Introduction
Xanthoparmelia is the most speciose genus of lichen-forming fungi (c. 800 species) (Thell et al. Reference Thell, Crespo, Divakar, Kärnefelt, Leavitt, Lumbsch and Seaward2012; Jaklitsch et al. Reference Jaklitsch, Baral, Lücking, Lumbsch and Frey2016). The genus has a worldwide distribution and is found in temperate to tropical, semi-arid and arid regions (Hale Reference Hale1990). South Africa and Australia are the distribution centers (Elix et al. Reference Elix, Johnston and Armstrong1986; Hale Reference Hale1990). While Xanthoparmelia species are equally species-rich in both regions, macro-evolutionary patterns of lineage diversification are quite distinct. South Africa and nearby regions have been suggested as the areas of evolutionary origin for Xanthoparmelia, which is estimated to be close to the Oligocene-Miocene boundary (Divakar et al. Reference Divakar, Crespo, Wedin, Leavitt, Hawksworth, Myllys, McCune, Randlane, Bjerke and Ohmura2015; Leavitt et al. Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018). Multiple early-diverging lineages diversified in South Africa, with several migration events into Australia (Leavitt et al. Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018; Autumn et al. Reference Autumn, Barcenas-Peña, Kish-Levine, Huang and Lumbsch2020). The species diversity in Australia might result from several subsequent radiation events (Leavitt et al. Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018). In contrast, the Northern Hemisphere was colonized recently by the genus, with an estimated age of the Holarctic clade of 7.2 Mya (Leavitt et al. Reference Leavitt, Lumbsch, Stenroos and St Clair2013, Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018). We focused our study on southern Africa, where currently more than 300 species are known, including the synonymized genera Karoowia, Namakwa, Neofuscelia, Paraparmelia and Xanthomaculina (Esslinger Reference Esslinger1981, Reference Esslinger1986, Reference Esslinger2000; Hale Reference Hale1985, Reference Hale1988, Reference Hale1989, Reference Hale1990; Elix Reference Elix1997, Reference Elix1999a, Reference Elixb, Reference Elix2001, Reference Elix2002, Reference Elix2003; Elix et al. Reference Elix, Becker and Follmann1999; Blanco et al. Reference Blanco, Crespo, Elix, Hawksworth and Lumbsch2004; Thell et al. Reference Thell, Feuerer, Kärnefelt, Myllys and Stenroos2004; Amo de Paz et al. Reference Amo de Paz, Lumbsch, Cubas, Elix and Crespo2010; Sipman Reference Sipman2017). We used the available data from recent collections by one of our researchers (VW) through somewhat opportunistic sampling. Therefore, the sampling has limitations and does not include all South African Xanthoparmelia species. However, a thorough sampling of specimens in the studied area has been carried out (Wirth & Sipman Reference Wirth and Sipman2018; Wirth et al. Reference Wirth, Sipman and Curtis-Scott2018). This study aims to contribute to a better understanding of species delimitation and phylogenetic relationships of Xanthoparmelia in South Africa.
Material and Methods
Study area
Our study is based on 61 Xanthoparmelia specimens collected during an inventory of natural vegetation remnants in the Cederberg and Renosterveld Reserve regions of the Western Cape, South Africa, by VW (Wirth & Sipman Reference Wirth and Sipman2018; Wirth et al. Reference Wirth, Sipman and Curtis-Scott2018), for which new sequences were generated. The specimens are deposited in B and STU, and we used collections in F for comparison.
Anatomical studies
Thallus morphology was studied using a Zeiss Stemi 2000-C stereomicroscope, and conidia and spore shape and size were observed using a Zeiss Axioscope. Secondary metabolites were identified using spot tests with 10% KOH, C (sodium hypochlorite), KC and PD (paraphenylenediamine) and high-performance thin-layer chromatography (HPTLC) (Culberson & Johnson Reference Culberson and Johnson1982; Arup et al. Reference Arup, Ekman, Lindblom and Mattsson1993; Orange et al. Reference Orange, James and White2010).
