Journal of Heredity Advance Access originally published online on April 13, 2006
Journal of Heredity 2006 97(3):270-278; doi:10.1093/jhered/esj031
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High Levels of Genetic Variation Exist in Aspergillus niger Populations Infecting Welwitschia mirabilis Hook
From the Department of Biology, 1205 Noyce Science Center, Grinnell College, 1116 8th Avenue, Grinnell, IA 50112
Address correspondence to K. Jacobson at the address above, or e-mail: jacobsok{at}grinnell.edu.
Aspergillus niger is an asexual, haploid fungus which infects the seeds of Namibia's national plant, Welwitschia mirabilis, severely affecting plant viability. We used 31 randomly amplified polymorphic DNA markers to assess genetic variation among 89 A. niger isolates collected from three W. mirabilis populations in the Namib Desert. While all isolates belonged to the same vegetative compatibility group, 84% were unique genotypes, and estimates of genotypic evenness and Simpson's index of diversity approached 1.0 in the three populations. Analysis of molecular variance revealed that 78% of the total variation sampled was among isolates from individual W. mirabilis plants. Lower, but significant, amounts of variation detected among isolates from different plants (12%) and different sites (10%) also indicated some site- and plant-level genetic differentiation. Total gene diversity (HT = 0.264) was mostly attributable to diversity within populations (HS = 0.217); the relatively low level of genetic differentiation among the sites (GST = 0.141) suggests that gene flow is occurring among the three distant sites. Although sexual reproduction has never been observed in this fungus, parasexuality is a well-known phenomenon in laboratory strains. We thus attribute the high levels of genetic variation to parasexuality and/or wind-facilitated gene flow from an as of yet undocumented broader host range of the fungus on other desert vegetation. Given the apparent ease of transmission, high levels of genetic diversity, and potentially broad host range, A. niger infections of W. mirabilis may be extremely difficult to control or prevent.
Welwitschia mirabilis Hook. is a large, leafy gymnosperm endemic to the arid and semiarid environments of Namibia and Angola. As the protected national plant of Namibia, W. mirabilis is of great conservation value, both as an important tourist attraction and as a biological curiosity (Barnard 1998). The range of W. mirabilis extends over a 150,000-km2 area along the southwestern Namib coast from the Nicolau River in southern Angola to the Kuiseb River and the central Namib Sand Sea in central Namibia (Kers 1967) (Figure 1). Geographically discrete populations of 2 to over 2,000 individuals (Henschel and Seely 2000) are typically found in and along ephemeral watercourses and shallow plains (Jacobson et al. 1993a). A recent study by Wetschnig and Depish (1999) eliminated wind as a possible pollination vector for W. mirabilis and established flies as the primary pollination agent. Reflecting likely dispersal patterns by these pollinators, as well as the large endosperm-rich seeds, Jacobson and Lester (2003) found that genetic differences between populations of W. mirabilis correlate with geographic distances between them, with gene flow occurring between populations 6 km apart but not between populations located 18 or more kilometers apart.
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Individuals of W. mirabilis are estimated to be up to 3000 years old (Bornman et al. 1972; von Willert and Wagner-Douglass 1994), and recent fossil evidence suggests that Welwitschia-related plants were present during the lower Cretaceous, approximately 112 million years ago (Dilcher 2005). There is concern, however, about the long-term viability of the species (Barnard 1998) because the seeds of many W. mirabilis populations are infected by a fungal seed pathogen in the Aspergillus niger van Tiegh aggregate. Initially noted by Hooker (1863) as a dark fungal infection of female cones in his original description of W. mirabilis, this pathogen severely reduces the reproductive viability of infected plants, with average infection rates ranging from 64% to 100% at 13 sites (1,013 plants sampled) (Cooper-Driver et al. 2000). While uninfected plants are highly fertile, juveniles are rare (Bornman 1978; Bustard 1990), and the fungal pathogen has recently been implicated as a primary cause of low recruitment.
A. niger conidia are small with thick, darkly pigmented walls that prevent desiccation and provide protection from UV radiation (Al-Musallam 1980). While it is likely that the conidia are wind dispersed over long distances, Bornman (1978) suggested that they are dispersed locally by a flightless, herbivorous insect, Probergrothius sexpunctatis (Pyrrhocoridae, Heteroptera). The ability of this insect to carry and introduce fungal conidia to female cones with its long proboscis has been demonstrated in the laboratory (Cooper-Driver et al. 2000). In addition, a recent spatial autocorrelation study of A. niger infection levels in W. mirabilis populations (Cooper-Driver et al. 2000) found spatial clustering of the pathogen in patterns that support this hypothesis of transmission by flightless insects with limited individual ranges.
