eukaryotes, and archaea, but also in viruses[3,4]. Each AQP monomer is composed of an internal repeat of three transmembrane helices (i.e., TM1–TM6) as well as two half helixes that are formed by loop B (LB) and LE through dipping into the membrane[5]. The dual Asn-Pro-Ala (NPA) motifs that are located at the N-terminus of two half helixes act as a size barrier of the pore via creating an electrostatic repulsion of protons, whereas the so-called aromatic/arginine (ar/R) selectivity filter (i.e., H2, H5, LE1, and LE2) determines the substrate specificity by rendering the pore constriction site diverse in both size and hydrophobicity[59]. Based on sequence similarity, AQPs in higher plants could be divided into five subfamilies, i.e., plasma membrane intrinsic protein (PIP), tonoplast intrinsic protein (TIP), NOD26-like intrinsic protein (NIP), X intrinsic protein (XIP), and small basic intrinsic protein (SIP)[1017]. Among them, PIPs, which are typically localized in the cell membrane, are most conserved and play a central role in controlling plant water status[12,1822]. Among two phylogenetic groups present in the PIP subfamily, PIP1 possesses a relatively longer N-terminus and PIP2 features an extended C-terminus with one or more conserved S residues for phosphorylation modification[5,15,17].

    Tigernut (Cyperus esculentus L.), which belongs to the Cyperaceae family within Poales, is a novel and promising herbaceous C4 oil crop with wide adaptability, large biomass, and short life period[2327]. Tigernut is a unique species accumulating up to 35% oil in the underground tubers[2830], which are developed from stolons and the process includes three main stages, i.e., initiation, swelling, and maturation[3133]. Water is essential for tuber development and tuber moisture content maintains a relatively high level of approximately 85% until maturation when a significant drop to about 45% is observed[28,32]. Thereby, uncovering the mechanism of tuber water balance is of particular interest. Despite crucial roles of PIPs in the cell water balance, to date, their characterization in tigernut is still in the infancy[21]. The recently available genome and transcriptome datasets[31,33,34] provide an opportunity to address this issue.

    In this study, a global characterization of PIP genes was conducted in tigernut, including gene localizations, gene structures, sequence characteristics, and evolutionary patterns. Moreover, the correlation of CePIP mRNA/protein abundance with water content during tuber development as well as subcellular localizations were also investigated, which facilitated further elucidating the water balance mechanism in this special species.

    PIP genes reported in Arabidopsis (Arabidopsis thaliana)[10] and rice (Oryza sativa)[11] were respectively obtained from TAIR11 (www.arabidopsis.org) and RGAP7 (http://rice.uga.edu), and detailed information is shown in Supplemental Table S1. Their protein sequences were used as queries for tBLASTn[35] (E-value, 1e–10) search of the full-length tigernut transcriptome and genome sequences that were accessed from CNGBdb (https://db.cngb.org/search/assembly/CNA0051961)[31,34]. RNA sequencing (RNA-seq) reads that are available in NCBI (www.ncbi.nlm.nih.gov/sra) were also adopted for gene structure revision as described before[13], and presence of the conserved MIP (major intrinsic protein, Pfam accession number PF00230) domain in candidates was confirmed using MOTIF Search (www.genome.jp/tools/motif). To uncover the origin and evolution of CePIP genes, a similar approach was also employed to identify homologs from representative plant species, i.e., Carex cristatella (v1, Cyperaceae)[36], Rhynchospora breviuscula (v1, Cyperaceae)[37], and Juncus effusus (v1, Juncaceae)[37], whose genome sequences were accessed from NCBI (www.ncbi.nlm.nih.gov). Gene structure of candidates were displayed using GSDS 2.0 (http://gsds.gao-lab.org), whereas physiochemical parameters of deduced proteins were calculated using ProtParam (http://web.expasy.org/protparam). Subcellular localization prediction was conducted using WoLF PSORT (www.genscript.com/wolf-psort.html).

    Nucleotide and protein multiple sequence alignments were respectively conducted using ClustalW and MUSCLE implemented in MEGA6[38] with default parameters, and phylogenetic tree construction was carried out using MEGA6 with the maximum likelihood method and bootstrap of 1,000 replicates. Systematic names of PIP genes were assigned with two italic letters denoting the source organism and a progressive number based on sequence similarity. Conserved motifs were identified using MEME Suite 5.5.3 (https://meme-suite.org/tools/meme) with optimized parameters as follows: Any number of repetitions, maximum number of 15 motifs, and a width of 6 and 250 residues for each motif. TMs and conserved residues were identified using homology modeling and sequence alignment with the structure resolved spinach (Spinacia oleracea) SoPIP2;1[5].

    Synteny analysis was conducted using TBtools-II[39] as described previously[40], where the parameters were set as E-value of 1e-10 and BLAST hits of 5. Duplication modes were identified using the DupGen_finder pipeline[41], and Ks (synonymous substitution rate) and Ka (nonsynonymous substitution rate) of duplicate pairs were calculated using codeml in the PAML package[42]. Orthologs between different species were identified using InParanoid[43] and information from synteny analysis, and orthogroups (OGs) were assigned only when they were present in at least two species examined.

    Plant materials used for gene cloning, qRT-PCR analysis, and 4D-parallel reaction monitoring (4D-PRM)-based protein quantification were derived from a tigernut variety Reyan3[31], and plants were grown in a greenhouse as described previously[25]. For expression profiling during leaf development, three representative stages, i.e., young, mature, and senescing, were selected and the chlorophyll content was checked using SPAD-502Plus (Konica Minolta, Shanghai, China) as previously described[44]. Young and senescing leaves are yellow in appearance, and their chlorophyll contents are just half of that of mature leaves that are dark green. For diurnal fluctuation regulation, mature leaves were sampled every 4 h from the onset of light at 8 a.m. For gene regulation during tuber development, fresh tubers at 1, 5, 10, 15, 20, 25, and 35 d after tuber initiation (DAI) were collected as described previously[32]. All samples with three biological replicates were quickly frozen with liquid nitrogen and stored at −80 °C for further use. For subcellular localization analysis, tobacco (Nicotiana benthamiana) plants were grown as previously described[20].

    Tissue-specific expression profiles of CePIP genes were investigated using Illumina RNA-seq samples (150 bp paired-end reads) with three biological replicates for young leaf, mature leaf, sheath of mature leaf, shoot apex, root, rhizome, and three stages of developmental tuber (40, 85, and 120 d after sowing (DAS)), which are under the NCBI accession number of PRJNA703731. Raw sequence reads in the FASTQ format were obtained using fastq-dump, and quality control was performed using fastQC (www.bioinformatics.babraham.ac.uk/projects/fastqc). Read mapping was performed using HISAT2 (v2.2.1, https://daehwankimlab.github.io/hisat2), and relative gene expression level was presented as FPKM (fragments per kilobase of exon per million fragments mapped)[45].

    For qRT-PCR analysis, total RNA extraction and synthesis of the first-strand cDNA were conducted as previously described[24]. Primers used in this study are shown in Supplemental Table S2, where CeUCE2 and CeTIP41[25,33] were employed as two reference genes. PCR reaction in triplicate for each biological sample was carried out using the SYBR-green Mix (Takara) on a Real-time Thermal Cycler Type 5100 (Thermal Fisher Scientific Oy). Relative gene abundance was estimated with the 2−ΔΔCᴛ method and statistical analysis was performed using SPSS Statistics 20 as described previously[13].

    Raw proteomic data for tigernut roots, leaves, freshly harvested, dried, rehydrated for 48 h, and sprouted tubers were downloaded from ProteomeXchange/PRIDE (www.proteomexchange.org, PXD021894, PXD031123, and PXD035931), which were further analyzed using Maxquant (v1.6.15.0, www.maxquant.org). Three dominant members, i.e., CePIP1;1, -2;1, and -2;8, were selected for 4D-PRM quantification analysis, and related unique peptides are shown in Supplemental Table S3. Protein extraction, trypsin digestion, and LC-MS/MS analysis were conducted as described previously[46].

    For subcellular localization analysis, the coding region (CDS) of CePIP1;1, -2;1, and -2;8 were cloned into pNC-Cam1304-SubN via Nimble Cloning as described before[30]. Then, recombinant plasmids were introduced into Agrobacterium tumefaciens GV3101 with the helper plasmid pSoup-P19 and infiltration of 4-week-old tobacco leaves were performed as previously described[20]. For subcellular localization analysis, the plasma membrane marker HbPIP2;3-RFP[22] was co-transformed as a positive control. Fluorescence observation was conducted using confocal laser scanning microscopy imaging (Zeiss LMS880, Germany): The wavelength of laser-1 was set as 730 nm for RFP observation, where the fluorescence was excited at 561 nm; the wavelength of laser-2 was set as 750 nm for EGFP observation, where the fluorescence was excited at 488 nm; and the wavelength of laser-3 was set as 470 nm for chlorophyll autofluorescence observation, where the fluorescence was excited at 633 nm.

    As shown in Table 1, a total of 14 PIP genes were identified from eight tigernut scaffolds (Scfs). The CDS length varies from 831 to 882 bp, putatively encoding 276–293 amino acids (AA) with a molecular weight (MW) of 29.16–31.59 kilodalton (kDa). The theoretical isoelectric point (pI) varies from 7.04 to 9.46, implying that they are all alkaline. The grand average of hydropathicity (GRAVY) is between 0.344 and 0.577, and the aliphatic index (II) ranges from 94.57 to 106.90, which are consistent with the hydrophobic characteristic of AQPs[47]. As expected, like SoPIP2;1, all CePIPs include six TMs, two typical NPA motifs, the invariable ar/R filter F-H-T-R, five conserved Froger's positions Q/M-S-A-F-W, and two highly conserved residues corresponding to H193 and L197 in SoPIP2;1 that were proven to be involved in gating[5,48], though the H→F variation was found in CePIP2;9, -2;10, and -2;11 (Supplemental Fig. S1). Moreover, two S residues, corresponding to S115 and S274 in SoPIP2;1[5], respectively, were also found in the majority of CePIPs (Supplemental Fig. S1), implying their posttranslational regulation by phosphorylation.

