See discussions, stats, and author profiles for this publication at: https://www.researchgate.net/publication/47414491 Glomus africanum and G. iranicum, two new species of arbuscular mycorrhizal fungi (Glomeromycota) ARTICLE in MYCOLOGIA · JUNE 2010 Impact Factor: 2.47 · DOI: 10.3852/09-302 · Source: PubMed CITATIONS READS 22 99 7 AUTHORS, INCLUDING: Janusz Błaszkowski Gábor M Kovács 47 PUBLICATIONS 434 CITATIONS 61 PUBLICATIONS 1,412 CITATIONS West Pomeranian University of Technology, … SEE PROFILE Eötvös Loránd University SEE PROFILE Elzbieta Orlowska Francois Buscot 22 PUBLICATIONS 351 CITATIONS 190 PUBLICATIONS 3,769 CITATIONS Aarhus University SEE PROFILE All in-text references underlined in blue are linked to publications on ResearchGate, letting you access and read them immediately. Helmholtz-Zentrum für Umweltforschung SEE PROFILE Available from: Mehdi Sadravi Retrieved on: 04 February 2016 Mycologia, 102(6), 2010, pp. 1450–1462. DOI: 10.3852/09-302 # 2010 by The Mycological Society of America, Lawrence, KS 66044-8897 Glomus africanum and G. iranicum, two new species of arbuscular mycorrhizal fungi (Glomeromycota) Janusz Błaszkowski1 red in Melzer’s reagent. In the field G. africanum was associated with roots of five plant species and an unrecognized shrub colonizing maritime sand dunes of two countries in Europe and two in Africa, and G. iranicum was associated with Triticum aestivum cultivated in southwestern Iran. In one-species cultures with Plantago lanceolata as the host plant G. africanum and G. iranicum formed arbuscular mycorrhizae. Phylogenetic analyses of partial SSU sequences of nrDNA placed the two new species in Glomus group A. Both species were distinctly separated from sequences of described Glomus species. Key words: arbuscular fungi, Glomeromycota, molecular phylogeny, mycorrhizae, new species Department of Plant Protection, West Pomeranian University of Technology, Szczecin, Słowackiego 17, PL-71434 Szczecin, Poland Gábor M. Kovács Department of Plant Anatomy, Institute of Biology, Eötvös Loránd University, Pázmány Péter sétány 1/C, 1117 Budapest, Hungary Tı́mea K. Balázs Institute of Ecology and Botany, Hungarian Academy of Sciences, Alkotmány street 2–4, 2163 Vácrátót, Hungary Elz_bieta Orłowska Institute of Molecular Biology, University of Aarhus, Gustav Wieds Vej 10 C, 8000 Aarhus C Denmark Mehdi Sadravi INTRODUCTION Department of Plant Protection, Faculty of Agriculture, Yasouj University, Daneshju Avenue, P.O. Box 353, 75918–74831 Yasouj, Iran Arbuscular mycorrhizal fungi (AMF) of phylum Glomeromycota are the most common soil fungi in the world coexisting symbiotically with ca. 70–90% of land plants (Wang and Qiu 2006, Smith and Read 2008, Brundrett 2009). Maritime sand dunes favor AMF development (Koske 1987, Dalpé 1989, Tadych and Błaszkowski 2000) because of low nutrient and organic matter content (Nicolson and Johnston 1979, Koske 1988), as well as the absence of numerous antagonistic microorganisms, especially parasites of AMF (Koske et al. 2004). Of the ca. 220 described species of Glomeromycota, at least 35 originally were isolated from maritime dunes and many others have been associated with roots of dune plants (Sridhar and Beena 2001, www. agro.ar.szczecin.pl/,jblaszkowski/). Members of Glomeromycota also commonly co-occur with cultivated plants, including Triticum aestivum L. that usually has harbored abundant and diverse spore populations of these fungi (Hetrick and Bloom 1983, Dodd and Jeffries 1989, Błaszkowski 1993). AMF sequences amplified from root samples however suggest that the number of existing species of AMF is much higher than that formally described and that most undescribed species belong to genus Glomus (Helgason et al. 2002, Fitter 2005, Hijri et al. 2006, Kovács et al. 2007, Öpik et al. 2009), especially in Glomus group A sensu Schwarzot et al. (2001). Possible causes for the omission of these undescribed species from the scientific record might be due to (i) a lack of or rare sampling of AMF in many terrestrial regions of Earth, (ii) the few specialized and Tesfaye Wubet François Buscot UFZ, Helmholtz Centre for Environmental Research, Theodor-Lieser-Straße 4, 06120 Halle-Saale, Germany Abstract: Two new arbuscular mycorrhizal fungal species (Glomeromycota) of genus Glomus, G. africanum and G. iranicum, are described and illustrated. Both species formed spores in loose clusters and singly in soil and G. iranicum sometimes inside roots. G. africanum spores are pale yellow to brownish yellow, globose to subglobose, (60–)87(–125) mm diam, sometimes ovoid to irregular, 80–110 3 90– 140 mm. The spore wall consists of a semipermanent, hyaline, outer layer and a laminate, smooth, pale yellow to brownish yellow, inner layer, which always is markedly thinner than the outer layer. G. iranicum spores are hyaline to pastel yellow, globose to subglobose, (13–)40(–56) mm diam, rarely eggshaped, prolate to irregular, 39–54 3 48–65 mm. The spore wall consists of three smooth layers: one mucilaginous, short-lived, hyaline, outermost; one permanent, semirigid, hyaline, middle; and one laminate, hyaline to pastel yellow, innermost. Only the outermost spore wall layer of G. iranicum stains Submitted 3 Dec 2009; accepted for publication 14 Apr 2010. 