While A. niger is classified as an asexual deuteromycete (Fennell 1977; Raper and Fennell 1965), its parasexual behavior has been extensively studied in the laboratory (Debets 1998; Lhoas 1967; Pontecorvo 1956). The relevance of parasexuality in maintaining genetic variation in natural populations of A. niger is not known, but in laboratory studies the rates of mitotic recombination (crossing over and haploidization) in A. niger were calculated to be 100x higher than meiotic crossing over in the sexual species Aspergillus nidulans (Lhoas 1967). While the exact significance of this calculation is unclear because no one has observed the frequency of heterokaryon formation (the initial step in the parasexual process) in wild populations (Debets 1998), recent studies of populations of a variety of fungal pathogens have shown that even strictly asexual species often have recombinant population structures. Examples of putatively asexual fungi with genetic evidence for recombination include Alternaria brassicae, Alternaria brassicicola, and Alternaria tenuissima (Berbee et al. 2003); Aspergillus flavus (Geiser et al. 1998); Beauveria bassiana (Paccola-Meirelles et al. 1991); Candida albicans (Gräser et al. 1996); Coccidioides immitis (Burt et al. 1996); Cochliobolus carbonum (Lodge and Leonard 1984); Cochliobolus sativus (Zhong and Steffenson 2001); Colletotrichum sublineolum (Souza-Paccola et al. 2003); Fusarium culmorum (Miedaner et al. 2001); Magnaporthe grisea (Zeigler et al. 1997); and Ustilago scabiosae (Garber and Ruddat 1992).
In order for parasexual recombination to occur between isolates, they must belong to the same vegetative compatibility group (VCG). Isolates that are vegetatively compatible have identical alleles at loci that facilitate hyphal fusion (plasmogamy), heterokaryon formation, and nuclear exchange analogous to histocompatibility loci in vertebrates (Brasier 1992). Fungal species and populations vary tremendously in the number of VCGs present. For example, only one to two VCGs were found in Cryphonectria parasitica populations in Europe, whereas a single population of this fungus in the eastern United States comprised 35 VCGs (Anagnostakis 1992).
Herein we report the results of our investigation of genetic variation among 89 isolates of A. niger from three geographically discrete populations of W. mirabilis (Figure 1). The Welwitschia Wash and Hope Mine sites are 6 km apart and located 500 km south of the Urinindes site. In a preliminary study of vegetative compatibility among the isolates, we found that all isolates (29–30) from within a site were vegetatively compatible. Subsequent interpopulation crosses using five isolates randomly chosen from each population revealed that all isolates from all three populations were vegetatively compatible and that there were no genetic barriers precluding parasexuality (Donovan and Jacobson 2001). All crosses were performed according to a protocol detailed in Jacobson et al. (1993b), in which each isolate is paired with the test isolate and itself (control) on CYM medium (ATCC medium 987). All crosses were conducted two times and interactions were unambiguous, with all testcrosses showing no barriers between the mycelia, as was observed for the control crosses.
The objectives of the study reported here were to describe the genetic variation and diversity within each site and to determine whether patterns exist in the distribution of variation within and between sites. To assess levels of genetic variation, we used randomly amplified polymorphic DNA (RAPD) markers derived by polymerase chain reaction using 10-base primers, which amplify random fragments throughout the genome (Williams et al. 1990). RAPD markers have provided an efficient initial assessment of genomic variation within and between fungal populations and closely related individuals of fungi (e.g., Dalgleish and Jacobson 2005; Jacobson et al. 1993b; McDonald and McDermott 1993), including molecular typing of A. niger and related species (Birch et al. 1995).
If populations of A. niger at our study sites were predominately clonal, we expected to see low levels of genetic variation among isolates from a single plant and also among all isolates from a single site. If, on the other hand, parasexual recombination were occurring among isolates within plants and sites, we expected to see higher levels of genetic variability within plants, although just how high could not be predicted as no work had been done that would indicate the level of genetic variability that parasexuality can produce in a natural population of A. niger (Debets 1998; Taylor et al. 1999). If conidia dispersal was limited to flightless insects, thus limiting gene flow between populations, we expected to see high levels of genetic differentiation between the isolates from geographically distinct sites because P. sexpunctatis is believed to be incapable of crossing the distances between populations (up to 500 km with no intervening W. mirabilis populations for at least 250 km between the northern and southern sites) (Cooper-Driver et al. 2000). In contrast, if this is a wind dispersed, panmictic organism, we expected to see little differentiation between sampling sites.