Molecular methods
Sixty-one specimens from South Africa were selected for molecular analysis. Total genomic DNA was extracted from thallus fragments using the ZR Fungal/Bacterial DNA Miniprep Kit (Zymo Research Corp., Irvine, CA, USA), following the manufacturer's instructions. DNA sequences were generated for three markers using the polymerase chain reaction (PCR): the nuclear ribosomal internal transcribed spacer region (ITS), a region of the mitochondrial small subunit rDNA (mtSSU), and a region of the nuclear large subunit rDNA (nuLSU). PCR reactions contained 6.25 μl of MyTaq™ Red DNA Polymerase (Bioline, Taunton, MA, USA), 5.25 μl of H2O, 0.25 μl of forward and reverse primers (10 μM), and 0.5 μl of template DNA (10×), for a total reaction volume of 12.5 μl. The ITS region was amplified using the primers ITS1F (Gardes & Bruns Reference Gardes and Bruns1993) and ITS4 (White et al. Reference White, Bruns, Lee, Taylor, Innis, Gelfand, Sninsky and White1990), mtSSU using primers mrSSU1 and mrSSU3R (Zoller et al. Reference Zoller, Scheidegger and Sperisen1999), and nuLSU rDNA using primers AL2R (Mangold et al. Reference Mangold, Martín, Lücking and Lumbsch2008) and LR6 (Vilgalys & Hester Reference Vilgalys and Hester1990). PCR products were sequenced with the same primers used for amplification on an ABI Prism 3730 DNA Analyzer (Applied Biosystems, Waltham, MA, USA) at the Pritzker Laboratory for Molecular Systematics and Evolution at The Field Museum, Chicago, Illinois, USA.
Sequence alignment and phylogenetic analysis
ITS, mtSSU and nuLSU sequences were aligned independently using the ‘auto’ option with the FFT-NS-i algorithm in MAFFT v. 7 (Katoh et al. Reference Katoh, Rozewicki and Yamada2019), with the remaining parameters set to default values. SequenceMatrix software (Vaidya et al. Reference Vaidya, Lohman and Meier2011) was used to concatenate all three alignments. Phylogenetic analyses were performed using maximum likelihood (ML) and Bayesian analyses (BA). ML trees were calculated with RAxML-HPC2 on XSEDE v. 8.2.10 (Stamatakis Reference Stamatakis2014) on the Cipres Science Gateway (http://www.phylo.org) (Miller et al. Reference Miller, Pfeiffer and Schwartz2010) using the GTR + G + I substitution model with 1000 bootstrap pseudoreplicates, and the data partitioned by loci. For the BA, substitution models for each locus were estimated using jModelTest v. 2.1.9 (Guindon & Gascuel Reference Guindon and Gascuel2003; Darriba et al. Reference Darriba, Taboada, Doallo and Posada2012), which recommended the TIM2ef + I + G model for the ITS and nuLSU loci, and the F81 + I model for the mtSSU locus. Since the TIM2ef substitution models are not implemented in MrBayes, they were replaced by the GTR model (Ronquist & Huelsenbeck Reference Ronquist and Huelsenbeck2003). The proportion of invariable sites (I) and gamma distributed rates (G) defined in jModelTest were conserved in both cases. Two parallel Markov chain Monte Carlo (MCMC) runs were performed in MrBayes v. 3.2.6 (Huelsenbeck & Ronquist Reference Huelsenbeck and Ronquist2001; Ronquist & Huelsenbeck Reference Ronquist and Huelsenbeck2003), each using 10 000 000 generations which were sampled every 100 steps. A 50% majority-rule consensus tree was generated from the combined sampled trees (150 002) of both runs after discarding the first 25% of trees as burn-in. The convergence diagnostic of the potential scale reduction factor (PSRF) was close to 1.0 for all parameters, and the average deviation of split frequencies was below 0.01 (Gelman & Rubin Reference Gelman and Rubin1992). Tree files were visualized with FigTree v. 1.4.2 (Rambaut Reference Rambaut2014). The ITS, mtSSU and nuLSU sequences are deposited in GenBank (see Supplementary Material Table S1, available online).