1 Corresponding author. Department of Plant Protection, West Pomeranian University of Technology, Szczecin, Słowackiego 17, PL 71434 Szczecin, Poland. E-mail: janusz.blaszkowski@zut.edu.pl 1450 BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. experienced mycologists that study taxonomy of Glomeromycota, and (iii) seasonal, rare or lack of sporulation by many AMF in the field (Gemma et al. 1989, Stürmer and Bellei 1994, Stutz and Morton 1996). Often the diversity of AMF spores obtained from environmental samples can be increased with successive (Stutz and Morton 1996) or long-term (Oehl et al. 2004) pot trap cultures. Examination of long-term trap cultures with rhizosphere soils and roots of plant species collected from maritime sand dunes of Africa and Europe and from T. aestivum cultivated in southwestern Iran revealed spores of two undescribed species of Glomeromycota forming glomoid spores. Phylogenetic analyses of sequences of rDNA placed the fungi in Glomus group A sensu Schwarzot et al. (2001) and confirmed their uniqueness relative to other known Glomus species. The fungi are described here as G. africanum sp. nov. and G. iranicum sp. nov. MATERIALS AND METHODS Establishment and growth of trap and single-species cultures, extraction of spores and staining of mycorrhizae.—Spores examined in this study were derived from both pot trap and single-species cultures. Trap cultures were established to obtain a large number of living spores and to initiate sporulation of species that were present but were not detected in field collections (Stutz and Morton 1996). The method used to establish trap cultures, growing conditions and the methods of spore extraction and staining of mycorrhizae were those described by Błaszkowski et al. (2006). Single-species cultures also were established and grown as described by Błaszkowski et al. (2006), with three exceptions. First, cultures of both species were successfully established from small clusters of spores. The clusters consisted of 2–3 (G. africanum) or 10 (G. irranicum) spores attached by a common mycelium. To prevent contamination by fragments of hyphae of other AMF the clusters were rinsed several times with water; each time the water was removed with a pipette. Second, instead of marine sand, the growing medium was an autoclaved commercially available coarse-grained sand (grains 1.0–10.0 mm diam, 80.50%; grains 0.1–1.0 mm diam, 17.28%; grains , 0.1 mm diam, 2.22%) mixed (5 : 1, v/v) with clinopthilolite (Zeocem, Bystré, Slovakia) of grains 2.5–5 mm. Clinopthilolite is a crystaline hydrated alumosilicate of alkali metals and alkaline earth metals having a high ion exchange and water-holding capacity. The pH of the sand-clinopthilolite mixture was 7.3. Third, the cultures were kept in transparent plastic bags, 15 cm wide and 22 cm high, as suggested by Walker and Vestberg (1994), instead of open pot cultures (Gilmore 1968). To prevent contamination of cultures with other AMF but still allow gas exchange an opening of about 1 cm2 was left in the upper part of each bag while the edges were sealed with plastic clips. The cultures were watered with tap water once a week and harvested after 5 mo to 1451 extract spores. Root fragments located ca. 1–5 cm below the upper level of the growing medium were cut off with a scalpel to reveal mycorrhizal structures. Plantago lanceolata L. was used as a host plant in both trap and single-species cultures. Microscopy survey.—Morphological properties of spores and wall structure were determined based on examination of at least 100 spores mounted in water, lactic acid, polyvinyl alcohol/lactic acid/glycerol (PVLG, Omar et al. 1979) and a mixture of PVLG and Melzer’s reagent (1 : 1, v/v). Spores at all developmental stages were crushed to varying degrees by applying pressure to the cover slip and then stored at 65 C for 24 h to clear contents from oil droplets. They were examined under an Olympus BX 50 compound microscope equipped with Nomarski differential interference contrast optics. Microphotographs were recorded on a Sony 3CDD color video camera coupled to the microscope. Terminology of spore structure is that suggested by Stürmer and Morton (1997) and Walker (1983). Spore color was examined under a dissecting microscope on fresh specimens immersed in water. Color names are from Kornerup and Wanscher (1983). Nomenclature of fungi and plants is that of Walker and Trappe (1993) and Mirek et al. (1995), respectively. The authors of the fungal names are those presented at the Index Fungorum Website http:// www.indexfungorum.org/AuthorsOfFungalNames.htm. Voucher specimens were mounted in PVLG and a mixture of PVLG and Melzer’s reagent (1 : 1, v/v) on slides and deposited in the Department of Plant Protection (DPP), West Pomeranian University of Technology, Szczecin, Poland, and in the herbarium at Oregon State University (OSC) in Corvallis, Oregon. Color microphotographs of spores of the new species can be viewed at the URL http:// www.agro.ar.szczecin.pl/,jblaszkowski/. DNA extraction, polymerase chain reaction and DNA sequencing.