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Materials and Methods |
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Taxon Description
Two collections of black mold fungi from W. mirabilis seed cones were previously identified as one of the following four taxa: A. niger V. Tiegh., A. phoenicus Corda, a species within the A. niger aggregate, or A. niger var. phoenicus Corda (Al-Musallam 1980; Schutte 1994). In her revision of black Aspergillus species, Al-Musallam (1980) used morphological and cultural characteristics to describe the A. niger aggregate as being composed of six varieties differentiated by conidia roughness and ornamentation. Specifically, the distinguishing features of A. niger var. phoenicus were conidia, 3–4.5 (5) µm in diameter, "usually rough, showing regular surface ornamentation with ridges in longitudinal striation." In 1996, L. Shutte and H. Kolberg identified infections from the Messum River as A. niger var. phoenicus, and as a result Cooper-Driver et al. (2000) referred to the infections they mapped as this taxon. We examined conidia from nine of our isolates (N = 20) from each of the three populations and found they were uniformly globose, 4–5 µm in diameter, irregularly roughened to punctuate, but entirely lacking longitudinal ridges. Thus, our isolates, notably all belonging to the same VCG (Donovan and Jacobson 2001), do not fit Al-Musallam's strict definition of A. niger var. phoenicus and are more appropriately described by the broader description of Raper and Fennell (1965) of the A. niger aggregate. Recent molecular studies (see review by Abarca et al. 2004) of black Aspergillus species have confirmed that the A. niger aggregate is a distinct phylogenetic entity but have not yet resolved taxonomic definitions within the group.
Site Description
The Welwitschia Wash (23°36'54.5''S, 15°10'45''E) and Hope Mine (23°34'273''S, 15°15'533''E) sites are located at the southern edge of the range of W. mirabilis, just north of the ephemeral Kuiseb River in the central Namib Desert, and are approximately 6 km apart (Jacobson and Lester 2003). The Urinindes (19°51'667''S, 13°39'075''E) site is located approximately 500 km north of these sites in the roadless area between the Ugab and Hoanib Rivers. While there are numerous well-known W. mirabilis populations between these northern and southern sites, all intervening northern and southern populations are separated by at least 250 km (Jacobson and Lester 2003).
Collection, Isolation, and Harvesting
Female W. mirabilis cones infected with A. niger were collected in July 1999 from three individual plants at each of the three sites and stored dry in sterile bags. Conidia were isolated by agitating an infected cone scale from each plant in sterile water. The suspended conidia were diluted 100- to 1000-fold and then plated on 100 µl CYM agar and incubated overnight at 20°C. Germinating conidia were isolated using a dissecting microscope, transferred on to separate CYM agar plates, and incubated approximately 1 week at 20°C. Each isolate was subcultured to two CYM agar plates and incubated for 10 days at 20°C. The mycelium and conidia were scraped from the surface of both plates with a sterile razor blade and stored frozen in CTAB buffer at –20°C until DNA was extracted. Nine to ten conidial isolates were isolated from three individual W. mirabilis plants at each of the three sites for a total of 89 isolates.
Molecular Methods
DNA was extracted from the fungal tissue using a CTAB DNA extraction protocol (Gardes and Bruns 1993) followed by use of the Omega Bio-tek (Doraville, GA) EZNA® Fungal DNA Kit to remove the remaining pigment when necessary. The DNA samples were then quantified with a fluorimeter and DNA-specific dye (Hoechst). RAPD reactions (25 µl) consisted of 12 ng genomic DNA, 25 mM MgCl2, 250 µM each of dATP, dTTP, dGTP, and dCTP, 5 units Taq DNA polymerase (Promega, Madison, WI), and 40–50 ng of each 10-mer (Operon Technologies, Alameda, CA). Primers OPA 1–20 were screened to yield a total of 30–40 loci. RAPD amplification was performed in an MJ Research (Watertown, MA) PTC-150 MinicyclerTM under conditions specified by Williams et al. (1990). The RAPD products were separated by agarose gel electrophoresis, and gels were stained with ethidium bromide and photographed under UV illumination using the Kodak Electrophoresis Documentation and Analysis System 290. Gel products were sized relative to a 100-bp DNA ladder standard (100–1500 bp) (Promega, Madison, WI) using Kodak ID Image Analysis software to allow objective and consistent sizing and quantification of bands. In addition to running water controls with each setup, four reactions, using 66 samples, were performed twice to confirm that RAPD markers scored (sizes ranges 200–1700 bp) were reproducible (Hadrys et al. 1992; Perez et al. 1998).