Results and Discussion
Phylogenetic analysis
The phylogenetic analyses of the concatenated dataset under maximum likelihood (ML) and the Bayesian analysis (BA) were congruent, and therefore only the ML tree is shown here with the BA support values added. Our new material clusters in 37 lineages, including three distinct lineages that are described as new species below (Fig. 1). In addition, our data suggest that the distinction of Xanthoparmelia capensis Hale and X. tinctina (Maheu & Gillet) Hale requires further study since the species do not form separate monophyletic groups (Figs 1 & 2). The two species are not separated but there is some geographical pattern that suggests more than one species might be present in the complex.
Figure 1. Phylogenetic relationships of Xanthoparmelia based on concatenated alignments of ITS, mtSSU and nuLSU markers. Topology based on maximum likelihood (ML) analyses. Bootstrap values > 75 and posterior probabilities > 0.95 are indicated on each branch. The 61 samples from South Africa generated in this study are indicated in bold. Highlighted in bold and enlarged font at the tree tips are 37 lineages, including XV major groups (well-supported lineages indicated with grey boxes, poorly-supported lineages with white boxes). Information provided includes the collector, specimen voucher, species name and country. Further information regarding these samples and those of the collapsed nodes (Holarctic Clade, Australian Clades A-D and pulla Clade) can be found in Supplementary Materials S1 (available online).
Figure 2. New species and probable synonyms of Xanthoparmelia in South Africa. A, habitus of X. nimisii sp. nov. (B, holotype). B, habitus of X. pseudochalybaeizans sp. nov. (B, holotype). C, habitus of X. sipmaniana sp. nov. (B, holotype). D, habitus of X. capensis (B 60 0201801). Scales: A–D = 5 mm. In colour online.
Lineages found in the South African material
The relationships of the 37 lineages (indicated with italicised numbers in parentheses) identified in the newly sequenced material cluster into 15 major groups (I–XV), which are briefly discussed below. Within Fig. 1, only these 37 lineages are indicated: the well-supported lineages are indicated by grey boxes, while white boxes represent clades that lack support. More studies will be necessary to understand the relationships and delimitation of several species mentioned in this study.
Clade I: (1) We interpret the samples in this clade as representing Xanthoparmelia chalybaeizans (J. Steiner & Zahlbr.) Hale s. str. since our new specimens are phenotypically identical to the type of X. chalybaeizans. Xanthoparmelia chalybaeizans and (2) X. bibax (Brusse) Hale contain salazinic and chalybaeizanic (trace) acids, but X. chalybaeizans lacks pruina on the lobes. Our sample of X. bibax (B 60 0198686) clusters with MAF-Lich 15530-2 identified as X. chalybaeizans in Amo de Paz et al. (Reference Amo de Paz, Lumbsch, Cubas, Elix and Crespo2010) and Leavitt et al. (Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018), and is interpreted here as belonging to X. bibax. (3) Some of our samples of X. ralla (Brusse) G. Amo et al. (B 60 0201813, B 60 0208820) are closely related to X. subamplexuloides Hale. The latter species differs from X. ralla by its larger foliose thallus (4–7 cm) and lobes (0.4–1.3 mm), the presence of isidia and lack of pycnidia and apothecia. However, the added specimens (B 60 0201813, B 60 0208820) did not cluster with the other X. ralla from South Africa (MAF-Lich 15537/15529) (Leavitt et al. Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018). The polyphyly of X. ralla requires additional sampling to clarify the species delimitation within this complex of subcrustose species. (4) Xanthoparmelia subchalybaeizans (Hale) G. Amo et al. is found to consist of at least two separate clades: X. subchalybaeizans MAF-LICH 16443 and X. subchalybaeizans MAF-LICH 15532 (Amo de Paz et al. Reference Amo de Paz, Lumbsch, Cubas, Elix and Crespo2010; Leavitt et al. Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018). Our additional specimen (B 60 0198685) is closely related to MAF-LICH 15531-1, 15531-2 and 15534, but does not form a monophyletic group with the other X. subchalybaeizans specimens. (5) Xanthoparmelia prolata (Hale) Elix is closely related to X. aff. tortula (MAF-LICH 16446), which has shorter and more adnate lobes and produces norlobaridone. (6) Xanthoparmelia tortula (Kurok.) Elix has previously been found to be closely related to X. neotumidosa Hale and X. brachinaensis (Elix) O. Blanco et al. (Amo de Paz et al. Reference Amo de Paz, Lumbsch, Cubas, Elix and Crespo2010; Leavitt et al. Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018). Our additional specimen of X. tortula (B 60 0198854) does not cluster with the other X. aff. tortula from South Africa (MAF-LICH 16446).