—Several spores or small spore clusters were used to obtain target DNA as described by Błaszkowski et al. (2009). We used a nested PCR to amplify a segment of SSU of the nrDNA. The GlomerWT0 and Glomer1536 primers were used in the first PCR as described by Wubet et al. (2006). In the second step we used the AML1 and AML2 primers as described by Lee et al. (2008). A high fidelity enzyme mix (MBI Fermentas, Vilnius, Lithuania) was used for PCR. With these primer combinations we obtained sequences longer than 700 nucleotides, which are suitable for reliable phylogenetic analyses of Glomus groups. Because the amplified region overlaps the segment amplified with AM1 (Helgason et al. 1998) and NS31 (Simon et al. 1992), primers used most widely in AMF diversity studies, we could compare the sequences of the new taxa with numerous environmental AMF sequences available in public databases. The appropriate size amplicons were cleaned and either cloned into a pGEMT-easy vector (Promega, Madison, Wisconsin) and transformed into competent JM109 Escherichia coli (Promega, Madison, Wisconsin) or cloned with the TOPO TA CloningH Kit (Invitrogen) and transformed into TOP10 chemically competent E. coli strains (Invitrogen) following manufacturers’ instructions. Ten positive clones from both species 1452 MYCOLOGIA were sequenced in both directions with universal primers and an ABI PRISM 3.1 BigDye Terminator 3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, California). Electrophoreses were carried out on an ABI PRISM 3100 or 3730XL Genetic Analyzer (Applied Biosystems, Foster City, California). The electrophoregrams were processed with Pregap4 1.4b1 and Gap 4.8b1 programs of the Staden Program Package (Staden et al. 2000). Nonredundant sequences of clones were deposited in GenBank (HM153415-HM153424). Phylogenetic analyses.—After pilot analyses of the sequences together with identified species of the phylum Glomeromycota the final analyses were carried out with a dataset of known Glomus group A sequences and unidentified AMF sequences from in planta studies including the most similar sequences to our clones obtained from BLAST queries. We used Glomus lamellosum as outgroup. Only the two most distant sequences of both new taxa were included in the analyses. The sequences were aligned with Multalin (Corpet 1988, http://prodes.toulouse.inra.fr/multalin/multalin. html) and manually edited with ProSeq 2.9 (Filatov 2002). The best fit nucleotide substitution model was selected with the program jModelTest (Posada 2008) considering the selection of Akaike information criterion (AIC). The model and the parameters were used to calculate distances for neighbor-joining analyses with PAUP*4.0b10 software (Swofford 2003). Support of branches was tested by bootstrap analysis with 1000 replicates. A maximum likelihood (ML) phylogenetic analysis was carried out with the online version of PHYML 3.0 (Guindon and Gascuel 2003). The GTR nucleotide substitution model was used with ML estimation of base frequencies. The proportion of invariable sites was estimated and optimized. Six substitution rate categories were set, and the gamma distribution parameter was estimated and optimized. A bootstrap analysis with 1000 replicates also was used here to test support of branches. The same substitution model was used in Bayesian analyses performed with MrBayes 3.1 (Huelsenbeck and Ronquist 2001, Ronquist and Huelsenbeck 2003) with the Computational Biology Service Unit, Cornell University (http://cbsuapps.tc.cornell.edu/index.aspx). The Markov chain was run 5 000 000 generations, sampling in every 100 steps, and with a burn-in at 7500 sampled trees. The alignment was deposited in TreeBase (http://purl. org/phylo/treebase/phylows/study/TB2:S10461). Phylogenetic trees were viewed and edited by Tree Explorer of the MEGA 4.0 program (Tamura et al. 2007) and a text editor. TAXONOMY Glomus africanum Błaszk. & Kovács sp. nov. FIGS. 1–13 MycoBank MB518241 Sporocarpia ignota. Sporae singulatim vel gregatim in solo efformatae. Fascicula 470–620 3 600–1250 mm, e sporis 2–6. Sporae pallide luteae vel spadiceae; globosae vel subglobosae; (60–)87(–125) mm diam; raro ovoideae, oblongae vel irregulares; 80–110 3 90–140 mm. Tunica sporae stratis duobus (strata 1 ad 2); stratum ‘‘1’’ caducum, glabrum, hyalinum, (1.5–)3.1(–8.6) mm crassum; stratum ‘‘2’’ laminatum, glabrum, pallide luteum vel spadiceum, (1.0–)1.7(–2.7) mm crassum. Hypha sporifera pallide lutea vel spadicea; recta vel recurvta; cylindrica vel infundibuliformis; (3.7–)5.7(–9.3) mm lata ad basim sporae; pariete pallide luteo vel spadiceo, (2.9–)4.4(–5.4) mm crasso, stratis 1 ad 2 in parietem sporae continuantibus. Porus hyphae (1.0–)2.1(–2.9) diam. Mycorrhizas vesiculo-arbusculares formans. Typus: Polonia: Sedinum (Szczecin), infra P. lanceolata, 10 Mar 2008, J. Błaszkowski, 3167 (Holotypus, DPP). Sporocarps unknown. Spores formed in loose clusters or singly in the soil (FIGS. 1, 2) develop blastically at the tip of sporogenous hyphae either branched from a parent hypha continuous with a mycorrhizal extraradical hypha (spores in clusters) or directly developed from mycorrhizal extraradical hyphae (single spores). Clusters 470–620 3 600–1250 mm with 2–6 spores (FIG. 1). Spores pale yellow (4A3) to brownish yellow (5C8); globose to subglobose, (60–)87(–125) mm diam, sometimes ovoid to irregular; 80–110 3 90–140 mm; with one subtending hypha (FIGS. 1–3, 7 and 8). Spore wall composed of two layers (FIGS. 3–5, 7, 8). Layer 1, forming the surface, semipermanent, evanescent, hyaline, (1.5–)3.1(–8.6) mm thick, more or less deteriorated in mature spores, infrequently completely sloughed in older specimens; in young and freshly matured spores, the upper surface of this layer usually is covered with irregular blister-like outgrowths, rarely is smooth (FIGS. 3–8). Layer 2-laminate, smooth, pale yellow (4A3) to brownish yellow (5C8), (1.0–)1.7(–2.7) mm thick (FIGS. 3–5, 7, 8). Layers 1 and 2 do not stain in Melzer’s reagent. Subtending hypha pale yellow (4A3) to brownish yellow (5C8); straight or recurved, flared to slightly funnel-shaped, sometimes slightly constricted at spore base; (3.7–)5.7(–9.3) mm wide at the spore base (FIGS. 1–3, 7, 8). Wall of subtending hypha pale yellow (4A3) to brownish yellow (5C8); (2.9–)4.4(–5.4) mm thick at spore base; continuous with spore wall layers 1 and 2; layers 1 and 2 usually extend far below spore base in mature spores (FIGS. 2, 3, 7, 8). Pore (1.0–)2.1(–2.9) mm diam, open (FIG. 7) or occluded by a curved septum continuous with some innermost laminae of spore wall layer 2 (FIG. 8). Germination unknown. Mycorrhizal associations. In the field G. aficanum was associated with roots of Ammophila arenaria (L.) Link, Cineraria geifolia L., Senecio elegans L., Thinopyrum distichum (Thunb.) A. Löve, Trachyandra divaricata ( Jacq.) Kunth and an unrecognized shrub. In one-species culture with P. lanceolata as the host plant G. africanum formed mycorrhizae with arbuscules, vesicles and intra- and extraradical hyphae (FIGS. 9–12). Arbuscules generally were dispersed widely along the root fragments examined. They BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. 1453 FIGS. 1–8. Glomus africanum. 1. Spores in loose cluster. 2. Single spore. 3–5. Spore wall layers (swl) 1–2; note the irregular blister-like outgrowths covering the upper surface of swl1 seen in a cross view. 6. Irregular outgrowths of the spore surface seen in a plan view. 7. Subtending hyphal wall layers (shwl) 1 and 2 continuous with spore wall layers (swl) 1 and 2; note the open lumen of the subtending hypha. 8. Subtending hyphal wall layers (shwl) 1 and 2 continuous with spore wall layers (swl) 1 and 2; note the septum in the lumen of the subtending hypha. 1, 2. Spores in lactic acid. 4, 6–8. Spores crushed in PVLG. 3, 5. Spores in PVLG + Melzer’s reagent. 1–8, differential interference microscopy. Bars: 1, 2 5 20 mm; 3–8 5 10 mm. 1454 MYCOLOGIA FIGS. 9–12. Mycorrhizae of Glomus africanum in roots of Plantago lanceolata stained in 0.1% trypan blue. 9. Arbuscule (arb) with trunk (t) developed from parent hypha (ph). 10. Vesicles (ves). 11. H-shaped branch (Hb). 12. Y-shaped branch (Yb). 9–12. Differential interference microscopy. Bars: 9, 11, 12 5 10 mm; 10 5 20 mm. consisted of a short trunk grown from a parent hypha and numerous branches with fine tips (FIG. 9). Vesicles were not numerous and usually highly separated. They were ellipsoidal to oblong, 10.5– 34.5 3 25.5–113.8 mm, when observed in a plan view (FIG. 10). Intraradical hyphae grew along the root axis, were (1.3–)4.6(–9.8) mm wide, straight or slightly recurved, and occasionally formed H- or Y-shaped branches and coils (FIGS. 9–12). The coils were ellipsoidal to oblong, 14.0–21.6 3 50.0–82.4.0 mm, when seen in a plan view (FIG. 11). Extraradical hyphae were (2.5–)4.1(–5.5) mm wide and occurred infrequently. In 0.1% trypan blue arbuscules stained violet white (17A2) to violet (17B6), vesicles pastel violet (17A4) to deep violet (17D8), intraradical hyphae pale violet (17A3) to violet (17B8), coils pastel violet (17A4) to deep violet (17D8), and extraradical hyphae pale violet (17A3) to