    Table 1.  Fourteen PIP genes identified in C. esculentus.
    Gene name Locus Position Intron no. AA MW (kDa) pI GRAVY AI TM MIP
    CePIP1;1 CESC_15147 Scf9:2757378..2759502(–) 3 288 30.76 8.82 0.384 95.28 6 47..276
    CePIP1;2 CESC_04128 Scf4:3806361..3807726(–) 3 291 31.11 8.81 0.344 95.95 6 46..274
    CePIP1;3 CESC_15950 Scf54:5022493..5023820(+) 3 289 31.06 8.80 0.363 94.57 6 49..278
    CePIP2;1 CESC_15350 Scf9:879960..884243(+) 3 288 30.34 8.60 0.529 103.02 6 33..269
    CePIP2;2 CESC_00011 Scf30:4234620..4236549(+) 3 293 31.59 9.27 0.394 101.57 6 35..268
    CePIP2;3 CESC_00010 Scf30:4239406..4241658(+) 3 291 30.88 9.44 0.432 98.97 6 31..266
    CePIP2;4 CESC_05080 Scf46:307799..309544(+) 3 285 30.44 7.04 0.453 100.32 6 28..265
    CePIP2;5 CESC_05079 Scf46:312254..314388(+) 3 286 30.49 7.04 0.512 101.68 6 31..268
    CePIP2;6 CESC_05078 Scf46:316024..317780(+) 3 288 30.65 7.68 0.475 103.06 6 31..268
    CePIP2;7 CESC_05077 Scf46:320439..322184(+) 3 284 30.12 8.55 0.500 100.00 6 29..266
    CePIP2;8 CESC_14470 Scf2:4446409..4448999(+) 3 284 30.37 8.30 0.490 106.90 6 33..263
    CePIP2;9 CESC_02223 Scf1:2543928..2545778(–) 3 283 30.09 9.46 0.533 106.47 6 31..262
    CePIP2;10 CESC_10007 Scf27:1686032..1688010(–) 3 276 29.16 9.23 0.560 106.05 6 26..256
    CePIP2;11 CESC_10009 Scf27:1694196..1696175(–) 3 284 29.71 9.10 0.577 105.49 6 33..263
    AA: amino acid; AI: aliphatic index; GRAVY: grand average of hydropathicity; kDa: kilodalton; MIP: major intrinsic protein; MW: molecular weight; pI: isoelectric point; PIP: plasma membrane intrinsic protein; Scf: scaffold; TM: transmembrane helix.
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    To uncover the evolutionary relationships, an unrooted phylogenetic tree was constructed using the full-length protein sequences of CePIPs together with 11 OsPIPs and 13 AtPIPs. As shown in Fig. 1a, these proteins were clustered into two main groups, corresponding to PIP1 and PIP2 as previously defined[10,49], and each appears to have evolved into several subgroups. Compared with PIP1s, PIP2s possess a relatively shorter N-terminal but an extended C-terminal with one conserved S residue (Supplemental Fig. S1). Interestingly, a high number of gene repeats were detected, most of which seem to be species-specific, i.e., AtPIP1;1/-1;2/-1;3/-1;4/-1;5, AtPIP2;1/-2;2/-2;3/-2;4/-2;5/-2;6, AtPIP2;7/-2;8, OsPIP1;1/-1;2/-1;3, OsPIP2;1/-2;4/-2;5, OsPIP2;2/-2;3, CePIP1;1/-1;2, CePIP2;2/-2;3, CePIP2;4/-2;5/-2;6/-2;7, and CePIP2;9/-2;10/-2;11, reflecting the occurrence of more than one lineage-specific whole-genome duplications (WGDs) after their divergence[50,51]. In Arabidopsis that experienced three WGDs (i.e. γ, β, and α) after the split with the monocot clade[52], AtPIP1;5 in the PIP1 group first gave rise to AtPIP1;1 via the γ WGD shared by all core eudicots[50], which latter resulted in AtPIP1;3, -1;4, and -1;2 via β and α WGDs; AtPIP2;1 in the PIP2 group first gave rise to AtPIP2;6 via the γ WGD, and they latter generated AtPIP2;2, and -2;5 via the α WGD (Supplemental Table S1). In rice, which also experienced three WGDs (i.e. τ, σ, and ρ) after the split with the eudicot clade[51], OsPIP1;2 and -2;3 generated OsPIP1;1 and -2;2 via the Poaceae-specific ρ WGD, respectively. Additionally, tandem, proximal, transposed and dispersed duplications also played a role on the gene expansion in these two species (Supplemental Table S1).

    Figure 1.  Structural and phylogenetic analysis of PIPs in C. esculentus, O. sativa, and A. thaliana. (a) Shown is an unrooted phylogenetic tree resulting from full-length PIPs with MEGA6 (maximum likelihood method and bootstrap of 1,000 replicates), where the distance scale denotes the number of amino acid substitutions per site. (b) Shown are the exon-intron structures. (c) Shown is the distribution of conserved motifs among PIPs, where different motifs are represented by different color blocks as indicated and the same color block in different proteins indicates a certain motif. (At: A. thaliana; Ce: C. esculentus; PIP: plasma membrane intrinsic protein; Os: O. sativa).

    Analysis of gene structures revealed that all CePIP and AtPIP genes possess three introns and four exons in the CDS, in contrast to the frequent loss of certain introns in rice, including OsPIP1;2, -1;3, -2;1, -2;3, -2;4, -2;5, -2;6, -2;7, and -2;8 (Fig. 1b). The positions of three introns are highly conserved, which are located in sequences encoding LB (three residues before the first NPA), LD (one residue before the conserved L involved in gating), and LE (18 residues after the second NPA), respectively (Supplemental Fig. S1). The intron length of CePIP genes is highly variable, i.e., 109–993 bp, 115–1745 bp, and 95–866 bp for three introns, respectively. By contrast, the exon length is relatively less variable: Exons 2 and 3 are invariable with 296 bp and 141 bp, respectively, whereas Exons 1 and 4 are of 277–343 bp and 93–132 bp, determining the length of N- and C-terminus of PIP1 and PIP2, respectively (Fig. 1b). Correspondingly, their protein structures were shown to be highly conserved, and six (i.e., Motifs 1–6) out of 15 motifs identified are broadly present. Among them, Motif 3, -2, -6, -1, and -4 constitute the conserved MIP domain. In contrast to a single Motif 5 present in most PIP2s, all PIP1s possess two sequential copies of Motif 5, where the first one is located at the extended N-terminal. In CePIP2;3 and OsPIP2;7, Motif 5 is replaced by Motif 13; in CePIP2;2, it is replaced by two copies of Motif 15; and no significant motif was detected in this region of CePIP2;10. PIP1s and PIP2s usually feature Motif 9 and -7 at the C-terminal, respectively, though it is replaced by Motif 12 in CePIP2;6 and OsPIP2;8. PIP2s usually feature Motif 8 at the N-terminal, though it is replaced by Motif 14 in CePIP2;2 and -2;3 or replaced by Motif 11 in CePIP2;10 and -2;11 (Fig. 1c).

    As shown in Fig. 2a, gene localization of CePIPs revealed three gene clusters, i.e., CePIP2;2/-2;3 on Scf30, CePIP2;4/-2;5/-2;6/-2;7 on Scf46, and CePIP2;10/-2;11 on Scf27, which were defined as tandem repeats for their high sequence similarities and neighboring locations. The nucleotide identities of these duplicate pairs vary from 70.5% to 91.2%, and the Ks values range from 0.0971 to 1.2778 (Table 2), implying different time of their birth. According to intra-species synteny analysis, two duplicate pairs, i.e., CePIP1;1/-1;2 and CePIP2;2/-2;4, were shown to be located within syntenic blocks (Fig. 2b) and thus were defined as WGD repeats. Among them, CePIP1;1/-1;2 possess a comparable Ks value to CePIP2;2/-2;3, CePIP1;1/-1;3, and CePIP2;4/-2;8 (1.2522 vs 1.2287–1.2778), whereas CePIP2;2/-2;4 harbor a relatively higher Ks value of 1.5474 (Table 2), implying early origin or fast evolution of the latter. While CePIP1;1/-1;3 and CePIP2;1/-2;8 were characterized as transposed repeats, CePIP2;1/-2;2, CePIP2;9/-2;10, and CePIP2;8/-2;10 were characterized as dispersed repeats (Fig. 2a). The Ks values of three dispersed repeats vary from 0.8591 to 3.0117 (Table 2), implying distinct times of origin.

