| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
Department of Microbiology, Colorado State University, Fort Collins, Colorado, 80523-1677 USA
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
We developed a molecular method for the detection and quantification of members of the genus Spizellomyces in the environment and used this technique, together with traditional cultural techniques, to measure the effects of cultivation and nitrogen availability on Spizellomyces populations in grassland soils. Primer sets specific for Spizellomyces acuminatus and S. kniepii were developed by sequencing internal transcribed spacer 2 (ITS2) of the gene encoding ribosomal RNA for 9 isolates within the genus Spizellomyces, 5 representatives of different genera within the order Spizellomycetales and one member of the order Chytridiales. These primers were used with fungal-specific primers in a nested PCR approach to generate a specific molecular signal for S. acuminatus and S. kneipii in a soil from which S. acuminatus had previously been recovered. Using MPN-PCR (a quantitative molecular technique) and traditional cultural techniques, we found that chytridiomycetous fungi, including members of the genus Spizellomyces, are abundant in the grassland ecosystems studied. No significant differences in occurrence were observed between native and disturbed control soils but it appeared in 2 separate MPN assays and one MPN-PCR assay that chytrid populations increased in response to disturbance. No significant differences in chytrid or Spizellomyces populations were observed with variations in nitrogen availability. The primer sets and protocols developed in this study worked well to complement traditional cultural data to better assess Spizellomyces populations in the environment. These molecular approaches should provide a foundation for further work with these interesting and oft neglected fungi.
Key words: Chytridiomycota, disturbance, ITS, MPN-PCR, quantification, soil
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
, Ross and Ruttencutter 1997, Berger et al 1998
). Spizellomyces is a genus in this phylum, consisting of 8 species that are very closely related to the genera Kochiomyces, Gaertneriomyces, and Powellomyces (Barr 1984
). Spizellomyces species are widespread in soils (Booth 1971
) and have been reported to parasitize the oospores of Sclerospora sorghi (Kenneth et al 1975
), Peronspora tabacina (Person et al 1955
), the chytrid Synchytrium endobioticum (Karling 1942
), and soil nematodes (Kenneth et al 1975
). In addition, Spizellomyces infects the spores of arbuscular mycorrhizal fungi (AMF) (Ross and Ruttencutter 1977
, Daniels and Menge 1980
, Daniels 1981
) and has been hypothesized to reduce AMF populations in the field (Daniels and Menge 1980
). Spizellomyces kniepii also has been found to transmit spiroplasma-like organisms between AMF spores, indicating that Spizellomyces species may serve as vectors for viruses (Tzean et al 1983
). Members of this genus also are active saprophytes in nature, colonizing recalcitrant substances such as pine pollen in laboratory experiments. Despite the varied and important roles that these organisms may play in the soil environment (Lozupone and Klein l999
), little is known about their ecology or abundance in nature.
Most quantitative ecological studies of the Chytridiomycota have been carried out using cultural and microscopic protocols based on baiting techniques. In these procedures, an environmental sample, if it is not a liquid, is flooded with sterile water and a "bait" substrate is added, such as pollen or cellophane, which is known to attract chytrids. After several days of incubation, these baits are viewed for the presence of chytrid sporangial growth. This approach was used by Willoughby (1998)
to quantify the monocentric soil chytrid Rhizophlyctis rosea by counting the thalli that grew on cellophane bait in flooded soil samples. Chytrids have also been quantified in soil by means of Most-Probable-Number (MPN) techniques where samples are diluted and a bait is added to the dilutions. The chytrid population is then estimated based on the highest dilution at which these organisms are observed. The MPN technique was first applied to the analysis of chytrids by Gaertner (1968)
, and this approach has been used in several subsequent studies with mixed results (Booth and Barrett 1976
).