deep violet (17D8; FIGS. 9–12). Phylogenetic position. Phylogenetic analyses of partial SSU sequences of nrDNA placed G. africanum unambiguously in Glomus group A sensu Schwarzott et al. (2001) within genus Glomus (FIG. 13). The sequences of the species separated unambiguously from described Glomus species of this group. Sequences of G. africanum showed high similarity to and grouped together with in planta sequences from Juniperus procera Hochst. ex Endlicher, Podocarpus falcatus (Thunb.) R.Br. ex Mirb. and Prunus africana Hook.f. trees of the Afromontane region of Ethiopia (Wubet et al. 2006, 2009). This clade formed a sister group of species with subgroup ‘‘a’’ of Glomus group A sensu Schwarzott et al. (2001) with strong bootstrap (NJ 99%, ML 98%) and posterior probability (PP 100%) support values. The high similarity of G. africanum to AMF sequences obtained from Africa (Wubet et al. 2006, 2009) is especially interesting because the species first was found on that continent (see below), although from a completely different habitat (maritime dune). Other environmental AMF originating from different regions and continents also showed high similarity to G. africanum when the BLAST query was restricted to the AM1-NS31 segment of the SSU sequences (data not shown). BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. 1455 FIG. 13. Neighbor-joining tree showing the phylogenetic positions of Glomus africanum and G. iranicum within Glomus group A inferred from 71 nrDNA SSU sequences with Glomus lamellosum as outgroup. The sequences obtained in this study are shown in boldface. Geographic origin (in parentheses), GenBank accession numbers and hosts of in planta sequences from earlier studies are provided. Values above branches have NJ bootstrap values (1000 replicates; before the slash) and ML bootstrap values (1000 replicates; after slash) as percentages, whereas values below branches are posterior probabilities calculated by Bayesian analysis as percentages. Bootstrap values below 75% and posterior probabilities below 90% are not shown. Bar 5 1 change/100 characters. 1456 MYCOLOGIA Specimens examined. POLAND, Szczecin, under potcultured P. lanceolata, 10 May 2009, Błaszkowski, J., 3167 (HOLOTYPE, DPP); Błaszkowski, J., 3168–3184 (ISTOTYPES, DPP) and two slides at OSC. Etymology. Latin, africanum, referring to the continent from where the fungus was first found. Distribution and habitat. With traditional methods of finding AMF (not molecular) G. africanum has been isolated from six trap cultures containing mixtures of rhizosphere soils/root fragments of four recognized plant species and an unrecognized shrub from two African countries (Egypt, South Africa) and from two trap cultures with soils and roots collected under A. arenaria growing in Bulgaria and Poland (Europe). All the plants colonized maritime sand dunes. No spores of AMF were isolated directly from field-collected samples. The South African plant species sampled were C. geifolia, S. elegans, Tr. divaricata, growing near Strand (34u069S, 18u499E), ca. 50 km southeast of Cape Town and Th. distichum, growing near Strand and in the Reserve Rooiels (34u189S, 18u499E). Strand samples were collected 31 Jul–2 Oct 2005 and those from the Reserve Rooiels 2 Aug 2005. The unrecognized shrub was sampled from Giftung Island (27u109N, 33u569E), Egypt. The rhizosphere soil-root mixture of this plant was sampled 28 Jul 2007. A. arenaria was sampled from dunes of the Black Sea near Varna (43u139N, 27u559E), Bulgaria, on 15 Sep 1998 and from dunes of the Baltic Sea adjacent to Świnoujście (53u559N, 14u149E), northwestern Poland, 10 Jul 2006. Spores of G. africanum were not found in ca. 3000 field-collected soils or in ca. 2500 pot trap cultures representing other regions of Africa and Europe as well as Asia and USA (Błaszkowski pers obs). Notes. The most distinctive structures of G. africanum are its two spore wall layers (FIGS. 3–8), of which the outer layer is hyaline, irregular and much thicker than the structural laminate inner layer, which is exceptionally thin compared with the thickness of the laminate structural spore wall layer of other known Glomus spp. (www.agro.ar.szczecin.pl/,jblaszkowski/). Of the species of Glomeromycota forming glomoidcolored spores with a two-layered spore wall in which the inner layer is laminate, G. africanum spores most resemble in color and size those of G. etunicatum W.N. Becker & Gerd. and G. versiforme (P. Karsten) S.M. Berch. However the darkest spores of G. africanum are markedly darker than the darkest G. versiforme spores (www.agro.ar.szczecin. pl/,jblaszkowski/, Błaszkowski et al. 2003). Moreover G. versiforme spores may be (i) produced singly and in compact epigeous sporocarps (vs. singly and in loose clusters for G. africanum; FIGS. 1, 2) and (ii) slightly larger, (80–)106(–150) mm diam when globose (Daniels and Trappe 1979, Błaszkowski et al. 2003, www.invam.caf.wvu.edu/). Glomus etunicatum produces only single hypogeous spores (Becker and Gerdemann 1977, www.agro.ar.szczecin.pl/,jblaszkowski/, www.invam.caf.wvu.edu/). Both spore wall layers of G. africanum and G. versiforme are of the same type and do not stain in Melzer’s reagent. However spore wall layer 1 of G. africanum is much thicker and layer 2 much thinner than layer 1, (0.7–)1.0(–1.2) mm thick, and layer 2, (2.7–)4.1(–5.4) mm thick, of the spore wall of G. versiforme (www.agro.ar.szczecin.pl/,jblaszkowski/, Błaszkowski et al. 2003). The semipermanent spore wall layer 1 of G. africanum is relatively long-lived and nonreactive in Melzer’s reagent (F IGS . 