    Figure 2.  Duplication events of CePIP genes and synteny analysis within and between C. esculentus, O. sativa, and A. thaliana. (a) Duplication events detected in tigernut. Serial numbers are indicated at the top of each scaffold, and the scale is in Mb. Duplicate pairs identified in this study are connected using lines in different colors, i.e., tandem (shown in green), transposed (shown in purple), dispersed (shown in gold), and WGD (shown in red). (b) Synteny analysis within and between C. esculentus, O. sativa, and A. thaliana. (c) Synteny analysis within and between C. esculentus, C. cristatella, R. breviuscula, and J. effusus. Shown are PIP-encoding chromosomes/scaffolds and only syntenic blocks that contain PIP genes are marked, i.e., red and purple for intra- and inter-species, respectively. (At: A. thaliana; Cc: C. cristatella; Ce: C. esculentus; Je: J. effusus; Mb: megabase; PIP: plasma membrane intrinsic protein; Os: O. sativa; Rb: R. breviuscula; Scf: scaffold; WGD: whole-genome duplication).
    Table 2.  Sequence identity and evolutionary rate of homologous PIP gene pairs identified in C. esculentus. Ks and Ka were calculated using PAML.
    Duplicate 1 Duplicate 2 Identity (%) Ka Ks Ka/Ks
    CePIP1;1 CePIP1;3 78.70 0.0750 1.2287 0.0610
    CePIP1;2 CePIP1;1 77.20 0.0894 1.2522 0.0714
    CePIP2;1 CePIP2;4 74.90 0.0965 1.7009 0.0567
    CePIP2;3 CePIP2;2 70.50 0.1819 1.2778 0.1424
    CePIP2;4 CePIP2;2 66.50 0.2094 1.5474 0.1353
    CePIP2;5 CePIP2;4 87.30 0.0225 0.4948 0.0455
    CePIP2;6 CePIP2;5 84.90 0.0545 0.5820 0.0937
    CePIP2;7 CePIP2;6 78.70 0.0894 1.0269 0.0871
    CePIP2;8 CePIP2;4 72.90 0.1401 1.2641 0.1109
    CePIP2;9 CePIP2;10 76.40 0.1290 0.8591 0.1502
    CePIP2;10 CePIP2;8 64.90 0.2432 3.0117 0.0807
    CePIP2;11 CePIP2;10 91.20 0.0562 0.0971 0.5783
    Ce: C. esculentus; Ka: nonsynonymous substitution rate; Ks: synonymous substitution rate; PIP: plasma membrane intrinsic protein.
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    According to inter-species syntenic analysis, six out of 14 CePIP genes were shown to have syntelogs in rice, including 1:1, 1:2, and 2:2 (i.e. CePIP1;1 vs OsPIP1;3, CePIP1;3 vs OsPIP1;2/-1;1, CePIP2;1 vs OsPIP2;4, CePIP2;2/-2;4 vs OsPIP2;3/-2;2, and CePIP2;8 vs OsPIP2;6), in striking contrast to a single one found in Arabidopsis (i.e. CePIP1;2 vs AtPIP1;2). Correspondingly, only OsPIP1;2 in rice was shown to have syntelogs in Arabidopsis, i.e., AtPIP1;3 and -1;4 (Fig. 2b). These results are consistent with their taxonomic relationships that tigernut and rice are closely related[50,51], and also imply lineage-specific evolution after their divergence.

    As described above, phylogenetic and syntenic analyses showed that the last common ancestor of tigernut and rice is more likely to possess only two PIP1s and three PIP2s. However, it is not clear whether the gene expansion observed in tigernut is species-specific or Cyperaceae-specific. To address this issue, recently available genomes were used to identify PIP subfamily genes from C. cristatella, R. breviuscula, and J. effuses, resulting in 15, 13, and nine members, respectively. Interestingly, in contrast to a high number of tandem repeats found in Cyperaceae species, only one pair of tandem repeats (i.e., JePIP2;3 and -2;4) were identified in J. effusus, a close outgroup species to Cyperaceae in the Juncaceae family[36,37]. According to homologous analysis, a total of 12 orthogroups were identified, where JePIP genes belong to PIP1A (JePIP1;1), PIP1B (JePIP1;2), PIP1C (JePIP1;3), PIP2A (JePIP2;1), PIP2B (JePIP2;2), PIP2F (JePIP2;3 and -2;4), PIP2G (JePIP2;5), and PIP2H (JePIP2;6) (Table 3). Further intra-species syntenic analysis revealed that JePIP1;1/-1;2 and JePIP2;2/-2;3 are located within syntenic blocks, which is consistent with CePIP1;1/-1;2, CePIP2;2/-2;4, CcPIP1;1/-1;2, CcPIP2;3/-2;4, RbPIP1;1/-1;2, and RbPIP2;2/-2;5 (Fig. 2c), implying that PIP1A/PIP1B and PIP2B/PIP2D were derived from WGDs occurred sometime before Cyperaceae-Juncaceae divergence. After the split with Juncaceae, tandem duplications frequently occurred in Cyperaceae, where PIP2B/PIP2C and PIP2D/PIP2E/PIP2F retain in most Cyperaceae plants examined in this study. By contrast, species-specific expansion was also observed, i.e., CePIP2;4/-2;5, CePIP2;10/-2;11, CcPIP1;2/-1;3, CcPIP2;4/-2;5, CcPIP2;8/-2;9, CcPIP2;10/-2;11, RbPIP2;3/-2;4, and RbPIP2;9/-2;10 (Table 3 & Fig. 2c).

    Table 3.  Twelve proposed orthogroups based on comparison of representative plant species.
    Orthogroup C. esculentus C. cristatella R. breviuscula J. effusus O. sativa A. thaliana
    PIP1A CePIP1;1 CcPIP1;1 RbPIP1;1 JePIP1;1 OsPIP1;3 AtPIP1;1, AtPIP1;2,
    AtPIP1;3, AtPIP1;4,
    AtPIP1;5
    PIP1B CePIP1;2 CcPIP1;2, CcPIP1;3 RbPIP1;2 JePIP1;2
    PIP1C CePIP1;3 CcPIP1;4 RbPIP1;3 JePIP1;3 OsPIP1;1, OsPIP1;2
    PIP2A CePIP2;1 CcPIP2;1 RbPIP2;1 JePIP2;1 OsPIP2;1, OsPIP2;4,
    OsPIP2;5
    AtPIP2;1, AtPIP2;2,
    AtPIP2;3, AtPIP2;4,
    AtPIP2;5, AtPIP2;6
    PIP2B CePIP2;2 CcPIP2;2 RbPIP2;2 JePIP2;2 OsPIP2;2, OsPIP2;3
    PIP2C CePIP2;3 CcPIP2;3 RbPIP2;3, RbPIP2;4
    PIP2D CePIP2;4, CePIP2;5 CcPIP2;4, CcPIP2;5 RbPIP2;5
    PIP2E CePIP2;5 CcPIP2;5 RbPIP2;6
    PIP2F CePIP2;6 CcPIP2;6
    PIP2G CePIP2;7 CcPIP2;7 RbPIP2;7 JePIP2;3, JePIP2;4
    PIP2H CePIP2;8 CcPIP2;8, CcPIP2;9 RbPIP2;8 JePIP2;5 OsPIP2;6 AtPIP2;7, AtPIP2;8
    PIP2I CePIP2;9, CePIP2;10,
    CePIP2;11
    CcPIP2;10, CcPIP2;11 RbPIP2;9, RbPIP2;10 JePIP2;6 OsPIP2;7, OsPIP2;8
    At: A. thaliana; Cc: C. cristatella; Ce: C. esculentus; Je: J. effuses; Os: O. sativa; Rb: R. breviuscula; PIP: plasma membrane intrinsic protein.
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    Tissue-specific expression profiles of CePIP genes were investigated using transcriptome data available for young leaf, mature leaf, sheath, root, rhizome, shoot apex, and tuber. As shown in Fig. 3a, CePIP genes were mostly expressed in roots, followed by sheaths, moderately in tubers, young leaves, rhizomes, and mature leaves, and lowly in shoot apexes. In most tissues, CePIP1;1, -2;1, and -2;8 represent three dominant members that contributed more than 90% of total transcripts. By contrast, in rhizome, these three members occupied about 80% of total transcripts, which together with CePIP1;3 and -2;4 contributed up to 96%; in root, CePIP1;1, -1;3, -2;4, and -2;7 occupied about 84% of total transcripts, which together with CePIP2;1 and -2;8 contributed up to 94%. According to their expression patterns, CePIP genes could be divided into five main clusters: Cluster I includes CePIP1;1, -2;1, and -2;8 that were constitutively and highly expressed in all tissues examined; Cluster II includes CePIP2;2, -2;9, and -2;10 that were lowly expressed in all tested tissues; Cluster III includes CePIP1;2 and -2;11 that were preferentially expressed in young leaf and sheath; Cluster IV includes CePIP1;3 and -2;4 that were predominantly expressed in root and rhizome; and Cluster V includes remains that were typically expressed in root (Fig. 3a). Collectively, these results imply expression divergence of most duplicate pairs and three members (i.e. CePIP1;1, -2;1, and -2;8) have evolved to be constitutively co-expressed in most tissues.

    Figure 3.  Expression profiles of CePIP genes in various tissues, different stages of leaf development, and mature leaves of diurnal fluctuation. (a) Tissue-specific expression profiles of 14 CePIP genes. The heatmap was generated using the R package implemented with a row-based standardization. Color scale represents FPKM normalized log2 transformed counts, where blue indicates low expression and red indicates high expression. (b) Expression profiles of CePIP1;1, -2;1, and -2;8 at different stages of leaf development. (c) Expression profiles of CePIP1;1, -2;1, and -2;8 in mature leaves of diurnal fluctuation. Bars indicate SD (N = 3) and uppercase letters indicate difference significance tested following Duncan's one-way multiple-range post hoc ANOVA (p< 0.01). (Ce: C. esculentus; FPKM: Fragments per kilobase of exon per million fragments mapped; PIP: plasma membrane intrinsic protein)

    As shown in Fig. 3a, compared with young leaves, transcriptome profiling showed that CePIP1;2, -2;3, -2;7, -2;8, and -2;11 were significantly down-regulated in mature leaves, whereas CePIP1;3 and -2;1 were up-regulated. To confirm the results, three dominant members, i.e., CePIP1;1, -2;1, and -2;8, were selected for qRT-PCR analysis, which includes three representative stages, i.e., young, mature, and senescing leaves. As shown in Fig. 3b, in contrast to CePIP2;1 that exhibited a bell-like expression pattern peaking in mature leaves, transcripts of both CePIP1;1 and -2;8 gradually decreased during leaf development. These results were largely consistent with transcriptome profiling, and the only difference is that CePIP1;1 was significantly down-regulated in mature leaves relative to young leaves. However, this may be due to different experiment conditions used, i.e., greenhouse vs natural conditions.

    Diurnal fluctuation expression patterns of CePIP1;1, -2;1, and -2;8 were also investigated in mature leaves and results are shown in Fig. 3c. Generally, transcripts of all three genes in the day (8, 12, 16, and 20 h) were higher than that in the night (24 and 4 h). During the day, both CePIP1;1 and -2;8 exhibited an unimodal expression pattern that peaked at 12 h, whereas CePIP2;1 possessed two peaks (8 and 16 h) and their difference was not significant. Nevertheless, transcripts of all three genes at 20 h (onset of night) were significantly lower than those at 8 h (onset of day) as well as 12 h. In the night, except for CePIP2;1, no significant difference was observed between the two stages for both CePIP1;1 and -2;8. Moreover, their transcripts were comparable to those at 20 h (Fig. 3c).