Although quantitative procedures based on baiting have provided some knowledge of chytrid presence and abundance in the environment, this approach has a series of limitations. First, chytrid zoospores may not be able to locate or colonize the bait if there is competition from other microorganisms (Willoughby 1962
). In addition, baiting may lead to an underestimation of the actual population size because zoospores emerging from the soil samples also will colonize materials already present in the sample, rather than the experimentally added bait (Willoughby 1998
). Finally, baiting techniques may be of limited value for species that are not easily recognizable or well characterized in terms of the baits to which they are attracted.
An approach which may make it possible to circumvent the problems associated with baiting in quantitative ecological studies of the Chytridiomycota, is a technique called MPN-PCR (Picard et al 1992
). In this technique, primers specific for an organism of interest are used to determine the minimum concentration at which that organism's genetic material can be detected in serially diluted DNA extracts. MPN statistics are then used to estimate the concentration of the particular molecular sequence per unit of sample. By correcting for gene copy number and detection limits, MPN-PCR results can be standardized to provide an estimate of possible organism numbers. MPN-PCR has been used in previous studies to quantify the indigenous populations of Frankia spp. in soils (Picard et al 1992
), to assess the population dynamics of Alnus-infective Frankia in a forest soil with and without host trees (Myrold and Huss-Danell 1994
), and to quantify the staphylococcal enterotoxin c1 gene from fresh cheese (Mäntynen et al 1997
).
The genes encoding ribosomal RNA molecules have been useful for selecting group-specific primers that can be used in MPN-PCR, as has been shown in several studies (Tooley et al 1997
, Schweiger and Tebbe 1998
). These genes, often found as a single tandem array of 50–220 copies per haploid genome (Berthier et al 1996
, Lott et al 1998
), encode a 35S RNA precursor that is subsequently cleaved to form mature 18S, 5.8S, and 28S ribosomal RNA units. These subunits are separated by non-coding regions known as internal transcribed spacer (ITS) regions. The regions between the 18S and 5.8S regions and the 5.8S and 28S regions are called ITS1 and ITS2, respectively. In eukaryotic organisms, particularly the fungi, ITS1 and ITS2 have been useful for finding genus- and species- specific markers because of a high degree of variability (Carbone and Kohn 1993
, Berthier et al 1996
, Lott et al 1998
).
Despite advances in the application of molecular techniques in microbial ecology, the use of such techniques has been limited for the phylum Chytridiomycota because little sequence information is available. Before this study, for instance, only one sequence of the internal transcribed spacer regions (ITS1, 2) was available for any member of the phylum Chytridiomycota, for an isolate of Olpidium brassicae (Ward and Adams 1998
), and no sequence information was available on this region for the genus Spizellomyces.
The objectives of this study were (i) to develop a molecular method to quantify members of the genus Spizellomyces in soil and (ii) to use this method, along with traditional cultural techniques, to evaluate the possible effects of cultivation and nitrogen availability on chytrid and Spizellomyces populations in a semiarid grassland ecosystem.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
).
|
was used to extract DNA from all fungal cultures. Zoospores were obtained by flooding agar plates with sterile distilled water, waiting 20 to 30 min for sporangia to release zoospores, and retrieving zoospores via centrifugation. Alternatively, sporangia were retrieved by removing a section of agar supporting abundant fungal growth with a flame-sterilized loop.
The ITS2 region of rDNA was amplified using ITS3 (5' GCATCGATGAAGAACGCAGC 3') as a forward primer and ITS4 (5' TCCTCCGCTTATTGATATGC 3') as a reverse primer (White et al 1990
). The ITS3 primer is in a conserved domain 128 bp from the 3' end of the 5.8S rDNA and the ITS4 primer is in a conserved domain 59 bp from the 5' end of 28S (White et al 1990
). PCR was carried out for 33 cycles of 95 C for 45 s, 55 C for 30 s, and 72 C for 45 s with a final extension at 72 C for 7 min. Taq polymerase was added after an initial 3- min incubation at 95 C. The amplified DNA was purified through Qiagen columns (PCR purification kit, Qiagen Corp. Chatsworth, California) and sequenced at the Molecular Resources Facility [MRF] (Colorado State University, Fort Collins, Colorado) or Davis Sequencing (University of California, Davis, California).
