Mycologia, 101(2), 2009, pp. 247–255. DOI: 10.3852/08-087 # 2009 by The Mycological Society of America, Lawrence, KS 66044-8897
Glomus perpusillum, a new arbuscular mycorrhizal fungus Janusz Błaszkowski1
Maritime sand dunes harbor exceptionally abundant and diverse populations of arbuscular fungi, mainly because of their low nutrient and organic matter content (Dalpe´ 1989; Koske 1987, 1988; Nicolson and Johnston 1979; Tadych and Błaszkowski 2000). Of many dune plants, Ammophila arenaria (L.) Link is the most important sandstabilizing species in maritime dunes of Europe (Rodrı´guez-Echeverrı´a and Freitas 2006), and its roots commonly host a diverse population of arbuscular mycorrhizal fungi (Kowalchuk et al 2002, Nicolson and Johnston 1979, Tadych and Błaszkowski 2000). At present phylum Glomeromycota comprises 14 genera of arbuscular mycorrhizal fungi, the most numerous is genus Glomus Tul. & C. Tul. It includes ca. 53% of the ca. 200 species of arbuscular mycorrhizal fungi described to date (www.agro.ar. szczecin.pl/,jblaszkowski). Nineteen Glomus spp. have been described from spores isolated from maritime dunes, and many others have associated with roots of dune plants (www.agro.ar.szczecin.pl/ ,jblaszkowski, Sridhar and Beena 2001). However results of molecular diversity studies of arbuscular fungi associated with plants of both cultivated and natural sites, including A. arenaria, indicate that many in planta sequence types could not be assigned to described glomeromycotan species (Hijri et al 2006, Kowalchuk et al 2002, Rodrı´guez-Echeverrı´a and Freitas 2006). Its spores are needed to name an undescribed arbuscular fungus. Stutz and Morton (1996) showed that lack of sporulation is a common phenomenon in some habitats. An effective method to induce sporulation of fungi hidden in roots is cultivation of field-collected rhizosphere soils and roots in successive trap cultures (Stutz and Morton 1996). Examination of a single pot trap culture using a mixture of a rhizosphere soil and root fragments of A. arenaria collected from dunes of the Mediterranean Sea adjacent to Calambrone, Italy, revealed many aggregates of spores of a new Glomus sp. that were not present in the soil when the culture was established. This fungus subsequently was propagated in single-species cultures and characterized by examining spores, mycorrhizal structures in roots and the partial nucleotide sequence of the small ribosomal subunit (18S) gene. This fungus is described and illustrated below as G. perpusillum sp. nov.
Department of Plant Protection, University of Agriculture, Słowackiego 17, PL-71434 Szczecin, Poland
Ga´bor M. Kova´cs Tı´mea Bala´zs Department of Plant Anatomy, Institute of Biology, Eo¨tvo¨s Lora´nd University, Pa´zma´ny Pe´ter se´ta´ny 1/C, 1117 Budapest, Hungary
Abstract: A new arbuscular mycorrhizal fungal species of genus Glomus, G. perpusillum (Glomeromycota), forming small, hyaline spores is described and illustrated. Spores of G. perpusillum were formed in hypogeous aggregates and occasionally inside roots. They are globose to subglobose, (10–)24(–30) mm diam, rarely egg-shaped, oblong to irregular, 18–25 3 25–63 mm. The single spore wall of G. perpusillum consists of two permanent layers: a finely laminate, semiflexible to rigid outer layer and a flexible to semiflexible inner layer. The inner layer becomes plastic and frequently contracts in spores crushed in PVLG-based mountants and stains reddish white to grayish red in Melzer’s reagent. Glomus perpusillum was associated with roots of Ammophila arenaria colonizing sand dunes of the Mediterranean Sea adjacent to Calambrone, Italy, and this is the only site of its occurrence known to date. In single-species cultures with Plantago lanceolata as host plant, G. perpusillum formed vesicular-arbuscular mycorrhiza. Phylogenetic analyses of partial SSU sequences of nrDNA placed the species in Glomus group A with no affinity to its subgroups. The sequences of G. perpusillum unambiguously separated from the sequences of described Glomus species and formed a distinct clade together with in planta arbuscular mycorrhizal fungal sequences found in alpine plants. Key words: arbuscular fungi, Glomeromycota, molecular phylogeny, mycorrhizae, new species INTRODUCTION
Arbuscular mycorrhizal fungi of phylum Glomeromycota are considered to belong to the most common soil fungi in the world and associate with at least 80% of vascular land plants (Smith and Read 1997).