    To reveal the expression patterns of CePIP genes during tuber development, three representative stages, i.e., 40 DAS (early swelling stage), 85 DAS (late swelling stage), and 120 DAS (mature stage), were first profiled using transcriptome data. As shown in Fig. 4a, except for rare expression of CePIP1;2, -2;2, -2;9, and -2;10, most genes exhibited a bell-like expression pattern peaking at 85 DAS, in contrast to a gradual decrease of CePIP2;3 and -2;8. Notably, except for CePIP2;4, other genes were expressed considerably lower at 120 DAS than that at 40 DAS. For qRT-PCR confirmation of CePIP1;1, -2;1, and -2;8, seven stages were examined, i.e., 1, 5, 10, 15, 20, 25, and 35 DAI, which represent initiation, five stages of swelling, and maturation as described before[32]. As shown in Fig. 4b, two peaks were observed for all three genes, though their patterns were different. As for CePIP1;1, compared with the initiation stage (1 DAI), significant up-regulation was observed at the early swelling stage (5 DAI), followed by a gradual decrease except for the appearance of the second peak at 20 DAI, which is something different from transcriptome profiling. As for CePIP2;1, a sudden drop of transcripts first appeared at 5 DAI, then gradually increased until 20 DAI, which was followed by a gradual decrease at two late stages. The pattern of CePIP2;8 is similar to -1;1, two peaks appeared at 5 and 20 DAI and the second peak was significantly lower than the first. The difference is that the second peak of CePIP2;8 was significantly lower than the initiation stage. By contrast, the second peak (20 DAI) of CePIP2;1 was significantly higher than that of the first one (1 DAI). Nevertheless, the expression patterns of both CePIP2;1 and -2;8 are highly consistent with transcriptome profiling.

    Figure 4.  Transcript and protein abundances of CePIP genes during tuber development. (a) Transcriptome-based expression profiling of 14 CePIP genes during tuber development. The heatmap was generated using the R package implemented with a row-based standardization. Color scale represents FPKM normalized log2 transformed counts, where blue indicates low expression and red indicates high expression. (b) qRT-PCR-based expression profiling of CePIP1;1, -2;1, and -2;8 in seven representative stages of tuber development. (c) Relative protein abundance of CePIP1;1, -2;1, and -2;8 in three representative stages of tuber development. Bars indicate SD (N = 3) and uppercase letters indicate difference significance tested following Duncan's one-way multiple-range post hoc ANOVA (p < 0.01). (Ce: C. esculentus; DAI: days after tuber initiation; DAS: days after sowing; FPKM: Fragments per kilobase of exon per million fragments mapped; PIP: plasma membrane intrinsic protein).

    Since protein abundance is not always in agreement with the transcript level, protein profiles of three dominant members (i.e. CePIP1;1, -2;1, and -2;8) during tuber development were further investigated. For this purpose, we first took advantage of available proteomic data to identify CePIP proteins, i.e., leaves, roots, and four stages of tubers (freshly harvested, dried, rehydrated for 48 h, and sprouted). As shown in Supplemental Fig. S2, all three proteins were identified in both leaves and roots, whereas CePIP1;1 and -2;8 were also identified in at least one of four tested stages of tubers. Notably, all three proteins were considerably more abundant in roots, implying their key roles in root water balance.

    To further uncover their profiles during tuber development, 4D-PRM-based protein quantification was conducted in three representative stages of tuber development, i.e., 1, 25, and 35 DAI. As expected, all three proteins were identified and quantified. In contrast to gradual decrease of CePIP2;8, both CePIP1;1 and -2;1 exhibited a bell-like pattern that peaked at 25 DAI, though no significant difference was observed between 1 and 25 DAI (Fig. 4c). The trends are largely in accordance with their transcription patterns, though the reverse trend was observed for CePIP2;1 at two early stages (Fig. 4b & Fig. 4c).

    As predicted by WoLF PSORT, CePIP1;1, -2;1, and -2;8 may function in the cell membrane. To confirm the result, subcellular localization vectors named pNC-Cam1304-CePIP1;1, pNC-Cam1304-CePIP2;1, and pNC-Cam1304-CePIP2;8 were further constructed. When transiently overexpressed in tobacco leaves, green fluorescence signals of all three constructs were confined to cell membranes, highly coinciding with red fluorescence signals of the plasma membrane marker HbPIP2;3-RFP (Fig. 5).

    Figure 5.  (a) Schematic diagram of overexpressing constructs, (b) subcellular localization analysis of CePIP1;1, -2;1, and -2;8 in N. benthamiana leaves. (35S: cauliflower mosaic virus 35S RNA promoter; Ce: C. esculentus; EGFP: enhanced green fluorescent protein; kb: kilobase; NOS: terminator of the nopaline synthase gene; RFP: red fluorescent protein; PIP: plasma membrane intrinsic protein).

    Water balance is particularly important for cell metabolism and enlargement, plant growth and development, and stress responses[2,19]. As the name suggests, AQPs raised considerable interest for their high permeability to water, and plasma membrane-localized PIPs were proven to play key roles in transmembrane water transport between cells[1,18]. The first PIP was discovered in human erythrocytes, which was named CHIP28 or AQP1, and its homolog in plants was first characterized in Arabidopsis, which is known as RD28, PIP2c, or AtPIP2;3[3,7,53]. Thus far, genome-wide identification of PIP genes have been reported in a high number of plant species, including two model plants Arabidopsis and rice[10,11,1317,5456]. By contrast, little information is available on Cyperaceae, the third largest family within the monocot clade that possesses more than 5,600 species[57].

    Given the crucial roles of water balance for tuber development and crop production, in this study, tigernut, a representative Cyperaceae plant producing high amounts of oil in underground tubers[28,30,32], was employed to study PIP genes. A number of 14 PIP genes representing two phylogenetic groups (i.e., PIP1 and PIP2) or 12 orthogroups (i.e., PIP1A, PIP1B, PIP1C, PIP2A, PIP2B, PIP2C, PIP2D, PIP2E, PIP2F, PIP2G, PIP2H, and PIP2I) were identified from the tigernut genome. Though the family amounts are comparative or less than 13–21 present in Arabidopsis, cassava (Manihot esculenta), rubber tree (Hevea brasiliensis), poplar (Populus trichocarpa), C. cristatella, R. breviuscula, banana (Musa acuminata), maize (Zea mays), sorghum (Sorghum bicolor), barley (Hordeum vulgare), and switchgrass (Panicum virgatum), they are relatively more than four to 12 found in eelgrass (Zostera marina), Brachypodium distachyon, foxtail millet (Setaria italic), J. effuses, Aquilegia coerulea, papaya (Carica papaya), castor been (Ricinus communis), and physic nut (Jatropha curcas) (Supplemental Table S4). Among them, A. coerulea represents a basal eudicot that didn't experience the γ WGD shared by all core eudicots[50], whereas eelgrass is an early diverged aquatic monocot that didn't experience the τ WGD shared by all core monocots[56]. Interestingly, though both species possess two PIP1s and two PIP2s, they were shown to exhibit complex orthologous relationships of 1:1, 2:2, 1:0, and 0:1 (Supplemental Table S5). Whereas AcPIP1;1/AcPIP1;2/ZmPIP1;1/ZmPIP1;2 and ZmPIP2;1/AcPIP2;1 belong to PIP1A and PIP2A identified in this study, AcPIP2;2 and ZmPIP2;2 belong to PIP2H and PIP2I, respectively (Supplemental Table S5), implying that the last common ancestor of monocots and eudicots possesses only one PIP1 and two PIP2s followed by clade-specific expansion. A good example is the generation of AtPIP1;1 and -2;6 from AtPIP1;5 and -2;1 via the γ WGD, respectively[17].

    In tigernut, extensive expansion of the PIP subfamily was contributed by WGD (2), transposed (2), tandem (5), and dispersed duplications (3). It's worth noting that, two transposed repeats (i.e., CePIP1;1/-1;3 and CePIP2;1/-2;8) are shared by rice, implying their early origin that may be generated sometime after the split with the eudicot clade but before Cyperaceae-Poaceae divergence. By contrast, two WGD repeats (i.e., CePIP1;1/-1;2 and CePIP2;2/-2;4) are shared by C. cristatella, R. breviuscula, and J. effusus but not rice and Arabidopsis, implying that they may be derived from WGDs that occurred sometime after Cyperaceae-Poaceae split but before Cyperaceae-Juncaceae divergence. The possible WGD is the one that was described in C. littledalei[58], though the exact time still needs to be studied. Interestingly, compared with Arabidopsis (1) and rice (2), tandem/proximal duplications played a more important role in the expansion of PIP genes in tigernut (5) as well as other Cyperaceae species tested (5–6), which were shown to be Cyperaceae-specific or even species-specific. These tandem repeats may play a role in the adaptive evolution of Cyperaceae species as described in a high number of plant species[14,41]. According to comparative genomics analyses, tandem duplicates experienced stronger selective pressure than genes formed by other modes (WGD, transposed duplication, and dispersed duplication) and evolved toward biased functional roles involved in plant self-defense[41].

    As observed in most species such as Arabidopsis[10,1417], PIP genes in all Cyperaceae and Juncaceae species examined in this study, i.e., tigernut, C. cristatella, R. breviuscula, and J. effuses, feature three introns with conserved positions. By contrast, zero to three introns was not only found in rice but also in other Poaceae species such as maize, sorghum, foxtail millet, switchgrass, B. distachyon, and barley[54,55], implying lineage/species-specific evolution.