Data Analysis
All RAPD bands were scored as loci with two alleles, with one allele indicated by the presence of a fragment and the other by the absence of a fragment (scored, respectively, as 1 or 0 in the Excel data file). Population structure at the three sites was studied in three ways. Firstly, we used data from all 89 isolates to quantify total gene diversity (HT), the genetic diversity of individuals relative to their subpopulation (HS), and genetic differentiation relative to the total population (GST) (Nei 1973). Each genetic parameter value was corrected for small, unequal sample sizes according to Nei and Chesser (1983). These values were also calculated for 29–30 isolates from each site to further quantify population structure at the site level.
The data set for all 89 isolates was used to create a similarity matrix in NTSYS using the Jaccard algorithm (Rohlf 1988). A phenogram of the resulting matrix was generated using the unweighted pair group method with arithmetic mean. The cophenetic correlation statistic was computed using the COPH function, and the goodness-of-fit of the phenogram for the similarity matrix was calculated using MXCOMP.
Finally, an analysis of molecular variance (AMOVA), which uses haploid molecular variance data rather than marker frequency (Excoffier et al. 1992), was also performed. This analysis was performed on a matrix created using the Euclidean distance algorithm (Liao and Hsiao 1998). AMOVA, which has been used effectively to obtain a first assessment of genetic variation in a number of other fungal species (Gosselin et al. 1999; Huff et al. 1994), was used to partition the total genetic variation into three specific hierarchical levels: among isolates on a W. mirabilis plant, among isolates on different plants at each site, and between sites. Significance levels for the resultant variance components were computed by nonparametric permutation procedures (Excoffier et al. 1992).
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Four primers (OPA2 = 5'-TGCCGAGCTG-3', OPA4 = 5'-AATCGGGCTG-3', OPA10 = 5'-GTGATCGCAG-3', and OPA20 = 5'-GTTGCGATCC-3') yielded 35 unambiguous RAPD loci from the 89 isolates. Four of these loci were invariant for the marker allele in all three populations, and of the remaining 31 loci, 60%–77% were variable within each site (Tables 1 and 2). The frequencies of only five of the 31 loci varied significantly (P < .05) among the three populations (Table 2), and Simpson's index of diversity and the estimate of genotypic evenness (Table 1) both approached values of 1.0. All three sites were composed primarily of isolates with unique genotypes (72%–93%) (Table 1), and despite the intense sampling of isolates within a single plant, only seven pairs of isolates had identical genotypes (Figure 2).
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The analysis of population structure at the site level revealed that genetic diversity within sites accounted for most of the total genetic diversity. Adjusted values of genetic diversity within sites (HS) ranged from 0.02 to 0.49 and averaged 0.22, accounting for 82% of the total genetic diversity (HT). The proportion of the total diversity attributable to differentiation at the site level (Table 2: GST) ranged from 0.01 to 0.47 and averaged 0.14, indicating little population subdivision at the site level.
The analysis of population structure at the plant level (Table 3) further revealed that genetic diversity of isolates within W. mirabilis plants accounted for most of the total genetic diversity within each site. Adjusted values of genetic diversity (HS) within plants averaged 0.26 at Hope Mine, 0.24 at Welwitschia Wash, and 0.22 at Urinindes accounting for, respectively, 83%, 76%, and 67% of the total genetic diversity within each site (HT). In addition, the average GST values for A. niger isolates among the three plants at each site were HM = 0.17, WW = 0.21, and U = 0.25, respectively, indicating low and intermediate levels of population subdivision at even the plant level (Table 3).
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The AMOVA confirmed that most of the total genetic variation was accounted for among the isolates from individual W. mirabilis plants (78% of the total variance; P < .001) but also revealed that the much smaller proportional variances attributed to within-site (12%) and between-site (10%) differences were highly significant (P < .001) (Table 4).
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Further illustrating the relationships among the 89 isolates, the phenogram (Figure 2) showed that few isolates formed site-specific genetic clusters. While 18/29 isolates from the Urinindes site formed a cluster sharing 82% similarity, the other 11 Urinindes isolates clustered with isolates from Hope Mine and Welwitschia Wash sharing up to 90% similarity. Similarly, 16/30 isolates from Welwitschia Wash formed small clusters (two to eight isolates per cluster) sharing 67%–76% similarity, but like the Urinindes isolates, the remaining 14 Welwitschia Wash isolates clustered with Urinindes and Hope Mine isolates with up to 96% similarity (Figure 2).