Accepted for publication 27 October 2008. 1 Corresponding author. E-mail:
[email protected]. szczecin.pl
247
248
MYCOLOGIA MATERIALS AND METHODS
Establishment and growth of trap and single-species cultures, extraction of spores, and staining of mycorrhizae.—Spores examined in this study came from both pot trap and singlespecies cultures. Trap cultures were established to obtain a great number of living spores and to initiate sporulation of species that were present but not sporulating in the field collections (Stutz and Morton 1996). The method used to establish trap cultures, the growth conditions and the methods of spore extraction and staining of mycorrhizal structures were described by Błaszkowski et al (2006). Single-species cultures were established and grown as described by Błaszkowski et al (2006), with two exceptions. First, instead of marine sand, the growing medium used was 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, Bystre´, Slovakia) of grains 2.5–5 mm diam. Clinopthilolite is a crystaline hydrated alumosilicate of alkali metals and alkaline earth metals having a high ion exchange capacity and reversible hydration and dehydration properties. The sand-clinopthilolite mixture was pH 7.3. Second, 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 the cultures with other arbuscular fungi but still allow exchange of gases we left a 1 cm wide opening in the center of the upper part of each bag, while the edges on both sides were closed with small plastic clips. The cultures were watered with tap water once a week and harvested after 5 mo when spores were extracted for study. To reveal mycorrhizal structures root fragments located ca. 1–5 cm below the upper level of the growing medium were cut off with a scalpel. The host plant used in both trap and singlespecies cultures was Plantago lanceolata L. Microscopy survey.—Morphological properties of spores and their wall structure were determined based on examination of at least 100 spores mounted in 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 their contents from oil droplets. They were examined under an Olympus BX 50 compound microscope equipped with Nomarski differential interference contrast optics. Microphotographs were recorded with a Sony 3CDD color video camera. Terminology of spore structure is that suggested by Stu¨rmer 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). 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), University of Agriculture, Szczecin, Poland, and in the herbarium at Oregon State University (OSU) in Corvallis, Oregon. Color microphotographs of spores of G. perpusillum can be viewed at the URL http://www.agro.ar.szczecin.pl/ ,jblaszkowski/. Molecular analyses.—Two to four spores of the fungus were transferred into 40 mL sterile ultra clean water in 0.2 mL PCR tubes and crushed with a pipette tip under dissecting microscope. They were vortexed and incubated at 96 C for 7 min and vortexed again. A total of 2–4 mL of these solutions were used as target DNA in the PCR with the PuReTaqTM Ready-To-GoTM PCR beads (GE Healthcare, Buckinghamshire, UK). PCR were carried out with primers NS31 (Simon et al 1992) and AM1 (Helgason et al 1998). The appropriate size amplicons were isolated from agarose gel with a Gel-M Extraction System (Viogene, Hong-Kong, China). PCR products were cloned into pGEM-T Easy Vector (Promega, Madison Wisconsin) and the clones were transformed into competent JM109 Escherichia coli cells (Promega, Madison Wisconsin). The preparation of the plasmid DNA was carried out with a MiniM Plasmid DNA Extraction kit (Viogene, Hong-Kong, China). Positive clones were sequenced on both strands with universal forward and reverse primers with an ABI PRISM 3.1 BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystem, Foster City, California). The electrophoresis was carried out in an ABI PRISM 3100 Genetic Analyzer at the service laboratory of Biology Research Center (Szeged, Hungary). Electrophoregrams were processed and analyzed with the Staden Program Package (Staden et al 2000). The sequences have been deposited in GenBank (FJ164234–FJ164242). The sequences were