Sequences were aligned by means of the default settings of Clustal W (gap opening penalty = 15, Gap extension penalty = 6.66, DNA transitions weight = 0.50) (Thompson et al 1994
). Mfold 3.0 was used to determine the secondary-structure of the entire sequence to aid in proper sequence alignment (Zuker et al 1999
). The location of the ITS2 region was determined based on homology of the 5.8S and 28S regions with other fungi. Similarity scores of the ITS2 region were calculated with the aid of Genedoc (Nicholas et al 1997
) and represent the percent identical bases between two aligned sequences; the 5.8S and 28S regions were removed prior to processing. The sequence alignment was used to select two primers from the 3' end of the ITS2 region that were each specific for both Spizellomyces acuminatus and Spizellomyces kniepii and designated S261 and S305.
Spizellomyces acuminatus CL5.2, S. kniepii BR351, S. palustris JL148, S. sp. JL199, and S. plurigibbosus JL210 (Table I
) were used to assess primer specificity. Fungal DNA extracts were amplified in three separate PCR reactions. In the first reaction, the fungal-specific primer ITS5 (5' GGAAGTAAAAGTCGTAACAAGG 3') was used as a forward primer (White et al 1990
) and primer S261 was used as a reverse primer. In the second reaction, primer ITS5 was again used as a forward primer and primer S305 was used as a reverse. In the third reaction, the ITS3 and ITS4 primers, which amplify the ITS2 region of all fungi, were used to verify that each DNA extract was amplifiable by PCR. PCR cycle conditions described earlier were used in all three PCR reactions. PCR conditions were optimized for S261 and S305 specific primers by keeping all concentrations constant and varying the annealing temperature. The optimum temperature that provided the desired specificity and the strongest possible level of amplification was noted.
Primer specificity was also evaluated by amplifying DNA extracts from a soil from which S. acuminatus had been previously isolated and noting if only one band of the correct molecular weight resulted from the amplification process. DNA was extracted from the soil samples with the aid of a mini bead-beater (Biospec Products, Bartlesville, Oklahoma) and a fast DNA spin kit for soil (BIO101, Vista, California), and stored at -20 C. Fungal-specific primers were used to verify that the soil DNA extract was amplifiable by PCR. DNA amplification employing both the ITS5/S261 and ITS5/S305 primer pairs was then attempted from soil extracts with the PCR cycle conditions described earlier and an annealing temperature of 56 C. Due to poor amplification of Spizellomyces DNA with both the ITS5/S261 and ITS5/S305 primer pairs, a nested procedure was used. In this procedure, soil DNA extract was first amplified with S305 and ITS5 primers and the reaction product (1.5 µL) was then amplified with internal primers S261 and ITS3. The first PCR product was filtered through a QIAquick PCR purification column (QIAGEN, Valencia, California) in order to remove the primers from the reaction mixture prior to adding 1.5 µL to the second PCR reaction.
Soils were collected from two sites located on or near the Central Plains Experimental Range (CPER), 50 km northeast of Fort Collins, Colorado (Paschke et al 2000
). These sites are being used in a larger investigation of the effects of nitrogen availability on plant communities and of microbial development during secondary succession. Both study sites are shortgrass steppe ecosystems where the mean annual temperature is approximately 17 C, the approximate elevation is 1650 m and annual precipitation is approximately 310 mm (Klein et al 1996
). The first study site is a native area which has never been disturbed by cultivation; it has vegetation dominated by perennial grasses (26%), succulents (52%), and lichens (13%) (Klein et al 1996
). The second site is a disturbed, early-seral site that was last cultivated in 1989. Its vegetation mainly consists of exotic annual forbs (Klein et al 1996
). Within the native and disturbed sites, twelve 10 x 10 m sub-plots with a 2 m buffer zone have been subjected to various treatments. In both the native and disturbed plots, four control sub-plots were established which have been left unaltered. In addition, 4 sub-plots within both plots have been amended with ammonium nitrate at 100 kg N ha-1 yr-1 added in three increments over the growing season since 1986. These nitrogen amendments have caused a significant increase in N-availability in both the native and disturbed site soils (Klein et al 1996
).