Clade II: (7) Xanthoparmelia sipmaniana Barcenas-Peña, Lumbsch & Grewe sp. nov. and (8) X. nimisii Barcenas-Peña, Lumbsch & Grewe sp. nov. (Clade III) will be discussed under ‘Taxonomy’.
Clade III: (9) Xanthoparmelia toninioides Hale is supported as a monophyletic clade. (10) Xanthoparmelia aff. perspersa (Stizenb.) G. Amo et al. (B 60 0198690) agrees phenotypically with X. perspersa but does not form a monophyletic clade. Instead, it clusters together with X. aff. perspersa ‘3’ (MAF-Lich 15539) of Leavitt et al. (Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018). However, this relationship lacks support. (11) Xanthoparmelia prodomokosii Hale et al. is closely related to (12) X. contrasta Hale. Both species have a similar chemistry but X. contrasta differs from X. prodomokosii by having a black lower surface and a lack of pycnidia and apothecia. (13) Xanthoparmelia karooensis Hale is a well-supported clade. (14) Xanthoparmelia subdomokosii (Hale) Hale does not cluster with the other X. subdomokosii specimen MAF-Lich 17187, which was also collected in South Africa. However, both samples are in the same clade as X. karooensis. Both species differ only slightly in X. subdomokosii having a wider thallus (3–12 cm) and lobes (2–4 mm) than X. karooensis (4–7 cm and 0.8–1.5 mm). Thus, the three samples could belong to the same species and more studies are necessary. (15) Xanthoparmelia substenophylloides Hale is closely related to X. aff. plittii from Kenya, a similar species that differs in having a larger thallus (4–10 cm) and contiguous to imbricate lobes (1–2 mm).
Clade IV: (16) Xanthoparmelia marroninipuncta (Brusse) Hale is closely related to X. austroafricana (Stirt.) Hale, which produces protocetraric and usnic acids and is also restricted to southern Africa. However, X. austroafricana has a white maculate upper surface and broader thallus (4–12 cm) than X. marroninipuncta (3–8 cm). Our study supports the monophyly of (17) X. endomiltoides (Nyl.) Hale. (18) Xanthoparmelia cf. iniquita does not match completely with the X. iniquita of Elix et al. (Reference Elix, Johnston and Armstrong1986), differing in having an adnate thallus and sublinear and broader lobes (up 4 mm). Whether X. cf. iniquita represents an undescribed taxon requires additional studies.
Clade V: (19) The distinction of Xanthoparmelia capensis and X. tinctina requires further study. The two species are similar in their morphology and chemistry, and also have overlapping geographical distributions (Hale Reference Hale1986, Reference Hale1990). Our analysis shows that the two newly sequenced samples identified as X. capensis (see Fig. 2D) fall into a clade with X. tinctina. We refrain from synonymizing the two species here since there is geographical structure in the phylogenetic tree and additional sampling might show that more than one species is involved in this clade.
Clade VI: (20) Xanthoparmelia ceresina (Vain.) Hale is closely related to X. subramigera (Gyeln.) Hale. Both species contain fumarprotocetraric and succinprotocetraric acids. However, X. ceresina has a very adnate and narrow thallus (3–8 cm) and lobes (0.8–2 mm), and lacks maculae. (21) Xanthoparmelia phaeophana (Stirt.) Hale is closely related to (22) X. leonora (A. Massal.) Hale. Both species have a white maculate upper surface and contain fumarprotocetraric and succinprotocetraric acids, but X. phaeophana can also contain protocetraric (±), physodalic (±), virensic (±) and caperatic (±) acids. In addition, X. leonora differs from X. phaeophana by its terricolous thallus, weakly convoluted lobes and less distinctive maculae (Hale Reference Hale1990). In other studies (Leavitt et al. Reference Leavitt, Kirika, Amo de Paz, Huang, Hur, Elix, Grewe, Divakar and Lumbsch2018), X. phaeophana was closely related to X. aff. krogiae Hale & Elix.