analyzed together with identified taxa of phylum Glomeromycota and subsequently with a dataset of Glomus Group A sequences with unidentified arbuscular fungi sequences from in planta diversity studies (Kova´cs et al 2007). Here we present in detail only results of the analyses of the Glomus Group A dataset. Sequences were aligned with Multalin (Corpet 1988) running on the INRA server (http://prodes.toulouse.inra. fr/multalin/multalin.html). The alignments were checked and edited manually with ProSeq 2.9 (Filatov 2002). In the final dataset three sequences from the newly described fungus were analyzed together with 47 glomeromycotan sequences with G. claroideum as outgroup (FIG. 17). The alignment was 513 characters long. It has been deposited in the TreeBase (S2170, M4117). Maximum likelihood (ML) phylogenetic analyses were carried out with the program PHYML (Guindon and Gascuel 2003). The GTR nucleotide substitution model was used with ML estimation of base frequencies. The proportion of the invariable sites was estimated and optimized. Four substitution rate categories were set, and the gamma distribution parameter was estimated and optimized. Bootstrap analysis with 1000 replicates was used to test the statistical support of the branches. The same substitution model was used in Bayesian analyses performed with MrBayes 3.1 (Huelsen-
BŁASZKOWSKI ET AL: A beck and Ronquist 2001, Ronquist and Huelsenbeck 2003). The Markov chain was run for 2 000 000 generations, sampling in every 100 steps, and with a burn-in at 7500 trees. The details of the analyses and the alignments are available on request. Phylogenetic trees were viewed and edited by the Tree Explorer of the MEGA 3.1 program (Kumar et al 2004). TAXONOMY
Glomus perpusillum Błaszk. & Kova´cs sp. nov. FIGS. 1–17 MycoBank No. MB 512346 Sporocarpia ignota. Sporae gregarius in solo vel in radici efformatae. Gregariae globosae, oblongae vel irregulares, 80–280 3 90–320 mm. Sporae hyalinae; globosae vel subglobosae; (10–)24(–30) mm diam; raro ovoidae, oblongae vel irregulares; 18–25 3 25–63 mm. Tunica sporae e stratis duobus (strati 1 at 2); stratum ‘‘1’’ laminatum, semielasticum vel rigidum, glabrum, hyalinum, (1.0–)1.4 (–1.7) mm crassum; stratum ‘‘2’’ elasticum vel semielasticum, glabrum, hyalinum, (0.5–)0.8(–1.2) mm crassum; in solutione Melzeri rubicoso vel rufo. Hypha subtenda hyalina; recta vel recurva; cylindrica vel infundibuliforma; (2.0–)3.2(–4.7) mm lata at basim sporae; pariete hyalino, (0.7–)1.3(–2.0) mm crasso, stratis 1 at 2 sporae continuans. Porus (0.7–)1.3(–2.0) diam, aperto. Mycorrhizas vesiculararbusculares formans. Typus: Poland: Szczecin, infra P. lanceolata, 15 Mar 2008, J. Błaszkowski, 2770 (Holotypus, DPP).
Spores formed in hypogeous, loose to compact aggregates; 80–280 3 90–320 mm; occasionally inside roots (FIGS. 1–3); develop blastically at the tip of hyphae branched from a parent hypha (FIGS. 1, 2, 4 and 6–11) continuous with a mycorrhizal extraradical hypha. Spores hyaline; globose to subglobose; (10–) 24(–30) mm diam; rarely egg-shaped, prolate to irregular; 18–25 3 25–63 mm; with one subtending hypha (FIGS. 1–11). Subcellular structure of spores consists of a spore wall composed of two hyaline, permanent, smooth layers (layers 1 and 2, FIGS. 4–9 and 11). Layer 1, forming the spore surface is finely laminate, semiflexible to rigid, (1.0–)1.4(–1.7) mm thick. Layer 2 is flexible to semiflexible, (0.5–)0.8 (–1.2) mm thick, always tightly adherent to layer 1 in intact or slightly crushed spores; in spores vigorously crushed in PVLG-based mountants, it becomes plastic and frequently contracts, separating from layer 1 (FIGS. 4–9 and 11). In Melzer’s reagent only layer 2 stains reddish white (9A2) to grayish red (9B6, FIGS. 2 and 5–11). Subtending hypha is hyaline; straight or recurved; cylindrical to slightly funnel-shaped; (2.0–) 3.2(–4.7) mm wide at the spore base (FIGS. 2 and 6– 11). Wall of subtending hypha is hyaline; (0.7–)1.3 (–2.0) mm thick at the spore base; composed of two layers continuous with spore wall layers 1 and 2 (FIG. 11). Pore is (0.7–)1.3(–2.0) mm diam, open. Germination unknown.
NEW
GLOMUS
SP.