    Despite the extensive expansion of PIP genes (PIP2) in tigernut even after the split with R. breviuscula, CePIP1;1, -2;1, and -2;8 were shown to represent three dominant members in most tissues examined in this study, i.e., young leaf, mature leaf, sheath, rhizome, shoot apex, and tuber, though the situation in root is more complex. CePIP1;1 was characterized as a transposed repeat of CePIP1;3, which represents the most expressed member in root. Moreover, its recent WGD repeat CePIP1;2 was shown to be lowly expressed in most tested tissues, implying their divergence. The ortholog of CePIP1;1 in rice is OsPIP1;3 (RWC-3), which was shown to be preferentially expressed in roots, stems, and leaves, in contrast to constitutive expression of OsPIP1;1 (OsPIP1a) and -1;2[5961], two recent WGD repeats. Injecting the cRNA of OsPIP1;3 into Xenopus oocytes could increase the osmotic water permeability by 2–3 times[60], though the activity is considerably lower than PIP2s such as OsPIP2;2 and -2;2[6163]. Moreover, OsPIP1;3 was shown to play a role in drought avoidance in upland rice and its overexpression in lowland rice could increase root osmotic hydraulic conductivity, leaf water potential, and relative cumulative transpiration at the end of 10 h PEG treatment[64]. CePIP2;8 was characterized as a transposed repeat of CePIP2;1. Since their orthologs are present in both rice and Arabidopsis (Supplemental Table S3), the duplication event is more likely to occur sometime before monocot-eudicot split. Interestingly, their orthologs in rice, i.e., OsPIP2;1 (OsPIP2a) and -2;6, respectively, are also constitutively expressed[61], implying a conserved evolution with similar functions. When heterologously expressed in yeast, OsPIP2;1 was shown to exhibit high water transport activity[62,6466]. Moreover, root hydraulic conductivity was decreased by approximately four folds in OsPIP2;1 RNAi knock-down rice plants[64]. The water transport activity of OsPIP2;6 has not been tested, however, it was proven to be an H2O2 transporter that is involved in resistance to rice blast[61]. More work especially transgenic tests may improve our knowledge of the function of these key CePIP genes.

    Leaf is a photosynthetic organ that regulates water loss through transpiration. In tigernut, PIP transcripts in leaves were mainly contributed by CePIP1;1, -2;1, and -2;8, implying their key roles. During leaf development, in contrast to gradual decrease of CePIP1;1 and -2;8 transcripts in three stages (i.e. young, mature, and senescing) examined in this study, CePIP2;1 peaked in mature leaves. Their high abundance in young leaves is by cell elongation and enlargement at this stage, whereas upregulation of CePIP2;1 in mature leaves may inform its possible role in photosynthesis[67]. Thus far, a high number of CO2 permeable PIPs have been identified, e.g., AtPIP2;1, HvPIP2;1, HvPIP2;2, HvPIP2;3, HvPIP2;5, and SiPIP2;7[6870]. Moreover, in mature leaves, CePIP1;1, -2;1, and -2;8 were shown to exhibit an apparent diurnal fluctuation expression pattern that was expressed more in the day and usually peaked at noon, which reflects transpiration and the fact that PIP genes are usually induced by light[11,7173]. In rice, OsPIP2;4 and -2;5 also showed a clear diurnal fluctuation in roots that peaked at 3 h after the onset of light and dropped to a minimum 3 h after the onset of darkness[11]. Notably, further studies showed that temporal and dramatic induction of OsPIP2;5 around 2 h after light initiation was triggered by transpirational demand but not circadian rhythm[74].

    As an oil-bearing tuber crop, the main economic goal of tigernut cultivation is to harvest underground tubers, whose development is highly dependent on water available[32,75]. According to previous studies, the moisture content of immature tigernut tubers maintains more than 80.0%, followed by a seed-like dehydration process with a drop of water content to less than 50% during maturation[28,32]. Thereby, the water balance in developmental tubers must be tightly regulated. Like leaves, the majority of PIP transcripts in tubers were shown to be contributed by CePIP1;1, -2;1, and -2;8, which was further confirmed at the protein level. In accordance with the trend of water content during tuber development, mRNA, and protein abundances of CePIP1;1, -2;1, and -2;8 in initiation and swelling tubers were considerably higher than that at the mature stage. High abundances of CePIP1;1, -2;1, and -2;8 at the initiation stage reflects rapid cell division and elongation, whereas upregulation of CePIP1;1 and -2;1 at the swelling stage is in accordance with cell enlargement and active physiological metabolism such as rapid oil accumulation[28,30]. At the mature stage, downregulation of PIP transcripts and protein abundances resulted in a significant drop in the moisture content, which is accompanied by the significant accumulation of late embryogenesis-abundant proteins[23,32]. The situation is highly distinct from other tuber plants such as potato (Solanum tuberosum), which may contribute to the difference in desiccation resistance between two species[32,76]. It's worth noting that, in one study, CePIP2;1 was not detected in any of the four tested stages, i.e., freshly harvested, dried, rehydrated for 48 h, and sprouted tubers[23]. By contrast, it was quantified in all three stages of tuber development examined in this study, i.e., 1, 25, and 35 DAI (corresponding to freshly harvested tubers), which represent initiation, swelling, and maturation. One possible reason is that the protein abundance of CePIP2;1 in mature tubers is not high enough to be quantified by nanoLC-MS/MS, which is relatively less sensitive than 4D-PRM used in this study[30,46]. In fact, nanoLC-MS/MS-based proteomic analysis of 30 samples representing six tissues/stages only resulted in 2,257 distinct protein groups[23].

    Taken together, our results imply a key role of CePIP1;1, -2;1, and -2;8 in tuber water balance, however, the mechanism underlying needs to be further studied, e.g., posttranslational modifications, protein interaction patterns, and transcriptional regulators.

    To our knowledge, this is the first genome-wide characterization of PIP genes in tigernut, a representative Cyperaceae plant with oil-bearing tubers. Fourteen CePIP genes representing two phylogenetic groups or 12 orthogroups are relatively more than that present in two model plants rice and Arabidopsis, and gene expansion was mainly contributed by WGD and transposed/tandem duplications, some of which are lineage or even species-specific. Among these genes, CePIP1;1, -2;1, and -2;8 have evolved to be three dominant members that are constitutively expressed in most tissues, including leaf and tuber. Transcription of these three dominant members in leaves are subjected to development and diurnal regulation, whereas in tubers, their mRNA and protein abundances are positively correlated with the moisture content during tuber development. Moreover, their plasma membrane-localization was confirmed by subcellular localization analysis, implying that they may function in the cell membrane. These findings shall not only provide valuable information for further uncovering the mechanism of tuber water balance but also lay a solid foundation for genetic improvement by regulating these key PIP members in tigernut.

    The authors confirm contribution to the paper as follows: study conception and design, supervision: Zou Z; analysis and interpretation of results: Zou Z, Zheng Y, Xiao Y, Liu H, Huang J, Zhao Y; draft manuscript preparation: Zou Z, Zhao Y. All authors reviewed the results and approved the final version of the manuscript.

    All the relevant data is available within the published article.

    This work was supported by the Hainan Province Science and Technology Special Fund (ZDYF2024XDNY171 and ZDYF2024XDNY156), China; the National Natural Science Foundation of China (32460342, 31971688 and 31700580), China; the Project of Sanya Yazhou Bay Science and Technology City (SCKJ-JYRC-2022-66), China. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

  • The authors declare that they have no conflict of interest.

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  • Cite this article

    Duan ZY, Yu J, Zhao CL. 2022. Molecular phylogeny and morphology reveal a new wood-rotting fungal species, Cyathus wenshanensis sp. nov. from the Yunnan-Guizhou Plateau. Studies in Fungi 7:8 doi: 10.48130/SIF-2022-0008
    Duan ZY, Yu J, Zhao CL. 2022. Molecular phylogeny and morphology reveal a new wood-rotting fungal species, Cyathus wenshanensis sp. nov. from the Yunnan-Guizhou Plateau. Studies in Fungi 7:8 doi: 10.48130/SIF-2022-0008

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Molecular phylogeny and morphology reveal a new wood-rotting fungal species, Cyathus wenshanensis sp. nov. from the Yunnan-Guizhou Plateau

Studies in Fungi  7 Article number: 8  (2022)  |  Cite this article

Abstract: A new species of bird's nest fungus, Cyathus wenshanensis is proposed based on a combination of the morphological and molecular evidence. It is characterised by the obconical to cupulate basidiomata covered with hirsute hairs, striations on the outer and inner surface of the peridium, funicular peridioles, a trimitic hyphal system of peridium with generative hyphae having clamp connections, a dimitic hyphal system of peridiole middle, and subglobose, elliptical to ellipsoid-elongate, thick-walled basidiospores. Sequence of the internal transcribed spacers (ITS) gene region was generated, and the phylogenetic analysis was performed with maximum likelihood, maximum parsimony and Bayesian inference methods. The phylogenetic analyses inferred from ITS dataset indicated that C. wenshanensis nested within the genus Cyathus, in which it formed a monophyletic lineage and grouped with C. albinus, C. amazonicus, C. badius, C. parvocinereus, C. pyristriatus and C. uniperidiolus.

    • The genus Cyathus (Nidulariaceae, Nidulariales) was first introduced by Haller[1] and later was adopted by Persoon[2], typified by C. striatus (Huds.) Willd. Cyathus together with Crucibulum Tul. & C. Tul., Mycocalia J.T. Palmer, Nidula V.S. White, and Nidularia Fr., are commonly known as bird's nest fungi due to their cup-like basidiomata resembling bird nest and lenticular periodioles resembling eggs[35]. It is characterized by having deeper or cuped, inverted bell-like basidiomata covered with shaggy or tomentose hairs on the outside; peridium composed of three layers of tissues, inside peridium filled with a number of dark-colored, small, hard lentil-shaped peridioles attached with funicular cords; colorless, thin-walled or thick-walled, smooth basidiospores[3,611]. The species of Cyathus are saprobic, usually growing in decaying wood, on manure or directly on soil are a cosmopolitan group and have a rich diversity related to the high diversity of plants growing in boreal, temperate, subtropical, and tropical regions[3,5,1214]. Both MycoBank database (www.MycoBank.org; 23 June 2022) and Index Fungorum (www.indexfungorum.org; 23 June 2022) register 204 specific and infraspecific names in the genus Cyathus, but the actual number of species are about 60[15], including 35 species from China[5,16].