Soil samples were collected during August 1998, and June 1999. Seven soil cores were taken from each sub-plot at each site. Soil cores for each sub-plot were taken from randomly generated coordinates and pooled into a single sample. The cores were 2.54 cm in diameter and taken to a depth of 20 cm with the aid of a rapid drill corer (Klein Environmental Assoc., Fort Collins, Colorado). These cores were gently broken apart, passed through a standard sieve with 8 meshes per inch, and stored at 4 C for 2–4 d before being used to measure chytridiomycete and Spizellomyces populations by means of cultural MPN techniques. A portion of the soil sample from each site also was stored at -80 C for later DNA extraction.
A cultural MPN procedure was used to quantify the pollen-metabolizing chytrid population in samples taken in August 1998 and the Spizellomyces populations alone in samples taken July 1999. With the August 1998 samples, three replicate soil samples from each of 4 treatments were analyzed: (i) native, control, (ii) native, nitrogen-treated, (iii) disturbed, control, and (iv) disturbed, nitrogen-treated. The samples were diluted in a 10-fold dilution series in autoclaved test tubes containing water made up of 1 part filtered, autoclaved pond water and 2 parts sterile, deionized water. Dilute pond water was used based on the observation that pure deionized water alone creates a toxic environment for chytridiomycete zoospores (Emerson 1958
). The soil dilutions were then mixed by repeated pipetting with a sterile electronic pipettor, transferred to small sterile petri dishes and amended with pollen, a substrate known to attract many different species of chytrids. After 4–5 d of incubation at room temperature, the pollen was transferred to a slide and examined under a light microscope for chytrid sporangia. The most probable number and a 95% confidence interval were calculated with a computer program (Klee 1993
).
Cultural MPN was performed on June 1999 samples, as described above, except (i) a five-fold dilution series was used with an initial 1: 40 dilution in order to conform to the MPN-PCR dilution series, (ii) 2 replicates of each treatment were used instead of 3, and (iii) only Spizellomyces sporangia inhabiting the pollen were noted.
MPN-PCR was used to estimate the number of S. acuminatus and S. kniepii genetic units in soils collected in June 1999. DNA extracted from all soil samples was serially diluted through a 5-fold dilution series and PCR was carried out with the nested PCR procedure described earlier. The presence or absence of an amplification product was noted and a computer progam (Klee 1993
) was used to determine the MPN and a 95% confidence interval for the number of ITS2 Genetic Units (GU) of Spizellomyces acuminatus and S. kniepii in each sample. Pairwise Student's t-tests were used for comparisons between soil treatments. MPN and confidence interval estimates, which are Poisson distributed (Cochran 1950
), were log-transformed prior to statistical analysis. Standard errors were derived from the confidence interval estimates and the degrees of freedom were estimated as one less than the number of replicate MPN tubes.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
). Spizellomyces isolate sequences were first aligned to each other, and the remaining isolates were aligned individually to this alignment. This method created an alignment that corresponded closely to the secondary structures of these molecules as determined by Mfold 3.0 (Zuker et al 1999
). The location of the ITS2 region was determined based on homology of the 5.8S and 28S regions with other fungi. Overall, the ITS2 region was found to be extremely variable, whereas the 5.8S and 28S regions were highly conserved. Within the genus Spizellomyces, the size of the ITS2 region ranged from 235 to 245 base pairs (bp). All sequences have been deposited in Genbank with accession numbers AF216757–AF216771. Similarity scores calculated for the ITS2 region for these isolates are noted in Table II ; scores represent the percent of residues that match exactly between two aligned sequences. The highest similarity scores were between the two isolates of S. plurigibbosus, JL210 and BR33, at 95%, and between S. acuminatus CL5.2 and S. kniepii BR351 at 83%. Similarity scores between other isolates within the genus Spizellomyces ranged from 45% to 67% and between Spizellomyces species and members of the closely related genera Kochiomyces, Gaertneriomyces, and Powellomyces, these values ranged from 39 to 48%. Similarity between Spizellomyces species and the 2 isolates of Rhizophlyctis rosea and Rhizophidium were very low (24 to 40%) and similarity between the two isolates of R. rosea was only 30%.