249
Mycorrhizal associations. In the field G. perpusillum was associated with roots of A. arenaria colonizing sand dunes of the Mediterranean Sea adjacent to Calambrone, Italy (43u359N, 10u189E.) In single-species cultures with P. lanceolata as the host plant, G. perpusillum formed mycorrhizae consisting of arbuscules, vesicles, as well as intraand extraradical hyphae (FIGS. 12–16). In most root fragments arbuscules were numerous and evenly distributed along the root axis. Arbuscules consisted of a short trunk grown from a parent hypha and numerous branches with fine tips. Vesicles occurred rarely and usually in groups and were egg-shaped to prolate, 10.0–20.0 3 17.5–52.5 mm. Intraradical hyphae grew along the root axis, were numerous, (2.7–) 3.7(–6.6) mm wide, straight or slightly curved, and sometimes formed H- or Y-shaped branches and coils. The coils were ellipsoid; 12.5–31.6 3 30.0–92.5 mm, when seen in plan view; and sparse to abundant, depending on the root fragment examined. Extraradical hyphae were (2.0–)3.8(–5.9) mm wide and were scarce to abundant. Arbuscules in 0.1% trypan blue stained violet white (16A2) to grayish violet (16C5), vesicles stained lilac (16B5) to reddish violet (16C6), intraradical hyphae stained pale violet (16A3) to deep violet (16D8), coils stained violet white (16A2) to deep violet (16E8), and extraradical hyphae stained pale violet (16A3) to deep violet (16D8). Phylogenetic position. Glomus perpusillum unambiguously grouped within Glomus group A clade sensu Schwarzott et al (2001) (FIG. 17). The sequences of G. perpusillum grouped together with unidentified Glomus sequences obtained from the roots of alpine plants in Austria. This well supported distinct group of sequences had no affinity to any described Glomus species and was separate from the three known subgroups of the Glomus group A clade (FIG. 17). Specimens examined. POLAND. Szczecin, under potcultured P. lanceolata, 15 Mar 2008, Błaszkowski, J., 2770 (HOLOTYPE, DPP); Błaszkowski, J. 2771–2813 (ISOTYPES, DPP) and two slides at OSU.
Etymology. Latin, perpusillum, referring to the small spores formed by the fungus. Distribution and habitat. Glomus perpusillum was found only in one pot trap culture consisting of rhizosphere soil and root fragments from A. arenaria growing in sand dunes of the Mediterranean Sea adjacent to Calambrone, Italy. Spores of this fungus were not found in ca. 3000 field-collected soils or in ca. 2000 pot trap cultures representing different regions of Europe, as well as Africa, Asia and USA. (Błaszkowski pers obs). Thus the lack of G. perpusillum spores in ca. 2000 trap cultures that usually reveal numerous arbuscular mycorrhizal fungi not present as
250
MYCOLOGIA
FIGS. 1–8. Glomus perpusillum. 1 and 2. Spores in loose aggregates. 3. Intraradical spores stained in 0.1% trypan blue (note the contracted spore wall layer 2). 4–8. Spore wall layers (swl) 1 and 2. Note wrinkled spore wall layer 1 (FIGS. 4, 7 and 8) and spore wall layer 2 highly contracted and stained in PVLG + Melzer’s reagent; sg 5 sand grain. 1. Spores in lactic acid. 2, 5–8. Spores in PVLG + Melzer’s reagent. 3, 4. Spores mounted in PVLG. 1–8. Differential interference microscopy. Bars: 1 5 50 mm, 2 and 3 5 20 mm, 4–8 5 10 mm.
BŁASZKOWSKI ET AL: A spores before culture (Stutz and Morton 1996) indicates this species is extremely rare in the regions sampled by the first author of this paper. Other arbuscular fungi sporulating in the trap culture along with G. perpusillum were Intraspora schenckii (Sieverd. & S. Toro) Oehl & Sieverd., G. aurantium Błaszk et al, G. constrictum Trappe, Scutellospora fulgida Koske & C. Walker, and S. persica (Koske & C. Walker) C. Walker & F.E. Sanders. The occurrence of sporulating fungi of the Glomeromycota in the field sample was not determined.