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Discussion |
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Levels of genetic diversity both among and between isolates from the sampled sites were much higher than expected for an asexual organism occurring on a single host species in isolated locations. These populations exhibited no clonal population structure. In fact, values obtained for total gene diversity (HT), genetic diversity of individuals relative to their subpopulation (HS), and genetic differentiation relative to the total population (GST) are similar to those observed in fungi with known sexual life cycles (e.g., Goodwin et al. 1992, 1993; Gosselin et al. 1999; Johannesson et al. 2001; McDermott and McDonald 1993; Peever and Milgroom 1994).
While the analysis of genotypic diversity, the hierarchical analysis performed by AMOVA, and the cluster analysis all revealed that most of the variation in the population is accounted for among individual fungal isolates, these analyses also revealed significant differentiation among isolates from each of the plants at the three sites and among the three sites. According to the AMOVA, only 12% and 10% of the total genetic variation were accounted for at the between-plant and between-site levels, yet both are highly significant (P < .001), indicating that isolates from each site are appropriately referred to as genetic subpopulations and populations, respectively. In diploid or dikaryotic organisms, regional population structure of this nature is usually the result of more frequent interbreeding events among isolates within a site than between sites (Gryta et al. 2000). Given that A. niger is an exclusively asexual fungus known to readily perform parasexuality in the laboratory (but not previously demonstrated in natural populations), we propose that parasexuality may, in part, account for the extreme levels of variation we observed in these subpopulations and populations.
While fungal populations that reproduce sexually are often more genetically variable than those that reproduce asexually (Burdon and Roelfs 1985), evidence is accruing that many asexual fungi have higher than expected levels of genetic variation and are undergoing some means of genetic exchange. However, only two of these studies have experimentally proved that parasexuality is occurring under controlled field conditions (Souza-Paccola et al. 2003; Zeigler et al. 1997), and studies have assessed neither the degree to which parasexuality occurs in natural populations nor the significance of such asexual horizontal gene transfer as an adaptive mechanism relative to migration and genetic drift (Correll and Gordon 1999).
Previous studies of other asexual Aspergillus species have been unable to make definitive conclusions about the level of genetic variation attributable to parasexual recombination because of uncertainty as to whether variation was caused by recombination or the introduction of inoculum from other populations (Bayman and Cotty 1990). As with these other studies of asexual fungi that exhibit higher than expected levels of variation, the relative contributions of gene flow and parasexual recombination in structuring these A. niger populations are not known. However, the relatively low level of genetic differentiation among all 89 isolates (GST = 0.14) indicates that gene flow is indeed occurring across distances ranging from 6 to 500 km and is homogenizing the populations. Among the 29–30 isolates within each population, our study also suggests that gene flow occurs among subpopulations occupying separate plants (GST = 0.17–0.25). More intensive sampling of fungi at the plant level is needed to determine whether these differences in levels of gene flow within the different sites reflect actual differences in the manner in which these populations are exchanging genetic material.
While the results of a spatial autocorrelation study by Cooper-Driver et al. (2000) favor local transmission by a dispersal agent (e.g., P. sexpunctatis) with a limited range that would account for gene flow within A. niger populations, the authors do not preclude transmission by other biotic and abiotic agents, some with the potential to cross great distances between W. mirabilis populations. An obvious possibility is the wind dispersal of the dark, thick-walled conidia. In addition, other potential agents of transmission include pollinator flies (Wetschnig and Depish 1999); animals which shelter under or feed on W. mirabilis such as Grays lark, snakes, lizards, arthropods, oryx, and springbok; and the tires of four-wheel drive vehicles (Cooper-Driver et al. 2000).
Another potential explanation for this high level of gene flow between sites is that the highly ubiquitous A. niger may be infecting the surrounding vegetation (e.g., Euphorbia, Commiphora, and Arthraerua species, or annual grasses) and dispersing by wind to nearby W. mirabilis plants. Indeed, A. niger is a common soil saprophyte in many environments (Abarca et al. 2004; Raper and Fennel 1965), and while the organic content of the Namib Desert soil is exceptionally low, it is possible that this fungus is a common soil saprophyte here as well. This has not been previously considered because it was assumed that the A. niger variant on W. mirabilis was unique and occupied a small niche on W. mirabilis alone. Alternatively, if the fungus is endemic to W. mirabilis in the Namib Desert, it may be more common in soils and on vegetation in the higher rainfall regions to the east of the Namib Desert. Strongly directional winter east winds could potentially transport conidia from the more mesic inland savanna to the arid coastal desert.