|
|
|
|
= 0.05, but there appeared to be more pollen-metabolizing chytrids in the disturbed-control plot than in the native-control plot (P < 0.2) and in the disturbed-nitrogen amended plot versus the native-nitrogen amended plot (P < 0.2).
|
= 0.05, but the Spizellomyces populations appeared to increase in response to disturbance in the control plots (P < 0.1).
The MPN-PCR approach was used to estimate the mean number of genetic units (GU) of Spizellomyces acuminatus and S. kniepii per gram of soil. The results of this analysis are listed in Table V
. There were between 1704 and 6783 Spizellomyces Genetic Units (GU) estimated per gram for the four soil treatments based on MPN-PCR analysis. There were no significant differences among the soils at = 0.05. The population of S. acuminatus and S. kniepii, however, appeared to increase in response to disturbance in the control plots (P < 0.2).
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
) rather than more variable morphological characteristics. Sequences were generated for 7 of the 8 species in this genus as well as for isolate JL199, which did not match any of Barr's descriptions but is most similar to S. palustris in the ITS2 region (67%) and probably represents a new species. ITS2 sequence data within the genus Spizellomyces were quite variable, indicating that the genus Spizellomyces is genetically diverse despite its morphological similarity. Similarity scores of 95%, as seen between S. plurigibbosus JL210 (from Puerto Rico) and BR33 (from Ontario, Canada), indicate that this region is strongly conserved within Spizellomyces species, whereas similarity scores ranging from 45% to 83% observed between species within the genus Spizellomyces indicate that this region is highly variable for this genus. In comparison, the ITS2 region of two isolates of Rhizophlyctis rosea, which is in the same order as Spizellomyces, were only 30% similar. Chytrids with morphological characteristics which lead the authors to identify them as Rhizophlyctis rosea were found to have diverse zoospore ultrastructure with four distinct sub-types, indicating a potential species complex (Barr and Désaulniers 1986). Problems in classification of the Chytridiomycetes, as illustrated here with R. rosea, can make the development of specific primers and the application of MPN-PCR difficult for such poorly defined species.
Conservation of the ITS2 region within species for the genus Spizellomyces, as well as variation between species, makes this region ideal for the development of specific primers for this group, as carried out in this study. The observed conservation within species is consistent with previous studies, in which ITS regions were used for development of PCR primers for Phytophthora species infecting potatoes (Tooley et al 1997
), Polymyxa species (Ward and Adams 1998
), and Alexandrium species (Penna and Magnani 1999
).
When DNA extracts from soil materials were amplified with the species-specific primers in a nested fashion, initially 2 bands of similar size resulted. This second band was initially suspected to be a non-specific amplification product from the soil DNA extract. Removal of the primers from the first PCR product, before it was added to the second PCR reaction, however, led to the detection of only 1 band, indicating that this extraneous band was a PCR artifact, and not non-specific amplification. Nested PCR was shown to increase both the sensitivity and the specificity of the PCR reaction, allowing a clear and specific signal for Spizellomyces acuminatus and S. kniepii to be detected from the soils.
The Spizellomyces population estimates generated in this study using MPN (135 and 673 thalli per gram for the native and disturbed shortgrass step soils, respectively) are similar in magnitude to the population estimates that were made for Rhizophlyctis rosea in a grassland soil using cellophane bait (395 thalli per gram of soil) by Willoughby (1998)
. The estimated number of pollen-metabolizing chytrids per gram of soil was significantly higher, at 1315 thalli in the native control plot and 4605 thalli in the disturbed control plot, indicating that Spizellomyces species only make up a small fraction of the entire chytrid community. In addition, certain chytrids are not attracted to pollen, indicating that the actual chytrid population may be even larger. Also, total chytrid and Spizellomyces spp. populations may be even further underestimated by this cultural MPN approach, because chytrid zoospores may not be able to locate or colonize the bait if there is competition from other microorganisms (Willoughby 1962
). In spite of these procedural concerns, the population estimates of chytrids obtained in this study are quite high, indicating that chytrids are present in high numbers and, therefore, may have important effects on this complex plant-soil ecosystem.