Clade VII: (23) Xanthoparmelia pseudochalybaeizans Barcenas-Peña, Lumbsch & Grewe sp. nov. is closely related to (24) X. amphixanthoides (J. Steiner & Zahlbr.) Hale. The former species differs in having an adnate, saxicolous thallus, wider lobules (1–3 mm), not being terete, and the presence of chalybaeizanic acid (see also ‘Taxonomy’ below).
Clade VIII: (25) Xanthoparmelia squamariatella (Elix) O. Blanco et al. includes distinct chemical components: B 60 0198388 contains norstictic, connorstictic and salazinic acids, whereas B 60 0201770 and B 60 0198392 contain fumarprotocetraric, protocetraric and succinprotrotocetraric acids. Additional studies are needed to understand their relationship.
Clade IX: (26) Xanthoparmelia caliginosa (Essl.) O. Blanco et al. is sister to X. pseudoglabrans (Essl.) O. Blanco et al., which is also endemic to southern Africa. The two species differ because X. pseudoglabrans has lobes that are often somewhat maculate, a black lower surface, and contains alectoronic and α-collatolic acids, whereas X. caliginosa lacks maculae, has a dark brown to black lower surface, and contains olivetoric acid. (27) Xanthoparmelia cafferensis (Essl.) O. Blanco et al. clusters (without support) with the similar X. verrucella (Essl.) O. Blanco et al., which occurs in southern Africa and Australasia (Culberson et al. Reference Culberson, Culberson and Esslinger1977). (28) Xanthoparmelia glabrans (Nyl.) O. Blanco et al. from southern Africa clusters separately from X. glabrans from Europe, South America and Australia (collapsed clades: Australian Clade C and pulla Clade). In contrast, X. glabrans clustered in an unsupported clade with X. imitatrix (Taylor) O. Blanco et al. from southern Africa. Both species are morphologically similar but X. imitatrix has a different chemistry (physodic acid and often trace amounts of 4-O-methylphysodic acid) (Esslinger Reference Esslinger1977, Reference Esslinger2000; Elix Reference Elix1994).
Clade X: (29) Xanthoparmelia xanthomelanella Elix forms a monophyletic clade.
Clade XI: (30) Xanthoparmelia perplexa (Stizenb.) Hale forms a monophyletic clade.
Clade XII: (31) Xanthoparmelia greytonensis Hale and (32) X. xanthomelaena (Müll. Arg.) Hale cluster together but this relationship lacks support. Note that X. aff. greytonensis from South Africa (MAF-LICH 16210) is only distantly related to our X. greytonensis material. Xanthoparmelia xanthomelaena also shows differences among specimens from different continents. The southern African material does not cluster with material from Australia (MAF-LICH 16447).
Clade XIII: (33) Xanthoparmelia waboomsbergensis Elix is related to X. tegeta Elix & J. Johnst. However, X. tegeta is readily distinguished by its loosely adnate, pulvinate, dark yellowish green and larger thallus (6–9 cm), and contiguous to imbricate lobes (0.7–1.5 mm) (Elix et al. Reference Elix, Johnston and Armstrong1986; Hale Reference Hale1990).
Clade XIV: (34) Xanthoparmelia molliuscula (Ach.) Hale forms a monophyletic clade. Clade XV: The southern African material identified here as (35) X. psoromica Hale is morphologically similar but differs in chemical constituents, with (36) X. colorata (Gyeln) Hale containing salazinic and norstictic acids, and X. psoromica containing psoromic acid. Both species are members of a group of species with the schenckiana pigment that also includes X. colorata and X. schenckiana (Müll. Arg.) Hale. In our analysis, X. colorata and X. psoromica cluster in the same clade, whereas (37) X. schenckiana is only distantly related.