      Molecular systematic studies of the genus Cyathus have been carried out previously[14,17,18]. An overview of the phylogeny of the Agaricales presented based on a multilocus analysis of a six-gene region supermatrix revealed that the family Nidulariaceae was sister to Cystodermateae, in which Cyathus striatus and Crucibulum laeve grouped together within Nidulariaceae[17]. Phylogenetic relationships within the genus Cyathus (bird's nest fungi) were investigated with neighbor joining, maximum likelihood, weighted maximum parsimony and MrBayes analyses of the internal transcribed spacers (ITS) and large subunit (LSU) of ribosomal DNA sequences datasets, in which the morphological characters of the peridium plications and variations in peridium hair anatomy, peridiole structure and fruit-body color were not supported by the molecular data, while the ITS and LSU datasets supported the recognition of three infrageneric groups herein named the ollum, pallidum and striatum groups[18]. Phylogenetic analyses based on ITS and LSU ribosomal DNA sequences revealed that three taxa C. cheliensis, C. gansuensis, and C. megasporus were respectively accepted as synonyms of C. limbatus, C. pygmaeus, and C. poeppigii[5]. On the basis of the morphological and molecular data, Martin et al.[19] discussed affinities among Cyathus species, which showed that this group formed a monophyletic group with high support. Phylogenetic reconstruction of Cyathus species based on alignment of 641 nucleotides of the ITS region indicated that three new species as C. batistae and C. apiculatus, C. pedunculatus were proposed, and discussed relationships with other species of Cyathus[20]. Phylogenetic relationships of bird's nest fungi investigated with four commonly used loci (ITS, LSU, translation elongation factor (TEF), and RNA polymerase II second largest subunit (RPB2)) revealed that the family Nidulariaceae was resolved as a monophyletic group with Squamanitaceae as a potential sister taxon, and suggested that species concepts needed to be revisited and refined throughout Nidulariaceae and several bird's nest fungi species had global geographical distributions, whereas others may have more limited ranges, and the basic morphological characters of bird's nest fungi had likely been lost or gained multiple times[21]. The phylogenetic study using five loci (ITS, LSU, SSU, translation elongation factor 1-alpha (TEF1) and RPB2) revealed that a new genus Retiperidiolia to accommodate this phylogenetically and morphologically unique bird's nest fungus lineage, in which Cyathus formed a monophyletic lineage and then was sister to the genus Retiperidiolia[14].

    • The fresh fruiting bodies of the bird's nest fungi were collected from Wenshan (Yunnan Province, P. R. China). The fresh specimens were dried in an electric food dehydrator at 40 °C, then sealed and stored in an envelope bag and deposited in the herbarium of the Southwest Forestry University (SWFC, Kunming, Yunnan Province, P.R. China).

      The macromorphological descriptions were based on field notes and photos captured in the field and lab. The macromorphological descriptions are based on Brodie[3]. The micromorphological data were obtained from the dried specimens and observed under Nikon Eclipse E100 light microscope following the methods of Zhao & Wu[22]. Color terms follow Kornerup & Wanscher[23]. Drawings were made with the aid of a fungus plotter. The measurements and drawings were made from slide preparations stained with Cotton Blue (0.1 mg aniline blue dissolved in 60 g pure lactic acid), Melzer's reagent (3 g potassium iodide, 1 g crystalline iodine, 44 g chloral hydrate, aq. dest. 40 ml) and 5% potassium hydroxide. In presenting spore size data, 5% of the measurements excluded from each end of the range are shown in parentheses. The following abbreviations are used: KOH = 5% potassium hydroxide; CB = cotton blue; CB– = acyanophilous; IKI = Melzer's reagent; IKI– = non-amyloid and non-dextrinoid; L = mean spore length (arithmetic average of all spores); W = mean spore width (arithmetic average of all spores); Q = L/W ratio; n = number of spores/measured from a given number of specimens.

    • The CTAB rapid plant genome extraction kit-DN14 (Aidlab Biotechnologies Co., Ltd, Beijing, China) was used to obtain genomic DNA from dried fungal specimens, according to the manufacturer's instructions. The ITS region was amplified with the primer pair ITS5 and ITS4[24]. The PCR cycling procedure for ITS was as follows: initial denaturation at 95 °C for 3 min, followed by 35 cycles at 94 °C for 40 s, 58 °C for 45 s and 72 °C for 1 min, and a final extension of 72 °C for 10 min. The PCR products were purified and directly sequenced at Kunming Tsingke Biological Technology Limited Company (Yunnan Province, P.R. China). All newly generated sequences were deposited in GenBank (Table 1).

      Table 1.  List of species, specimens, and GenBank accession numbers of ITS sequences used in this study.

      Species nameSample no.GenBank
      accession no.
      References
      Crucibulum laeveSWFC 21261DQ463357Zhao et al.[18]
      Cyathus africanusDAOM 200370[T]DQ463347Zhao et al.[18]
      C. albinusUFRN-Fungos 2239KY176371Accioly et al.[25]
      C. amazonicusURM 80036[T]KY495280Accioly et al.[25]
      C. amazonicusUFRN-Fungos 2798KY176375Accioly et al.[25]
      C. annulatusMichaelKuo-8200901MT444076Kraisitudomsook et al.[21]
      C. apiculatusUFRN:Fungos 1448KT365516da Silva et al.[20]
      C. aurantogriseocarpusUFRN:Fungos:2798KX966026da Cruz et al.[26]
      C. badiusKH:JPN15-1321KX906250da Cruz et al.[27]
      C. batistaeUFRN:Fungos 1449KT365515daSilva et al.[20]
      C. berkeleyanusSWFC 20789DQ463355Zhao et al.[18]
      C. bulleriDAOMC 195062MK020156Vats & Mishra[28]
      C. cannaCBS 370.80MH861275Vu et al.[29]
      C. colensoiDAOM 200423DQ463344Zhao et al.[18]
      C. crassimurusDAOM 200372[T]DQ463350Zhao et al.[18]
      C. discoideusAB 7831KY652080da Cruz[30]
      C. gansuensisSWFC 20880[T]DQ463348Zhao et al.[18]
      C. gansuensisStrain 69KC869661da Cruz et al.[27]
      C. gracilisAB7873KY652081da Cruz[30]
      C. hookeriSWFC 20799DQ463346Zhao et al.[18]
      C. hortensisUFRN:Fungos:1819KX906252da Cruz et al.[27]
      C. ibericusAH:48138KX858598Crous et al.[31]
      C. ibericusAH:48137[T]KX858597Crous et al.[31]
      C. intermediusUFRN:Fungos 1033KT365519da Silva et al.[20]
      C. jiayuguanensisSWFC 20846[T]DQ463341Zhao et al.[18]
      C. lignilantanaeMA Fungi 87327NR_154827da Cruz et al.[27]
      C. limbatusUFRN-Fungos 2238KY176373Accioly et al.[25]
      C. magnomuralisUFRN:Fungos:1817KX906251da Cruz et al.[27]
      C. minimusAB7868KY652082da Cruz[30]
      C. novae-zeelandiaePDD-76442MT444096Kraisitudomsook et al.[21]
      C. ollaPDD-86833MT444086Kraisitudomsook et al.[21]
      C. ollaBPI 727227DQ463345Zhao et al.[18]
      C. pallidusKKUITN2KU202745Sutthisa & Sanoamuang[32]
      C. pallidusKKUITN3KU202751Sutthisa & Sanoamuang[32]
      C. parvocinereusUFRN:Fungos:1814KX906253da Cruz et al.[27]
      C. pedunculatusUFRN:Fungos 403KT365518da Silva et al.[20]
      C. poeppigiicp-457KT962176da Silva et al.[20]
      C. pyristriatusMFLUCC:14-0770KU865513Richter et al.[33]
      C. renweiiSWFC 201406[T]DQ463352Zhao et al.[18]
      C. setosusDAOM 200815[T]DQ463349Zhao et al.[18]
      C. stercoreusNK-08MT444037Kraisitudomsook et al.[21]
      C. stercoreusDM4KY706156Hay et al.[34]
      C. striatusNK-61MT444056Kraisitudomsook et al.[21]
      C. subglobisporusBBH-14815MT444063Kraisitudomsook et al.[21]
      C. subglobisporuBBH18348EF613553Zhao et al.[5]
      C. triplexSWFC 21077DQ463353Zhao et al.[18]
      C. uniperidiolusAMH:10196MN398297Boonmee et al.[35]
      C. wenshanensisCLZhao 20202[T]ON795104This study
      Nidula niveotomentosaSWFC 3000DQ463358Zhao et al.[18]

      Sequencher 4.6 (GeneCodes, Ann Arbor, MI, USA) was used to assemble and edit the generated sequence reads. Sequences were aligned in MAFFT 7 (https://mafft.cbrc.jp/alignment/server/) using the 'G-INS-I' strategy and manually adjusted in BioEdit[36]. Crucibulum laeve (Huds.) Kambly and Nidula niveotomentosa (Henn.) Lloyd were selected as an outgroup for the phylogenetic analysis of the ITS phylogenetic tree[25].

      Maximum parsimony (MP), Maximum Likelihood (ML) and Bayesian Inference (BI) analyses were applied to the ITS dataset sequences. Approaches to phylogenetic analyses followed[22]. MP analysis was performed in PAUP* version 4.0b10[37]. All of the characters were equally weighted and gaps were treated as missing data. Trees were inferred using the heuristic search option with TBR branch swapping and 1000 random sequence additions. Maxtrees were set to 5000, branches of zero length were collapsed and all most-parsimonious trees were saved. Clade robustness was assessed using bootstrap (BT) analysis with 1,000 replicates[38]. Descriptive tree statistics tree length (TL), the consistency index (CI), the retention index (RI), the rescaled consistency index (RC) and the homoplasy index (HI) were calculated for each most-parsimonious tree generated. ML was inferred using RAxML-HPC2 through the Cipres Science Gateway (www.phylo.org)[39]. Branch support (BS) for ML analysis was determined by 1000 bootstrap replicates and evaluated under the gamma model.