The MPN-PCR data provided an estimate of the mean number of rRNA genetic units per gram of soil. These data do not indicate actual population sizes because the rRNA operon is present in multiple copies in the eukaryotic genome. The rRNA genome has been estimated to contain ranging from 50 to 220 copies of rRNA for various fungal taxa (Berthier et al 1996
, Lott et al 1998
). The estimated numbers of Spizellomyces genetic units in the shortgrass steppe soils are quite high (between 1704 and 6783 for the four soil treatments). This gives additional evidence that Spizellomyces are abundant in these soils.
Although few of the results of this study were statistically significant, both MPN experiments and the MPN-PCR analysis of Spizellomyces populations in shortgrass steppe soils showed a population increase in response to disturbance. This possible increase in chytrid population density is in contrast to what has been observed for filamentous fungi in these same soils. Earlier research revealed that communities of filamentous fungi in the native sites are larger and less active than in disturbed sites (Klein 1996
, Klein and Paschke 2000
). The possible increase in chytrid populations, however, is not surprising given the life-cycle of this group. Disturbance, especially by cultivation, often increases the concentration of biologically available nutrients in soil. Chytrids can rapidly reproduce, releasing highly chemotactic zoospores when conditions are favorable. These characteristics could allow them to quickly exploit the nutrients released in response to disturbance, increasing their overall population, as observed in this study and as noted for Rhizophlyctis by Harder (1948)
.
It is hoped that this work is a first attempt that will lead to a greater use of molecular techniques for the study of zoosporic fungi in nature. One potential weakness of both cultural MPN and MPN-PCR is that neither technique differentiates between active and non-active chytrids. The DNA extraction techniques that were used in this study will retrieve DNA from chytrid spores as well as active chytrids, and enrichment techniques, such as baiting, provide a favorable environment that may cause spore release. Quantification techniques that are capable of differentiating between active and non-active chytrids in the environment need to be developed, and these approaches will represent an important aspect of chytridiomycete ecology in the future.
Future work should include standardizing the MPN-PCR technique for gene-copy number and PCR detection limits so that it can be used to estimate actual organism numbers rather than genetic units. This can be done by adding known numbers of zoospores to a soil of similar quality, which does not contain natural populations of S. acuminatus or S. kniepii, and comparing the actual population size to the size predicted using MPN-PCR.
The primers developed for S. acuminatus and S. kniepii could be applied to many different environments to address a wide range of ecological questions. In addition, information on ITS2 sequences provided in this study, as well as the observation that the ITS2 region is highly conserved within these species, should provide a foundation for the development of additional primer sets for other members of the genus Spizellomyces, and also for related groups of motile fungi. These should make it possible to carry out more detailed molecular-technique based studies of these interesting and largely unappreciated fungi. This approach, as a complement to cultural quantification techniques, should lead to a better understanding of the distribution and abundance of these Spizellomyces species, and assist in developing a better understanding of other chytridiomycetous fungi in nature.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
Accepted for publication October 1, 2001.