2–8), while that of G. etunicatum is short-lived, mucilaginous and stains in this reagent (www.agro.ar.szczecin.pl/,jblaszkowski/, Stürmer and Morton 1997). In addition spore wall layer 1 of G. etunicatum is much thinner, 0.5–2.5 mm thick when intact, than spore wall layer 1 of G. africanum, and the upper range of thickness of the laminate spore wall layer 2 of the latter species does not attain even the lower limit of the range of thickness of the laminate spore wall layer of the former fungus, 4.5 mm thick (www.agro.ar.szczecin.pl/,jblaszkowski/). Finally, the subtending hypha of G. africanum spores is less regular (flared to slightly funnel-shaped; FIGS. 1–3, 7, 8) than that of spores of both G. etunicatum and G. versiforme (cylindrical to flared; www.agro.ar.szczecin.pl/,jblaszkowski/, Błaszkowski et al. 2003, www.invam.caf.wvu.edu/). Molecular-phylogenetic analyses results (FIG. 13) indicate that G. africanum has no apparent relatives among described Glomus spp. The phylogenetic position of G. africanum within Glomeromycota is different than that of G. etunicatum and G. versiforme; Glomus africanum has grouped among members of Glomus group A, whereas G. etunicatum represents Glomus group B (Schwarzott et al. 2001) and Glomus versiforme is a close relative of Diversispora spurca (C.M. Pfeiff., C. Walker & Bloss) C. Walker & Schuessler, the type species of family Diversisporaceae C. Walker & Schuessler (Walker and Schübler 2004, Redecker et al. 2007). Of members of Glomus group A, juvenile G. constrictum Trappe spores are similar to mature, small-spored isolates of G. africanum in color and appearance. Moreover spore wall layer 2 of immature G. constrictum spores is of thickness similar to that of layer 2 of mature G. africanum spores (Błaszkowski pers obs). However at maturity G. constrictum spores are much darker, brownish orange (6C8) to dark brown (9F5) to black, than those of G. africanum, and spore wall layer 2 of the former species always is BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. thicker, (7.5–)10.0(–12.0) mm, than layer 1, (0.8–) 2.5(–8.5) mm thick, and much thicker than spore wall layer 2 of G. africanum (Trappe 1977, www.agro.ar. szczecin.pl/,jblaszkowski/). In addition G. constrictum spores generally are much larger, (100–)160 (–220–330) mm diam when globose, than those of G. africanum and the width of the subtending hypha of spores of the former species far exceeds that of the latter fungus, (11.3–)15.0(–17.5) mm wide at the spore base in G. constrictum. Finally, while the subtending hypha of G. constrictum spores typically is markedly constricted at the spore base (Trappe 1977; www.agro. ar.szczecin.pl/,jblaszkowski/), that of G. africanum spores is rarely and only slightly constricted at the base. Glomus iranicum Błaszk., Kovács & Balázs, sp. nov. FIGS. 13–25 MycoBank MB518242 Sporocarpia ignota. Sporae singulatim vel gregatim in solo vel in radice efformatae. Fascicula globosa, oblonga vel irregulares 70–280 3 90–480 mm. Sporae hyalinae vel subluteae; globosae vel subglobosae; (13–)40(–56) mm diam; raro ovoideae, oblongae vel irregulares; 39–54 3 48–65 mm. Tunica sporae stratis tribus (strati 1–3); stratum ‘‘1’’ caducum, glabrum, hyalinum, (0.4–)1.0(–1.5) mm crassum, in solutione Melzeri rufum; stratum ‘‘2’’ semirigidum, glabrum, hyalinum, (0.8–)1.2(–1.5) mm crassum; stratum ‘‘3’’ laminatum, glabrum, hyalinum vel subluteum, (1.2–)2.0(–2.6) mm crassum. Hypha sporifera hyalina; recta vel recurvta; cylindrica vel infundibuliformis; (4.8–)6.9 (–9.8) mm lata ad basim sporae; pariete hyalino vel subluteo, (1.8–)2.6(–3.5) mm crasso, stratis 1–3 in parietem sporae continuantibus. Porus hyphae (1.2–)2.5(–5.0) diam, aperto. Mycorrhizas arbusculares formans. Typus: Polonia: Sedinum (Szczecin), infra P. lanceolata, 10 Mar 2008, J. Błaszkowski, 3185 (Holotypus, DPP). Sporocarps unknown. Spores formed in the soil in loose to compact clusters (FIGS. 14–17 and 22); 70– 280 3 90–480 mm; rarely singly (FIG. 18), occasionally inside roots; develop blastically at the tip of sporogenous hyphae branched from a parent hypha continuous with a mycorrhizal extraradical hypha (FIGS. 14, 15), rarely