      MrModeltest 2.3[40] was used to determine the best-fit evolution model for the dataset for Bayesian Inference (BI). Bayesian Inference was performed with MrBayes 3.1.2 with a general time reversible (GTR+I+G) model of DNA substitution and a gamma distribution rate variation across sites[41]. Four Markov chains were used in each of two runs from random starting trees for 1.5 million generations (Fig. 1), with trees and parameters sampled every 100 generations. The first quarter of the generations were discarded as burn-in. A majority rule consensus tree of all remaining trees and posterior probabilities were calculated. Branches were considered significantly supported if they received maximum likelihood bootstrap value (BS) of > 70%, a maximum parsimony bootstrap value (BT) of > 70%, or Bayesian posterior probabilities (BPP) of > 0.95.

      Figure 1. 

      Maximum parsimony strict consensus tree illustrating the phylogeny of the new species and related species in genus Cyathus based on ITS sequences. Branches are labelled with maximum likelihood bootstrap value > 70%, parsimony bootstrap value > 50% and Bayesian posterior probabilities > 0.95, respectively. The present species are in bold.

    • The ITS dataset (Fig. 1) included sequences from 49 fungal specimens representing 42 species. The dataset had an aligned length of 805 characters, of which 270 characters were constant, 232 were variable and parsimony-uninformative, and 303 parsimony-informative. The MP analysis yielded one equally parsimonious trees (TL = 1211, CI = 0.6474, HI = 0.3526, RI = 0.7855, RC = 0.5086). Best model for the ITS dataset estimated and applied in the Bayesian analysis: GTR+I+G, lset nst = 6, rates = invgamma; prset statefreqpr = dirichlet (1,1,1,1). The bayesian and ML analyses resulted in a similar topology as MP analysis, with an average standard deviation of split frequencies = 0.009975 (BI), and the effective sample size (ESS) across the two runs was double the average ESS (avg ESS) = 248. The phylogenetic tree (Fig. 1) inferred from ITS sequences revealed that C. wenshanensis nested within the genus Cyathus, in which it formed a monophyletic lineage and grouped with C. albinus, C. amazonicus, C. badius, C. parvocinereus, C. pyristriatus and C. uniperidiolus.

    • Cyathus wenshanensis Z.Y. Duan & C.L. Zhao, sp. nov. Figs 25.

      Figure 2. 

      Basidiomata of Cyathus wenshanensis. (a) Basidiomata, (b) outer corving of peridium. Scale bars: (a) = 1 cm, (b) = 1 mm.

      Figure 3. 

      Peridiole of Cyathus wenshanensis. (a) Peridioles with funicular cord, (b) transversal section of peridiole showing single-layered cortex. Scale bars: (a) = 1 mm, (b) = 1 mm.

      Figure 4. 

      Microscopic structures of Cyathus wenshanensis. (a) Outer wall of peridium, (b) inner wall of peridium, (c) three-layered peridium, (d) the structure of the hair. Scale bars: (a)–(d) = 10 μm.

      Figure 5. 

      Microscopic structures of Cyathus wenshanensis. (a) Basidiospores. (b) Funicular cord. (c) The internal structure of peridiole. Scale bars: (a)–(c) = 10 μm.

      Index Fungorum number: IF844702; Facesoffungi number: FoF12564

      Etymology – wenshanensis (Lat.): referring to the provenance (Wenshan) of the type specimens.

      Basidiomata obconical to cupulate, 5–15 mm high, 5–10 mm wide at the mouth, without expanding at the top or tapering abruptly at the base; the base usually attached to the substrate by a slightly conspicuous emplacement, brown (5E6) to beige (4C3); exoperidium brown (5E6), hirsute, external wall striate near the mouth, 0.4–0.7 mm between folds, covered with brown (5E6) to dark brown (7F6), irregular and flexible tufts of hair; hair hyphae with clamp connections, colorless, thick-walled (wall up to 0.5–2 μm thick), 3.5–13.5 µm in diameter; endoperidium greyish brown (8F3) to black brown (7F4), conspicuously striate with 0.4–0.8 mm between the groves; mouth finely fimbriate; peridium walls consist of three different layers: (1) outer wall layer, hyphal system trimitic, CB–, IKI–, tissues unchanged in KOH; generative hyphae with clamp connections, colorless to pale brown, slight thick-walled, frequently branched, 1.5–4 µm in diameter; skeletal hyphae colorless to pale brown, thick-walled, unbranched, 2–4 µm in diameter; binding hyphae colorless to pale brown, thick-walled, unbranched, 1.5–2.5 µm in diameter; (2) inner wall layer, hyphal system trimitic, CB–, IKI–, tissues unchanged in KOH; generative hyphae with clamp connections, colorless to pale brown, slight thick-walled, rarely branched, 2–4 µm in diameter; skeletal hyphae colorless to pale brown, thick-walled, unbranched, 2.5–4.5 µm in diameter; binding hyphae colorless to pale brown, thick-walled, rarely branched, 1.5–3 µm in diameter; (3) pseudopare-chymatous layer.

      Peridioles depressed, shiny, angular to irregular, suborbicular, broadly ellipsoid to ovoid, dark grey (8F1) to black (6F3), surface smooth to wrinkled, tunica present, often inconspicuous, cortex single-layered, 2.5–3.5 × 2–3 mm; funicular cord present, funiculus hyphae with clamp connections, thick-walled, unbranched, pale yellowish, 1–3.5 µm in diameter; hyphal system of peridiole middle dimitic, generative hyphae with clamp connections, colorless, thin-walled, frequently branched, with oil drops inside, 1–3 µm in diameter, CB–, IKI–, tissues unchanged in KOH; skeletal hyphae colorless, slight thick-walled, unbranched, with oil drops inside, 1.5–4 µm in diameter, CB–, IKI–, tissues unchanged in KOH.

      Basidiospores subglobose, elliptical to ellipsoid-elongate, colorless, smooth, thick-walled (wall up to 1–5 μm thick), CB–, IKI–, with inclusions or oil-like globule, without apiculus, (10–)11–21(–22) × 9–14(–15) µm, L = 16.34 µm, W = 11.51 µm, Q = 1.4 (n = 60/1). Basidia not observed.

      Known distribution – Thus far known only from China.

      Material examined – China. Yunnan Province, Wenshan, Pingba Town, Huguangqing Village, 23.26°N, 104.06°E, on the fallen branch of angiosperm, 12 August 2020, collected by C.L. Zhao. Specimen voucher number: CLZhao 20202 (SWFC 020202).

    • In the present study, C. wenshanensis sp. nov. is described based on the phylogenetic analyses and morphological characteristics.

      Phylogenetically, the molecular systematics and taxonomic overview of the bird's nest fungi revealed that the family Nidulariaceae was resolved as a monophyletic group with Squamanitaceae as a potential sister taxon, in which Cyathus and Crucibulum each formed its own independent and well-supported clade, and Nidula and Nidularia formed a clade together, but each genus is polyphyletic[21]. In the present study, C. wenshanensis nests within the genus Cyathus located in the family Nidulariaceae, in which it forms a monophyletic lineage and then groups with taxa C. albinus, C. amazonicus, C. badius, C. parvocinereus, C. pyristriatus and C. uniperidiolus. However, morphologically C. albinus differs from C. wenshanensis by having the golden blond to dark blond exoperidium, brownish gray peridioles with double-layered cortex, and basidiospores with conspicuous apiculous[25]. C. amazonicus differs in having very dark brown to grayish dark brown exoperidium and and gray, shiny endoperidium[42]. C. badius differs in having the smooth exoperidium, light brown to orange endoperidium, ovoid basidiospores[43]. C. parvocinereus differs in having the campanulate, smaller basidiomata (4–7 × 3.5–5 mm), pearl grey to brightness silvery endoperidium and greyish brown to grey peridioles with double-layered cortex[44]. C. pyristriatus differs in its clavate basidiomata with yellowish-brown or buff exoperidium, grey to dark grey endoperidium, greyish-brown peridioles, and ovoid basidiospores[45]. C. uniperidiolus distinct from C. wenshanensis in having the globose to sub-globose basidiomata with serrate margin at mouth, smooth peridium walls, and globose, smooth peridioles[35] (see Table 2).

      Table 2.  The comparison among Cyathus wenshanensis and phylogenetically related species.

      C. wenshanensisC. albinusC. amazonicusC. badiusC. parvocinereusC. pyristriatusC. uniperidiolus
      BasidiomataSize (high
      × wide)
      5–15 × 5–10 mm6–8.5 × 5–6.52 mm9–11 × 5–7 mm8–10 × 5–8 mm4–7 × 3.5–5 mm5.5–7 × 4–6 mm2–12 × 2–3.5 mm
      ShapeObconical to
      cupulate
      InfundibuliformObconicalInfundibuliformCampanulateClavate to broadly obconicGlobose to sub-globose
      ExoperidiumColourBrownGolden blond to dark blondVery dark brown to grayish dark brownBrownReddish brownYellowish-brown or buffDark brown
      SurfaceStrigose Tufts; striate, 0.4–0.7 mmStrigose tufts; striate; 0.3–0.5 mmStrigose tufts; striateShaggy, wooly tufts; smooth to striate;
      0.3 mm
      Strigose tufts; striate; 0.4–0.5 mmshaggy or fluffy hairsSmooth to velvety
      MouthFimbriateFimbriateFimbriateFimbriateFimbriateSerrate
      EndoperidiumColourGreyish brown to black brownGrayish brownGray to brownish grayLight brown to orangePlatinumGrey to dark greyDark brown
      SurfaceStriate, 0.4–0.8 mmStriate, 0.3–0.6 mmStriateSmooth to minutely striate; 0.5 mmStriate; 0.5 mmStriateSmooth
      PeridiolesShapeLentil-shapedLentil-shapedLentil-shapedLentil-shapedLentil-shapedLentil-shapedGlobose
      Size2–3.5 mm1.8–2.6 mm1.7–3 mm2–2.5 mm1–2 mm3–3.5 mm2–2.5 mm
      ColorDark grey to blackBrownish grayDark grayLight grey to blackGreyish brown to greyGreyish-brown to dark greyBlack
      CortexSingle layeredDouble layeredSingle layeredSingle layeredDouble layeredDouble layered
      BasidiosporesShapeSubglobose, elliptical to ellipsoid-elongate; apiculum absentOvoid to ellipsoid; apiculus presentSubglobose to broadly ellipsoidSubglobose, ovoid to ellipticalelliptical, globose; apiculum absentOvoid, subglobose, ellipsoid to broadly ellipsoidOval, sub-globose, broadly ellipsoid to ellipsoid-elongate
      Size11–21 × 9–14 µm14.8–20 × 10.4–14.3 µm14–19 × 12–16 µm13–19 × 9–11 µm11.43–17.78 × 9–15.24 µm14–17 × 8–10 µm14.2–28.7 × 11.7–23.7 µm
      Walls1–5 μm thick0.8–1.3 μm thickthick-walled1.9–3.2 μm thick2–3.5 μm thick1.5–3 μm thickthick-walled
      DistributionChinaBrazilAmazon rainforestJapan, BrazilBrazilThailandIndia
      ReferencePresent studyAccioly et al.[25]Trierveiler-Pereira et al.[42]da Cruz et al.[43]da Cruz & Baseia[44]Hyde et al.[45]Boonmee et al.[35]