DISCUSSION
The distinctive properties of G. perpusillum are its exceptionally small, hyaline spores formed only in aggregates and remaining hyaline throughout their life cycle, as well as the simple spore wall structure consisting of two permanent layers. The outer layer usually wrinkles in crushed spores, and the inner layer frequently contracts in spores crushed in PVLG-based mountants and stains red in Melzer’s reagent (FIGS. 1, 2 and 4–11). In most described Glomus spp. the outermost spore wall layer, forming the spore surface, deteriorates with age in both the field and pot culture conditions (Bentivenga and Morton 1995, www.agro. ar.szczecin.pl/,jblaszkowski). In contrast layer 1 of the spore wall of G. perpusillum always was retained intact in spores isolated from long-term cultures. Thus it is of the type of permanent wall layers sensu Morton (www.invam.caf.wvu.edu) and should be present in specimens isolated from the field. After storage of G. perpusillum spores in lactic acid-based mountants, the staining intensity of their inner spore wall layer in Melzer’s reagent was slightly reduced but never disappeared, as in some other species of the Glomeromycota (Morton 1986). Other described species of genus Glomus forming only hyaline spores with a 2-layered spore wall are G. cerebriforme McGee and G. minutum Błaszk., Tadych & Madej (www.agro.ar.szczecin.pl/,jblaszkowski, McGee 1986). Living spores or sequences of G. cerebriforme unfortunately were not available to us. Dr P. McGee no longer has the fungus in culture (Błaszkowski pers comm), and numerous requests sent to the curator of the Herbarium of the Botanic Gardens in Adelaide, Australia, were unanswered. In addition DNA extracts from spores of G. minutum collected from an old single-species culture failed to produce PCR products (D. Redecker pers comm). Thus we have to rely on spore morphology solely to explain the differences among G. perpusillum and G. cerebriforme or G. minutum. Five properties separate G. cerebriforme from G.
NEW
GLOMUS
SP.
251
perpusillum. First, G. cerebriforme produces spores in epigeous sporocarps in the field or in loose hypogeous hyphal masses in pot cultures (McGee 1986) whereas those of G. perpusillum occur mainly in hypogeous aggregates (FIGS. 1, 2 and 4–9), rarely inside roots (FIG. 3). Second, spores of the former species form on racemose hyphae, and those of G. perpusillum develop at the tip of straight or slightly curved hyphal branches of a parent hypha (FIGS. 2, 4 and 6–10). Third, only the smallest spores of G. cerebriforme (25 mm diam when globose) overlap with the mean spore size (24 mm diam) of G. perpusillum. The largest spores of the former species are ca. two- to almost threefold larger than the largest spores of the latter fungus. Fourth, the outer spore wall layer of G. cerebriforme is much thicker (2–4 mm) than that of G. perpusillum (1.0–1.7 mm) (McGee 1986). Although the inner wall layers of spores of both species are flexible, the unique properties of the spore wall layer 2 of G. perpusillum are its plasticity and contractibility (FIGS. 4–9 and 11). This layer also stains intensively in Melzer’s reagent (FIGS. 2 and 5–11). The reactivity of the spore wall components of G. cerebriforme in this reagent unfortunately is unknown. Fifth, the lumen of the subtending hypha of G. cerebriforme spores is closed by a membranous septum continuous with layer 2 of the spore wall while that of spores of G. perpusillum is open (FIGS. 6–11). Glomus minutum and G. perpusillum differ in the range of spore size, the phenotypic and biochemical properties of the spore wall components, as well as the morphology of the subtending hyphae. The mean spore size of the former species (39 mm diam when globose; www.agro.ar.szczecin.pl/,jblaszkowski, Błaszkowski et al 2000) is markedly larger than the largest globose spores (30 mm diam) of the new fungus. Although the outer spore wall layers of both species are semiflexible, spore wall layer 1 of G. minutum is uniform (vs. finely laminate in G. perpusillum) and much thinner ([0.2–]0.6[–0.7]) mm vs. [1.0–]1.4 [–1.7] mm) than G. perpusillum. Spore wall layer 2 of G. minutum is also laminate, nonflexible to semiflexible, and nonreactive in Melzer’s reagent. While the subtending hypha of G. perpusillum is cylindrical to slightly funnel-shaped and (2.0–)3.2(–4.7) mm wide at the spore base (FIGS. 2 and 6–11), that of G. minutum is more regular in shape (cylindrical to flared) and much wider ([4.2–]5.7[–8.1] mm) at the spore base. Finally, the subtending hypha of G. minutum is closed by a septum continuous with the innermost lamina of spore wall layer 2, and that of G. perpusillum is open (FIGS. 6–11). Spores of G. microaggregatum also are hyaline in youth and their spore wall consists of two layers, frequently wrinkling in crushed spores (Koske et al
252
MYCOLOGIA
FIGS. 9–16. Glomus perpusillum. 9. Semiflexible spore wall layer 1 (swl1) with small wrinkles and slightly plastic spore wall layer 2 (swl2). 10. Spore(s) and slightly funnel-shaped subtending hypha (sh). 11. Wrinkled spore wall layer 1 (swl1), highly contracted and stained spore wall layer 2 (swl2) and subtending hyphal wall layers (shwl) 1 and 2. 12. Arbuscules. 13. Vesicles. 14. H-shaped branch. 15. Y-shaped branch. 16. Coil. 9–11. Spores in PVLG + Melzer’s reagent. 12–16. Mycorrhiza of G. perpusillum in roots of Plantago lanceolata stained in 0.1% trypan blue. 9–16. Differential interference microscopy. Bars: 9–16 5 10 mm.