Taxonomy
Xanthoparmelia nimisii Barcenas-Peña, Lumbsch & Grewe sp. nov.
MycoBank No.: MB 847480
Differs from Xanthoparmelia annexa by the presence of soredia and the broader lobes (5–6 mm). In addition, the new species forms a well-supported clade based on a dataset of ITS, mtSSU and nuLSU sequences.
Type: South Africa, Western Cape, Cape Winelands, Breede River DC, Klein Cedarberg, on sunny siliceous rock, 985 m alt., 32°55.4ʹS, 19°30.55ʹE, 24 September 2014, V. Wirth B 60 0198683 (B—holotype). GenBank Accession nos: OQ356389 (ITS) and OQ366463 (mtSSU).
Thallus foliose, tightly to loosely adnate, 4–5 cm diam., lobate; lobes subirregular, elongate, plane, separate, contiguous, 5–6 mm wide, not lobulate; apices subrotund, smooth, eciliate; upper surface yellow-green, smooth, shiny, epruinose and emaculate, not isidiate; soralia laminal and orbicular to irregular masses, soredia granular; medulla white, with continuous algal layer; lower surface black, plane, moderately rhizinate; rhizines black, simple, 0.3–0.4 mm long.
Apothecia not observed; pycnidia not observed.
Chemistry
Upper cortex K+ yellow, UV−; medulla K−, C+ deep red, KC+ red, P−. Atranorin and lecanoric acid.
Etymology
The new species is named in honour of Pier Luigi Nimis for his successful career devoted to the study of lichens.
Distribution and habitat
South Africa, on siliceous rocks.
Remarks
The new species resembles X. annexa because both species share the same chemistry. However, X. annexa is an isidiate species, it does not have soredia and has narrower lobes (0.7–3.5 mm) than the new species (5–6 mm), which is sorediate (Hale & Kurokawa Reference Hale and Kurokawa1964).
Additional specimen examined
South Africa: Western Cape: Klein Cedarberg, on shaded vertical cliff faces, siliceous rock, 985 m alt., 32°55ʹ08ʺS, 19°30ʹ26ʺE, 2014, V. Wirth 36287 (STU).
Xanthoparmelia pseudochalybaeizans Barcenas-Peña, Lumbsch & Grewe sp. nov.
MycoBank No.: MB 847482
Same chemistry and similar morphology to Xanthoparmelia chalybaeizans, but differs by not forming an imbricate thallus, and having a white medulla. Its distinction is supported by a phylogenetic study based on a concatenated dataset of ITS, mtSSU and nuLSU rDNA.
Type: South Africa, Western Cape, Overberg, 32 km NE of Bredasdorp, Renosterveld Nature Reserve, on rock, 190 m alt., 34°21.18ʹS, 20°19.05ʹE, 13 October 2015, V. Wirth RENO-2, B 60 0201783 (B—holotype). GenBank Accession nos: OQ356405 (ITS), OQ366468 (mtSSU) and OQ366517 (nuLSU).
(Figs 1 (Clade VII (23)) & 2B)
Thallus foliose, adnate, (1.8)3–7 cm diam.; lobes subirregular, elongate, plane, contiguous, 1–3 mm wide, not lobulate, apices rotund, smooth, eciliate; upper surface yellow-green, smooth, shiny, epruinose and emaculate, isidia and soredia lacking; medulla white, with continuous algal layer; lower surface brown, plane, moderately rhizinate; rhizines brown to dark brown, simple, 0.5–0.7 mm long.
Apothecia substipitate, with a brown to dark brown disc, not pruinose, (0.5)2–6 mm diam.; spores 10–12 × 5–7 μm.
Pycnidia common; conidia bifusiform, 5–6 × 0.5 μm.
Chemistry
Upper cortex K−, UV−; medulla K+ yellow to dark red, C−, KC−, P+ orange. Usnic, norstictic (trace), chalybaeizanic, salazinic, consalazinic acids, and an unknown substance (R f 15/norstictic 30).
Etymology
The new species is named for its close similarity to X. chalybaeizans.
Distribution and habitat
South Africa, on siliceous rocks.