      Morphologically, six taxa of Cyathus as C. apiculatus, C. hortensis, C. limbatus, C. lignilantanae, C. pedunculatus, and C. poeppigii are similar to C. wenshanensis on the basis of the character by having the obvious stripes on the inner and outer walls of peridium. However, C. apiculatus differs from C. wenshanensis by the basidiomata being expanded at the mouth and abruptly tapering to the base, silvery endoperidium, smaller peridioles (1–1.5 × 1.5–2 mm), and longer basidiospores (22–37 × 10–22 μm)[20]; C. hortensis is distinguished from C. wenshanensis by its basidiomata constricting abruptly at the base and forming a slender stipe, cinnamon exoperidium, smaller peridioles (1.2–2 × 1–1.5 mm) with double-layered cortex, and ovoid, wider basidiospores (17–34 × 13–20 μm)[44]; C. limbatus differs from C. wenshanensis by its double-layered peridioles, and basidiospores with apiculus[46]; C. lignilantanae is different from C. wenshanensis by having a reddish brown exoperidium, brownish grey to greyish brown, smaller peridioles (2.1–2.3 × 1.8–2 mm) with double-layered cortex[19]; C. pedunculatus is separated from C. wenshanensis by having the basidiomata abruptly tapering in the base forming a conspicuous pedicel, pale yellow to dark blond exoperidium, double-layered cortex, brownish grey, smaller peridioles (1.5–2 × 1–1.5 mm), and larger basidiospores (25–34 × 22–29 μm)[20]; C. poeppigii is distinguished from C. wenshanensis by having the narrowly obconical basidiomata with incurved mouths and a slender stipe at the base, and dark brown, smaller peridioles (1.5–2 mm) with double cortex, and larger basidiospores (30–45 × 18–30 μm)[47].

      Several taxa, Cyathus batistae, C. discoideus, C. gracilis, C. hookeri, C. magnomuralis, C. renweii and C. triplex are similar to C. wenshanensis based on the character having the fimbriate of basidiomata mouth. However, C. batistae differs from C. wenshanensis by its expanded mouth of basidiomata, with the stipe, smooth exoperidium wall, double-layered cortex peridioles, and smaller basidiospores (9–13 × 5–8 μm) with apiculus[20]; C. discoideus differs from C. wenshanensis by having grey brown, smaller peridioles (1.56–2.16 × 1.41–1.74 mm)[30]; C. gracilis is distinguished from C. wenshanensis by the basidiomata with slender base, umber to rusty outer surface of peridium, double-layered cortex peridioles, and basidiospores with apical notch[48]; C. hookeri differs from C. wenshanensis by its smooth peridium walls, and smaller basidiospores (9–13 × 5–8 μm)[49]; C. magnomuralis is distinguished from C. wenshanensis by having the dark blond exoperidium, smaller peridioles (1–1.5 × 1–1.5 mm) with double-layered cortex, and ovoid, larger basidiospores (27–49 × 23–41 μm) with small apiculus[44]; C. renweii differs from C. wenshanensis by its greyish peridioles with the brown tunica, and longer basidiospores (21–31 × 10.5–13.5 μm)[50]; C. triplex is separated from C. wenshanensis by its smaller basidiomata (5–8 × 4.5–5 mm) with the slender base orbicular, flattened peridioles with double-layered cortex, and basidiospores with the apical notch[25].

      Eight species of the genus Cyathus as C. africanus, C. colensoi, C. gansuensis, C. ibericus, C. jiayuguanensis, C. novae-zeelandiae, C. olla, and C. pallidus are similar to C. wenshanensis in light of the characteristics of having single-layered cortex peridioles. However, C. africanus differs from C. wenshanensis by its peridium walls with woolly hairs, silvery peridioles, and broadly ovate, smaller basidiospores (8.5–12 × 6.5–8.5 μm) with apiculus[51]; C. colensoi differs from C. wenshanensis by its smooth peridium walls, and ovoid, smaller basidiospores (8.5–11.5 × 7–8.5 μm)[30,49]; C. gansuensis differs from C. wenshanensis by its narrow base basidiomata with grayish to dark smoke-gray interior, grayish, smaller peridioles (1.5–2 × 0.8–1.5 mm), and ovoid basidiospores[52]; C. ibericus differs in its whitish to pale brownish grey external peridium with woolly hairs, smaller peridioles (0.8–1.2 mm diam), and ovoid, smaller basidiospores (7–9 × 5–6 μm)[30]; C. jiayuguanensis differs from C. wenshanensis by its basidiomata with the short stipe, smoke-gray peridioles, and ovoid, smaller basidiospores (8–11.5 × 7–8.5 μm)[52]; C. novae-zeelandiae differs in C. wenshanensis by its basidiomata abruptly constricted into a stipe, peridioles with white tunica, and smaller basidiospores (11–13 × 5–6 μm)[4,53]; C. olla differs from C. wenshanensis by its peridium with tomentose outside, silver, smooth inside, pure silver peridioles, and the smaller basidiospores (9.8–11.2 × 6.4–8 μm)[54]; C. pallidus differs from C. wenshanensis by its smaller basidiospores (6.8–14.5 × 6–8.1 μm)[32].

      Cyathus annulatus, C. aurantogriseocarpus, C. minimus, and C. stercoreus are similar to C. wenshanensis inferred from the characteristics of having thick-walled basidiospores without apiculus. However, C. annulatus is separated from C. wenshanensis by its expanded peridium at the top, ochraceous-tawny exoperidium, pale buff inner surface, subtriangular peridioles, and the striking dark-brown ring at the mouth[55]; C. aurantogriseocarpus differs from C. wenshanensis by the orange-grey exoperidium with long tomentum, brownish grey, smaller peridioles (1.5–1.75 × 1.2–1.5 mm) with double-layered cortex, and larger basidiospores (32.5–47 × 22.5–28.5 μm)[26]; C. minimus differs from C. wenshanensis by the clay brown exoperidium, yellowish brown endoperidium, and reddish brown, coffee or brown tobacco, smaller peridioles (1.3–1.37 × 1.13–1.23 mm)[30]; C. stercoreus differs in its smooth peridium walls, double-layered cortex peridioles and larger basidiospores (30–41 × 25–31 μm)[4].

      The family Nidulariaceae is a characteristic group of Agaricomycetes (Basidiomycota), which has a number of macrofungi based on a result of the morphological, phylogenetic and cytological studies in China[56,57], but the species diversity of macrofungi are still not well known, especially in subtropical and tropical areas of the country[5861]. The new species, Cyathus wenshanensis is from the subtropics. Therefore, the present paper enriches the fungal diversity in the Chinese ecosystem, and it is likely that more new taxa will be found after further fieldwork and molecular analyses.

      In addition, the results of BLAST queries in NCBI based on ITS separately showed the sequences producing significant alignments descriptions: in ITS blast results, the top ten taxa are C. pyristriatus (Max score 1116; Total score 1116; Query cover 92%; E value 0.0; Ident 95.47%), C. parvocinereus (Maximum record descriptions: Max score 1081; Total score 1081; Query cover 89%; E value 0.0; Ident 95.71%), C. amazonicus (Maximum record descriptions: Max score 1077; Total score 1077; Query cover 87%; E value 0.0; Ident 96.11%), C. uniperidiolus (Maximum record descriptions: Max score 1075; Total score 1075; Query cover 91%; E value 1e-78; Ident 94.84%), C, albinus (Max score 1053; Total score 1053; Query cover 85%; E value 0.0; Ident 96.01%), and C. badius (Maximum record descriptions: Max score 1000; Total score 1000; Query cover 83%; E value 0.0; Ident 95.15%).

      • The research was supported by the National Natural Science Foundation of China (Project No. 32170004), Yunnan Fundamental Research Project (Grant No. 202001AS070043) and received support from Yunnan Academy of Biodiversity, Southwest Forestry University.

      • The authors declare that they have no conflict of interest.

      • Copyright: © 2022 by the author(s). Published by Maximum Academic Press, Fayetteville, GA. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
    Figure (5)  Table (2) References (61)
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    Duan ZY, Yu J, Zhao CL. 2022. Molecular phylogeny and morphology reveal a new wood-rotting fungal species, Cyathus wenshanensis sp. nov. from the Yunnan-Guizhou Plateau. Studies in Fungi 7:8 doi: 10.48130/SIF-2022-0008
    Duan ZY, Yu J, Zhao CL. 2022. Molecular phylogeny and morphology reveal a new wood-rotting fungal species, Cyathus wenshanensis sp. nov. from the Yunnan-Guizhou Plateau. Studies in Fungi 7:8 doi: 10.48130/SIF-2022-0008

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