intercalary. Spores hyaline to pastel yellow (3A4), globose to subglobose, (13–) 40(–56) mm diam, rarely egg-shaped, prolate to irregular; 39–54 3 48–65 mm; with one subtending hypha (FIGS. 14–18 and 20–22). Spore wall composed of three layers (1–3, FIGS. 16–22). Layer 1, forming the spore surface, mucilaginous, roughened, hyaline, (0.4–)1.0(–1.5) mm thick when intact, usually more or less deteriorated in mature spores, almost always sloughed in older specimens (FIGS. 16–22). Layer 2 permanent, semirigid, smooth, hyaline, (0.8–)1.2 (–1.5) mm thick, loosely associated with layer 3 1457 (FIGS. 16–22); in vigorously crushed spores this layer frequently cracks and then separates from layer 3 and usually protrudes because of its rigidity (FIGS. 21, 22). Layer 3 laminate, smooth, hyaline to pastel yellow (3A4), (1.2–)2.0(–2.6) mm thick, sometimes stratifying into groups of or single laminae in crushed spores (FIGS. 16–22). In Melzer’s reagent only layer 1 stains pastel red (7A5) to brownish red (10C6, FIGS. 15, 17–19). Subtending hypha hyaline to pastel yellow (3A4); straight or recurved, cylindrical to slightly funnel-shaped, rarely constricted at spore base; (4.8–)6.9(–9.8) mm wide at the spore base (FIGS. 14, 15, 18, 20–22). Wall of subtending hypha hyaline to pastel yellow (3A4); (1.8–)2.6(–3.5) mm thick at the spore base; composed of three layers continuous with spore wall layers 1–3 (FIGS. 21, 22). Pore (1.2–)2.5(–5.0) mm diam, open (FIGS. 21, 22). Germination unknown. Mycorrhizal associations. In the field G. iranicum was associated with roots of T. aestivum. In one-species pot cultures with P. lanceolata as the host plant G. iranicum formed mycorrhizae with arbuscules and intra- and extraradical hyphae (FIGS. 23–25). No vesicles were found. Arbuscules generally were infrequent and widely dispersed along the root fragments examined. They consisted of a short trunk developed from a parent hypha and numerous branches with fine tips (FIG. 23). Intraradical hyphae grew parallel to the longitudinal root axis, were straight to slightly curved, and (2.3–)4.3(–7.0) mm wide (FIGS. 23–25). They sometimes formed Y-shaped branches and coils (FIGS. 24, 25). Coils were ellipsoidal, 19.0–22.8 3 25.8–56.8 mm, when seen in a plan view, and not numerous and not widely dispersed along roots fragments (FIG. 25). Extraradical hyphae occurred rarely and were (2.4–)4.8(–6.8) mm wide. In 0.1% trypan blue arbuscules stained lilac (16B5) to deep violet (16E8), intraradical hyphae pale violet (16A3) to deep violet (16D8), coils violet (17A6–17B7), and extraradical hyphae pale violet (16A3) to grayish violet (16C6; FIGS. 23–25). Phylogenetic position. Phylogenetic analyses of partial SSU sequences of nrDNA positioned G. iranicum unambiguously in Glomus group A sensu Schwarzott et al. (2001) (FIG. 13). The sequences of the species separated unambiguously from described Glomus species. Although sequences of G. iranicum grouped together with a sequence obtained from roots of P. africana from Africa (Wubet et al. 2009), they are relatively distant. The clade with G. iranicum grouped together without significant support with several lineages of sequences obtained from different in planta diversity studies (Bidartondo et al. 2002; Wubet et al. 2006, 2009; Kovács et al. 2007) but not with any described Glomus species (FIG. 13). 1458 MYCOLOGIA FIGS. 14–21. Glomus iranicum. 14, 15. Spores in loose clusters. 16–19. Spore wall layers (swl) 1–3. 20. Spore wall layers (swl) 2 and 3. Note the thick swl2. Spore wall layer 1 is sloughed in this spore. 21. Spore wall layers (swl) 2 and 3 and subtending hyphal wall layers (shwl) 2 and 3. Spore wall layer 1 and subtending hyphal wall layer 1 are sloughed in this spore. Note the cracked semirigid swl2 and the separated laminae of swl3. 14, 16. Spores in PVLG. 15, 17–21. Spores in PVLG + Melzer’s reagent. 14–21. Differential interference microscopy. Bars: 14, 15 5 20 mm; 16–21 5 10 mm. BŁASZKOWSKI ET AL.: TWO NEW GLOMUS SPP. 1459 FIGS. 22–25. Glomus iranicum. 22. Spore wall layers (swl) 1–3 continuous with subtending hyphal wall layers (shwl) 1–3. Note highly decomposed swl1 and shwl1. 23–25. Mycorrhizae of Glomus iranicum in roots of Plantago lanceolata stained in 0.1% trypan blue. 23. Arbuscule (arb) with trunk (t) developed from parent hypha (ph) and straight intraradical hypha (sih). 24. Y-shaped branch. 25. Coil (c). 