Any efforts taken to control fungal infections of W. mirabilis should bear in mind the high levels of genetic diversity found by this study and the possibility that surrounding soils and vegetation and prevailing winds from upland savannas may be serving as sources of inoculum of this apparently common fungus. Before any control measure can be put in place, efforts should be made to identify all means of conidia dispersal and sources of inoculum, as well as the identity of the pathogen relative to the known phylogeny of this group (Abarca et al. 2004). Given the inherent difficulties in attempting to control such a potentially common and ubiquitous pathogen, further studies should address whether this fungus really is reducing plant recruitment at higher than normal levels. Changes in the size and frequency of the large precipitation events (50 mm) required for seed germination may be an equally important factor affecting plant recruitment. Our observations of plants at 12 sites throughout the plant's range suggest that there have been only three to five recruitment events over the last 150 years resulting in cohorts of plants all of approximately the same size. Healthy populations of W. mirabilis produce seeds each year, and seeds are long lived (at least 10 years—personal observations). It is thus entirely possible that seed recruitment, albeit with a heavy loss to fungal infection, is adequate to take advantage of the uncommon high rain events that facilitate germination and survival.
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Acknowledgments |
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We gratefully acknowledge the Namibian Ministry of Environment and Tourism for granting permission to sample infected cones from W. mirabilis in the Namib-Naukluft and Skeleton Coast Parks. Thanks also to the Desert Research Foundation of Namibia and Mary Seely for critical logistical support for the fieldwork, Grinnell College and the Woodrow Wilson Foundation for financial support, and Thomas Harrington and three anonymous reviewers for helpful comments on the manuscript.
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Footnotes |
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Corresponding Editor: Reid Palmer
Received July 18, 2005
Accepted February 7, 2006
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Abarca ML, Accensi F, Cano J, Cabañes FJ. (2004) Taxonomy and significance of black aspergilli. Antonie Leeuwenhoek 86:33–49.[CrossRef][Medline]
Al-Musallam A. (1980) Revision of the black Aspergillus species (PhD dissertation). (State University Utrecht, Netherlands).
Anagnostakis SL. (1992) Diveristy within populations of fungal pathogens on perennial parts of perennial plants. The fungal community: its organization and role in the ecosystem (Marcel DekkerIn Carroll GC and Wicklow DT (Eds.). , New York)183–192.
Barnard P. (1998) Biological diversity in Namibia: a country study (Namibian National Biodiversity Task Force, Windhoek, Namibia).
Bayman P and Cotty PJ. (1990) Vegetative compatibility and genetic diversity in the Aspergillus flavus population of a single field. Can J Bot 69:1707–1711.
Berbee ML, Payne BP, Zhang G, Roberts RG, Turgeon BG. (2003) Shared ITS DNA substitutions in isolates of opposite mating type reveal a recombining history for three presumed asexual species in the filamentous ascomycete genus Alternaria. Mycol Res 107:2169–182.[Medline]
Birch M, Anderson MJ, Denning DW. (1995) Molecular typing of Aspergillus species. J Hosp Infect 30:339–351.[Medline]
Bornman CH. (1978) Welwitschia, paradox of a parched paradise. (C. Struik, Cape Town, South Africa).
Bornman CH, Elsworthy JA, Butler V, Botha CEJ. (1972) Welwitschia mirabilis: observations on general habit, seed, seedlings, and leaf characteristics. Madoqua Ser II 1:53–66.
Brasier C. (1992) A champion thallus. Nature (London) 356:382–383.
Burdon JJ and Roelfs AP. (1985) The effect of sexual and asexual reproduction on the isozyme structure of populations of Puccinia graminis. Phytopathology 75:1068–1073.
Burt A, Carter DA, Koenig GL, White TJ, Taylor JW. (1996) Molecular markers reveal cryptic sex in the human pathogen Coccidioides immitis. Proc Natl Acad Sci USA 93:770–773.
Bustard L. (1990) The ugliest plant of the world: the story of Welwitschia mirabilis. Kew Mag 7:85–90.
Cooper-Driver GAC, Wagner C, Kolberg H. (2000) Patterns of Aspergillus niger var. phoenicis (Corda) Al-Musallam infection in Namibian populations of Welwitschia mirabilis Hook.f. J Arid Environ 46:181–198.