Remarks
This new species has the same chemistry (salazinic, consalazinic, norstictic (±trace), usnic and chalybaeizanic acids) and a similar morphology to X. chalybaeizans (Hale Reference Hale1990). However, X. chalybaeizans has a thallus that is 4–8 cm diam., with contiguous to imbricate lobes, that are 1–3 mm wide, and a white to slightly yellow medulla. In contrast, X. pseudochalybaeizans has no imbricate lobes and a white medulla. Furthermore, the distinction of X. chalybaeizans and X. pseudochalybaeizans is supported by the phylogenetic study based on ITS, mtSSU and nuLSU rDNA sequence data.
Additional specimens examined
South Africa: Western Cape: Overberg, 32 km NE of Bredasdorp, Renosterveld Nature Reserve, on rock, 185 m alt., 34°21.25ʹS, 20°19.03ʹE, 2015, V. Wirth B 60 0201774 (B); ibid., 190 m alt., 34°21.21ʹS, 20°19.017ʹE, V. Wirth B 60 0201772 (B).
Xanthoparmelia sipmaniana Barcenas-Peña, Lumbsch & Grewe sp. nov.
MycoBank No.: MB 847483
Morphologically and chemically similar to X. hypoprotocetrarica (Kurok. & Elix) Hale but differs by its phylogenetic relationships based on ITS, mtSSU and nuLSU rDNA sequence data. In addition, both differ in their geographical distribution (Australia vs South Africa).
Type: South Africa, Western Cape, Cape Winelands, Breede River DC, Klein Cedarberg, on sunny siliceous rock, 985 m alt., 32°55.4ʹS, 19°30.55ʹE, 24 September 2014, V. Wirth B 60 0198688 (B—holotype). GenBank Accession nos: OQ356383 (ITS) and OQ366502 (nuLSU).
Thallus foliose, adnate to loosely attached, 7–9 cm diam.; lobes subirregular, elongate, plane, imbricate, subascending, 1–3 mm wide, lobulated margins, apices rotund, smooth, eciliate; upper surface yellow-green, smooth, shiny, epruinose, effigurate maculate, isidia and soredia lacking; medulla white, with continuous algal layer; lower surface black, plane, sparsely rhizinate; rhizines black, simple to furcate, 0.5–0.7 mm long.
Apothecia substipitate, with a dark brown disc, not pruinose, 2–5 mm diam.; spores 8–11 × 4–6 μm.
Pycnidia common; conidia bifusiform, 5–6 × 0.5 μm.
Chemistry
Upper cortex K−, UV−; medulla K−, C−, KC−, P−. Usnic and hypoprotocetraric acids.
Etymology
The new species is named in honour of Harrie Sipman for his important contribution to lichenology.
Remarks
Xanthoparmelia sipmaniana and X. hypoprotocetrarica both share an identical morphology and chemistry and can be interpreted as cryptic species, as indicated by molecular data. The two species exhibit different distributional ranges, with X. hypoprotocetrarica occurring in Australia, and X. sipmaniana currently known only from South Africa (Kurokawa & Elix Reference Kurokawa and Elix1971; Hale Reference Hale1974, Reference Hale1990).
Acknowledgements
We are grateful to Robert Lücking (Berlin) for organizing the loan of specimens from B to F and Holger Thüs (Stuttgart) for examining some of specimens in STU. With regards to the fieldwork in South Africa, Volkmar Wirth acknowledges the permission from Werner Wullschleger to collect on his property at Klein-Cederberg and the collecting permit organized by Odette Curtis-Scott, who was also great company during the fieldwork in the Haarwegskloof Renosterveld Reserve.
Author ORCIDs
Alejandrina Barcenas-Peña, 0000-0003-1674-1164; Harrie J. M. Sipman, 0000-0002-6224-3513; Volkmar Wirth, 0000-0002-4982-9472; Felix Grewe, 0000-0002-2805-5930; H. Thorsten Lumbsch, 0000-0003-1512-835X.
Competing Interests
The authors declare none.
Supplementary Material
The Supplementary Material for this article can be found at https://doi.org/10.1017/S0024282923000300.