BŁASZKOWSKI ET AL: A
NEW
GLOMUS
SP.
253
FIG. 17. Maximum likelihood tree inferred from partial SSU sequences showing the phylogenetic position of G. perpusillum in the Glomus Group A clade. G. perpusillum sequences are shown in boldface. The host plant and country are given for the in planta sequences grouping with G. perpusillum. Values above branches are the bootstrap values (1000 replicates) and values below branches are posterior probabilities shown as a percentage (values not shown if below 70% and 90% respectively).
254
MYCOLOGIA
1986). However the inner spore wall layer of this fungus neither becomes plastic nor contracts and stains in spores crushed in a mixture of PVLG and Melzer’s reagent, as the spore wall layer 2 of G. perpusillum does (FIGS. 5–9 and 11; Błaszkowski pers obs). In addition spores of G. microaggregatum darken to brownish yellow with age (Koske et al 1986) whereas those of G. perpusillum remain hyaline thorough their life cycle. As mentioned previously the sequences of G. perpusillum formed a distinct clade within Glomus group A fungi (sensu Schwarzott et al 2001) together with unidentified in planta sequences of arbuscular mycorrhizal fungi from alpine plants sampled in Austria (FIG. 17). However this clade had no affinity to any described Glomus species or to the three subgroups of Glomus group A fungi. Thus phylogenetic analyses confirm the uniqueness of G. perpusillum. ACKNOWLEDGMENTS
This study was supported in part by the Polish Committee of Scientific Researches, grants 2 PO4C 041 28 and 164/NCOST/2008/0, and the Hungarian Research Fund, OTKA K72776. LITERATURE CITED
Bentivenga SP, Morton JB. 1995. A monograph of the genus Gigaspora, incorporating developmental patterns of morphological characters. Mycologia 87:719–731. Błaszkowski J, Renker C, Buscot F. 2006. Glomus drummondii and G. walkeri, two new species of arbuscular mycorrhizal fungi (Glomeromycota). Mycol Res 110:555–566. ———, Tadych M, Madej M. 2000. Glomus minutum, a new species in Glomales (Zygomycetes) from Poland. Mycotaxon 76:187–195. Corpet F. 1988. Multiple sequence alignment with hierarchical clustering. Nuc Acids Res 16:10881–10890. Dalpe´ Y. 1989. Inventaire et repartition de la flore endomycorhizienne de dunes et de rivages maritimes du Quebec, du Nouveau-Brunswick et de la NouvelleEcosse. Naturaliste can (Rev Ecol Syst) 116:219–236. Filatov DA. 2002. ProSeq: a software for preparation and evolutionary analysis of DNA sequence datasets. Mol Ecol Notes 2:621–624. Gilmore AE. 1968. Phycomycetous mycorrhizal organisms collected by open-pot culture methods. Hilgardia 39: 87–105. Guindon S, Gascuel O. 2003. A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst Biol 52:696–704. Helgason T, Daniell TJ, Husband R, Fitter AH, Young JPW. 1998. Ploughing up the wood-wide Web? Nature 394: 431. Hijri I, Sy´korova Z, Oehl F, Ineichen K, Ma¨der P, Wiemken A, Redecker D. 2006. Communities of arbuscular