Correll JC and Gordon TR. (1999) Population structure of Ascomycetes and Deuteromycetes. In Worrall JJ (Ed.). Structure and dynamics of fungal populations (Kluwer Academic Publishers, Dordrecht, The Netherlands) pp. 225–250.
Dalgleish HJ and Jacobson KM. (2005) A first assessment of genetic variation among Morchella esculenta (morel) populations. J Hered 96:31–8.
Debets AJM. (1998) Parasexuality in fungi: mechanisms and significance in wild populations. In Bridge P, Couteaudier Y, Clarkson J (Eds.). Molecular variability of fungal pathogens (Cab International, Wallingford, U.K) pp. 41–52.
Dilcher DL. (2005) Welwitschiaceae from the lower Cretaceous of northeastern Brazil. Am J Bot 92:81294–1310.
Donovan AK and Jacobson KM. (2001) An analysis of genetic variation of three geographic populations of Aspergillus niger var. phoenicus, a pathogen of Welwitschia mirabilis. Proceedings of the Iowa Academy of Science Conference. Des Moines, April 20, 2001; 21 (Iowa Academy of Science, Cedar Falls).
Excoffier L, Smouse PE, Quattro JM. (1992) Analysis of molecular variance inferred from metric distances among DNA haplotypes: applications to human mitochondrial DNA restriction data. Genetics 131:479–491.[Abstract]
Fennell DI. (1977) Aspergillus taxonomy. In Smith JE and Pateman JA (Eds.). Genetics and physiology of Aspergillus (Academic Press, London) pp. 1–22.
Garber ED and Ruddat M. (1992) The parasexual cycle in Ustilago scabiosae (Ustilaginales). Int J Plant Sci 153:98–101.
Gardes M and Bruns TD. (1993) ITS primers with enhanced specificity for basidiomycetes: application to the identification of mycorrhizae and rusts. Mol Ecol 2:113–118.[Medline]
Geiser DM, Pitt JI, Taylor JW. (1998) Cryptic speciation and recombination in the aflatoxin-producing fungus Aspergillus flavus. Proc Natl Acad Sci USA 95:388–393.
Goodwin SB, Saghai-Maroof MA, Allard RW, Webster RK. (1993) Isozyme variation within and among populations of Rhynchosporium secalis in Europe, Australia and the United States. Mycol Res 97:49–58.
Goodwin SB, Spielman LJ, Matuszak JM, Bergeron SN. (1992) Clonal diversity and genetic differentiation of Phytophthora infestans populations in northern and central Mexico. Phytopathology 82:955–961.
Gosselin L, Jobidon R, Bernier L. (1999) Genetic variability and structure of Canadian populations of Chondrostereum purpureum, a potential biophytocide. Mol Ecol 8:113–122.
Gräser Y, Volovsek M, Arrington J, Schönian G, Presber W, Mitchell TG, Vilgalys R. (1996) Molecular markers reveal that populations structure of the human pathogen Candida albicans exhibits both clonality and recombination. Proc Natl Acad Sci USA 93:12473–12477.
Gryta H, Debaud J-C, Marmeise R. (2000) Population dynamics of the symbiotic mushroom Hebeloma cylindrosporum: mycelial persistence and inbreeding. Heredity 84:294–302.[Medline]
Hadrys H, Balick M, Schierwater B. (1992) Applications of randomly amplified polymorphic DNA (RAPD) in molecular ecology. Mol Ecol 1:55–63.[Medline]
Henschel JR and Seely MK. (2000) Long-term growth patterns of Welwitschia mirabilis, a long-lived plant of the Namib Desert (including a bibliography). Plant Ecol 150:7–26.[CrossRef]
Hooker JD. (1863) On Welwitschia, a new genus of Gnetaceae. Trans Linn Soc 24:1–48.
Huff DR, Bunting TE, Plumley KA. (1994) Use of random amplified polymorphic DNA markers for the detection of genetic variation in Magnaprothe poae. Genetics 84:1312–1316.
Jacobson KM, Jacobson PJ, Miller OK Jr. (1993a) The mycorrhizal status of Welwitschia mirabilis. Mycorrhiza 3:13–17.
Jacobson KM and Lester E. (2003) A first assessment of genetic variation in Welwitschia mirabilis Hook. J Hered 94:3212–217.