22–25. Spores and root fragments in PVLG. 22–25. Differential interference microscopy. Bars: 22–25 5 10 mm. Specimens examined. POLAND, Szczecin, under potcultured P. lanceolata, 2 Jul 2009, Błaszkowski, J., 3185 (HOLOTYPE, DPP); Błaszkowski, J., 3186–3207 (ISTOTYPES, DPP) and two slides at OSC. Etymology. Latin, iranicum, referring to Iran, the only country in which the species has been found. Distribution and habitat. Glomus iranicum was found in only one trap culture with rhizosphere soil and root fragments of T. aestivum collected from a cultivated field in a weakly alkaline clay soil near Khoramshahr, Khuzestan Province (southwestern Iran; 30u309N, 48u099E) 5 Jun 1997. Notes. The distinctive morphological characters of G. iranicum are the hyaline to pale, small spores formed mainly in loose clusters (FIGS. 14–17, 22) and the permanent, semirigid, hyaline and relatively thick middle layer of the three-layered spore wall (FIGS. 16– 22). In crushed spores this layer usually cracks, separates from the laminate layer 3 and frequently protrudes because of its rigidity (FIGS. 19, 21, 22). In addition the fungus has a mucilaginous, short-lived outermost spore wall layer, forming the surface, which stains intensively in Melzer’s reagent (FIGS. 15–19, 22), and the laminate layer 3 frequently stratifies into groups of single laminae in crushed spores (FIG. 21). Of the described Glomus spp. forming clusters of spores having a three-layered spore wall in which the innermost layer is laminate, G. iranicum is most similar to G. xanthium Błaszk., Blanke, Renker & Buscot (Błaszkowski et al. 2004). The darkest G. iranicum spores are pigmented similarly to the lightest G. xanthium spores. Mature spores of both species have a similar size. Spore wall layer 2 in G. iranicum and G. xanthium is permanent, rigid, smooth, hyaline and of similar thickness, and the subtending hypha of spores of both fungi has a similar width. However most G. xanthium spores are markedly darker, to yellow ocher (5C7), and never are hyaline as are most spores of G. iranicum. Most important at maturity spore wall layer 1 of G. xanthium generally is present as a more or less deteriorated, light yellow (4A4) structure (1.2–) 1460 MYCOLOGIA 1.8(–2.7) mm thick, which does not stain in Melzer’s reagent. In contrast spore wall layer 1 of G. iranicum is markedly thinner and stains intensively in Melzer’s reagent (FIGS. 15–19). Moreover this layer is a shortlived component of the spore wall and is rarely present in mature G. iranicum spores (FIGS. 17, 20– 22). Finally, the subtending hypha of G. xanthium spores is more uniform (cylindrical to flared vs. cylindrical to funnel-shaped in G. iranicum), its upper range of width, (0.5–)0.9(–1.5) mm at spore base, does not attain even the lower limit of that of the subtending hypha of G. iranicum spores, and its lumen is occluded by a septum (vs. it is open in G. iranicum; FIGS. 14, 15, 18, 20–22). Another species of the Glomus group that shares some characters with G. iranicum is G. viscosum T.H. Nicolson. Similarly to G. iranicum, many G. viscosum spores arise in loose clusters, the spores are hyaline to pale (pale yellow; www.invam.caf.wvu.edu/, Walker et al. 1995), and their three-layered spore wall comprises two permanent layers (2 and 3) with nearly identical phenotypic and biochemical properties. However G. viscosum spores generally are larger (www.invam.caf. wvu.edu/, Walker et al. 1995), and spore wall layer 2 is much thinner, 0.5 mm thick or less versus (0.8–)1.2 (–1.5) mm thick (FIGS. 16–22) and semiflexible (www. invam.caf.wvu.edu/) versus semirigid, frequently cracking in vigorously crushed spores (FIGS. 20–22) than G. iranicum. In addition, although spore wall layer 1 of G. viscosum is an impermanent structure like that of G. iranicum, it does not stain in Melzer’s reagent (FIGS. 15–19), it generally remains attached to mature spores, and it exudes a mucigel-like substance absorbing soil particles (www.invam.caf. wvu.edu/, Walker et al. 1995) distinguishing it from G. iranicum. Similar to G. africanum, G. iranicum has no apparent molecular relatives among described Glomus spp. (FIG. 13). Although G. iranicum shares some morphological characters with G. viscosum and G. xanthium, the phylogenetic positions of the three species within Glomeromycota are different. Glomus iranicum is a member of Glomus group A, whereas G. viscosum is related to species of Glomus group B (Schwarzott et al. 2001). Although G. xanthium belongs to Glomus group A, its closest molecular relatives are G. caledonium and G. mosseae (Błaszkowski et al. 2004), which are clearly separated from G. iranicum within the phylogenetic analyses presented here (FIG. 13). ACKNOWLEDGMENTS We thank Prof Zdzisław Koszański, Department of Water Management, West Pomeranian University of Technology, Szczecin, for collecting the A. arenaria soil-root sample from Varna (Bulgaria). We also thank Dr P. Schreiner, Mycologia associate editor, and two anonymous reviewers for valuable comments. This study was supported in part by the Polish Committee of Scientific Researches, grants 2 PO4C 041 28 and 164/N-COST/2008/0, and the Hungarian Research Fund, OTKA K72776. A part of the study was conducted during the Experienced Researcher Fellowship of the Alexander von Humboldt Foundation awarded to G. 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