mycorrhizal fungi in arable soils are not necessarily low in diversity. Mol Ecol 15:2277–2289. Huelsenbeck JP, Ronquist F. 2001. MrBayes: Bayesian inference of phylogeny. Bioinformatics 17:754–755. Kornerup A, Wanscher JH. 1983. Methuen handbook of colour. 3rd ed. London: Eyre Methuen. 252 p. Koske RE. 1987. Distribution of VA mycorrhizal fungi along a latitudinal temperature gradient. Mycologia 79:55– 68. ———. 1988. Vesicular-arbuscular mycorrhizae of some Hawaiian dune plants. Pacific Sci 42:217–229. ———, Gemma JN, Olexia PD. 1986. Glomus microaggregatum, a new species in the Endogonaceae. Mycotaxon 26:125–132. Kova´cs GM, Bala´zs T, Pe´nzes Z. 2007. Molecular study of the arbuscular mycorrhizal fungi colonizing the sporophyte of the eusporangiate rattlesnake fern (Botrychium virginianum, Ophioglossaceae). Mycorrhiza 17:597– 605. Kowalchuk GA, de Souza FA, van Veen JA. 2002. Community analysis of arbuscular mycorrhizal fungi associated with Ammophila arenaria in Dutch coastal sand dunes. Mol Ecol 11:571–581. Kumar S, Tamura K, Nei M. 2004. MEGA3: integrated software for molecular evolutionary genetics Analysis and sequence alignment. Brief Bioinform 5:150–163. McGee PA. 1986. Further sporocarpic species of Glomus (Endogonaceae) from South Australia. Trans Brit Mycol Soc 87:123–129. Mirek Z, Pie˛kos´-Mirkowa H, Zaja˛c A, Zaja˛c M. 1995. Vascular plants of Poland: a checklist. Polish Botanical Studies. Krako´w: Guidebook 15. 303 p. Morton JB. 1986. Effects of mountants and fixatives on wall structure and Melzer’s reaction in spores of two Acaulospora species (Endogonaceae). Mycologia 78: 787–794. Nicolson TH, Johnston C. 1979. Mycorrhiza in Gramineae III. Glomus fasciculatum as the endophyte of pioneer grasses in maritime sand dunes. Trans Br Mycol Soc 72: 261–268. Omar MB, Bollan L, Heather WA. 1979. A permanent mounting medium for fungi. Bull Brit Mycol Soc 13: 31–32. Rodrı´guez-Echeverrı´a S, Freitas H. 2006. Diversity of AMF associated with Ammophila arenaria spp. arundinacea in Portuguese sand dunes. Mycorrhiza 16:543–552. Ronquist F, Huelsenbeck JP. 2003. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19:1572–1574. Schwarzott D, Walker C, Schu¨ßler A. 2001. Glomus, the largest genus of the arbuscular mycorrhizal fungi (Glomales) is nonmonophyletic. Mol Phyl Evol 21: 190–197. Simon L, Lalonde M, Bruns TD. 1992. Specific amplification of 18S fungal ribosomal genes from vesiculararbuscular endomycorrhizal fungi colonizing roots. Appl Environ Microbiol 58:291–293. Smith SE, Read DJ. 1997. Mycorhizal symbiosis. San Diego: Academic Press. 605 p. Sridhar KR, Beena KR. 2001. Arbuscular mycorrhizal
BŁASZKOWSKI ET AL: A research in coastal sand dunes: a review. Proc Nat Acad Sci India 71:179–205. Staden R, Beal KF, Bonfield JK. 2000. The Staden package, 1998. Method Mol Biol 132:115–130. Stu¨rmer SL, Morton JB. 1997. Developmental patterns defining morphological characters in spores of four species in Glomus. Mycologia 89:72–81. Stutz JC, Morton JB. 1996. Successive pot cultures reveal high species richness of arbuscular mycorrhizal fungi in arid ecosystems. Can J Bot 74:1883–1889. Tadych M, Błaszkowski J. 2000. Arbuscular fungi and
NEW
GLOMUS
SP.
255
mycorrhizae (Glomales) of the Słowin´ski National Park, Poland. Mycotaxon 74:463–483. Walker C. 1983. Taxonomic concepts in the Endogonaceae: spore wall characteristics in species descriptions. Mycotaxon 18:443–455. ———, Trappe JM. 1993. Names and epithets in the Glomales and Endogonales. Mycol Res 97:339–344. ———, Vestberg M. 1994. A simple and inexpensive method for producing and maintaining closed pot cultures of arbuscular mycorrhizal fungi. Ag Sci Finland 3:233–240.