Jacobson KM, Miller OK, Turner BJ. (1993b) RAPD markers are superior to somatic compatibility tests for discriminating genotypes in natural populations of the ectomycorrhizal fungus Suillus granulatus. Proc Natl Acad Sci USA 90:9159–9163.
Johannesson H, Gustafsson M, Stenlid J. (2001) Local population structure of the wood decay ascomycete Daldinia loculata. Mycologia 93:440–446.
Kers LE. (1967) The distribution of Welwitschia mirabilis. Hook.F. Sven Bot Tidskr 61:97–125.
Lhoas P. (1967) Genetic analysis by means of the parasexual cycle in Aspergillus niger. Genet Res 10:45–61.[Medline]
Liao LC and Hsiao JY. (1998) Relationship between population genetic structure and riparian habitat as revealed by RAPD analysis of the rheophyte Acornis gramineus Soland. (Araceae) in Taiwain. Mol Ecol 7:1275–1281.
Lodge DJ and Leonard KJ. (1984) A cline and other patterns of genetic variation in Cochliobolus carbonum isolates pathogenic to corn in North Carolina. Can J Bot 62:5995–1005.
McDermott JM and McDonald BA. (1993) Gene flow in plant pathosystems. Annu Rev Phytopathol 31:353–373.[CrossRef][ISI]
McDonald BA and McDermott JM. (1993) Population genetics of plant pathogenic fungi. Bioscience 43:5311–319.[CrossRef]
Miedaner T, Schilling AG, Geiger HH. (2001) Molecular genetic diversity and variation for aggressiveness in populations of Fusarium graminearum and Fusarium culmorum sampled from wheat fields in difference countries. J Phytopathol 149:11–12641–648.[CrossRef]
Nei M. (1973) Analysis of gene diversity in subdivided populations. Proc Natl Acad Sci USA 70:3321–3323.
Nei M and Chesser RK. (1983) Estimation of fixation indices and gene diversities. Ann Hum Genet 47:253–259.[ISI][Medline]
Paccola-Meirelles LD, Valarini MJ, Azevedo JL, Alfenas AC. (1991) Parasexuality in Beauveria bassiana. J Invertebr Pathol 57:172–176.[CrossRef]
Peever TL and Milgroom MG. (1994) Genetic structure of Pyrenophora teres populations determined with random amplified polymorphic DNA markers. Can J Bot 72:915–923.
Perez T, Albornoz J, Dominguez A. (1998) An evaluation of RAPD fragment reproducibility and nature. Mol Ecol 7:1347–1357.[CrossRef][Medline]
Pontecorvo G. (1956) The parasexual cycle in fungi. Annu Rev Microbiol 10:393–400.[CrossRef][ISI][Medline]
Raper K and Fennell D. (1965) The genus Aspergillus (Williams and Wilkins Company, Baltimore, MD).
Rohlf FJ. (1988) NTSYS-pc, numerical taxonomy and multivariate analysis system version 1.40 (Exeter, Setauket, NY).
Schutte AL. (1994) An overview of Aspergillus (Hyphomycetes) and associated teleomorphs in South Africa. Bothalia 24:171–85.
Souza-Paccola EA, Fávaro LCL, Casela CR, Paccola-Meirelles LD. (2003) Genetic recombination in Colletotrichum sublineolum. J Phytopathol 151:329–334.[CrossRef]
Taylor JW, Jacobson DJ, Fisher MC. (1999) The evolution of asexual fungi: reproduction, speciation and classification. Annu Rev Phytopathol 37:197–246.[CrossRef][ISI][Medline]
von Willert DJ and Wagner-Douglass U. (1994) Water relations, CO2 exchange, water use efficiency and growth of Welwitschia mirabilis Hook. f. in three contrasting habitats of the Namib Desert. Bot Acta 107:291–299.
Wetschnig W and Depisch B. (1999) Pollination biology of Welwitschia mirabilis Hook.f. (Welwitschiaceae, Gnetopsida). Phyton 39:167–183.
Williams JG, Kubelik AR, Livak KJ, Rafalski JA, Tingey SV. (1990) DNA polymorphisms amplified by arbitrary primers are useful as genetic markers. Nucleic Acids Res 18:6531–6535.
Zeigler RS, Scott RP, Leung H, Bordeos AA, Kumar J, Nelson RJ. (1997) Evidence of parasexual exchange of DNA in the rice blast fungus challenges its exclusive clonality. Phytopathology 87:3284–294.
Zhong SH and Steffenson BJ. (2001) Virulence and molecular diversity in Cochliobolus sativus. Phytopathology 91:5469–476.
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