|
LITERATURE CITED |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
Barr DJS., Desaulniers NL., 1985 Four zoospore subtypes in the Rhizophlyctis-Karlingia complex (Chytridiomycetes) Can J Bot 64:561-572
Berger L, Speare R, Daszak P, Green DE, Cunningham AA, Goggin CL, Slocombe R, Ragan MA, Hyatt AD, McDonald KR, Hines HB, Lips KR, Marantelli G, Parkes H., 1998 Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America Proc Nat Acad Sci USA 95:9031-9036
Berthier YT, Bruckart WL, Chaboudez P, Luster DG., 1996 Polymorphic restriction patterns of ribosomal internal transcribed spacers in the biocontrol fungus Puccinia carduorum correlate with weed host origin Appl Environ Microbiol 62:3037-3041[Abstract]
Black WC, Duteau NM., 1997 RAPD-PCR and SSCP analysis for insect population genetic studies In: Crampton JM, Beard CB, and Louis C, eds. The molecular biology of insect disease vectors. A methods manual. New York: Chapman and Hall. p 361–371
Booth T., 1971 Distribution of certain soil inhabiting chytrid and chytridiaceous species related to some physical and chemical factors Can J Bot 49:1743-1755
Booth T., Barrett P., 1976 Taxonomic and ecologic observations of zoosporic fungi in soils of a high-arctic ecosystem Can J Bot 54:533-538
Carbone I, Kohn LM., 1993 Ribosomal DNA sequence divergence within internal transcribed spacer 1 of the Scleroyiniaceae Mycologia 85:415-427
Cochran WG., 1950 Estimation of bacterial densities by means of the "most probable number" Biometrics 6:105-116[Medline]
Daniels BA., 1981 The influence of hyperparasites of vesicular-arbuscular mycorrhizal fungi on growth of citrus Phytopathology 71:212-213
Daniels BA., Menge JA., 1980 Hyperparasitism of vesicular-arbuscular mycorrhizal fungi Phytopathology 70:584-588
Emerson R., 1958 Mycological organization Mycologia 50:589-621
Gaertner A., 1968 Eine Methode des quantitativen Nachweises niederer, mit pollen koderbarer Pilze im Meerwasser und im Sediment Veroff Inst Meeresforsch Bremerhaven 3:75-92
Harder R., l948 Über das Vorkommen niederer Phycomyceten in deutschen Böden Nachrichten Akad. Wiss. Göttingen, Math.-Physik Kl 1948:5-7
Karling JS., 1942 Parasitism among the chytrids Am J Bot 29:24-35
Kenneth RE, Cohn E, Shahor G., 1975 A species of Phlyctochytrium attacking nematodes and oospores of downy mildew fungi Phytoparasitica 3:70
Klee AJ., 1993 A computer program for the determination of most probable number and its confidence limits J Microbiol Methods 18:91-98
Klein DA, McLendon T, Paschke MW, Redente EF., 1996 Nitrogen availability and fungal-bacterial responses in successional semiarid steppe soils Arid Soil Restoration and Rehabilitation 10:321-332
Klein DA, Paschke MW., 2000 A soil microbial community structural-functional index: the microscopy-based total/active fungal/bacterial (TA/AFB) biovolumes ratio Appl Soil Ecol 14:257-268
Lott TL, Burns BM, Zancope-Oliveira R, Elie CM, Reiss E., 1998 Sequence analysis of the internal transcribed spacer 2 (ITS2) from yeast species within the genus Candida Curr Microbiol 36:63-69[Medline]
Lozupone CA, Klein DA., 1999 Chytridiomycota of little importance in soils? ASM News 65:662-663
Mäntynen V, Niemela S, Kaijalainen S, Pirhonen T, Lindstrom K., 1997 MPN-PCR—quantification method for staphylococcal enterotoxin c1 gene from fresh cheese Int J Food Microbiol 36:135-143[Medline]
Myrold DD, Huss-Danell K., 1994 Population dynamics of Alnus-infective Frankia in a forest soil with and without host trees Soil Biol Biochem 26:533-540
Nicholas KB, Nicholas HB Jr.,, Deerfield DW II., 1997 Genedoc: analysis and visualization of genetic variation EMBNEW NEWS 4:14
Paschke M W, McLendon T, Redente E F., 2000 Nitrogen availability and old-field succession in a shortgrass steppe Ecosystems 3:144-158[Medline]
Penna I, Magnani M., 1999 Identification of Alexandrium (dinophyceae) species using PCR and rDNA-targeted probes J Phycol 35:615-621
Person LH, Lucas GB, Koch WJ., 1955 A chytrid attacking oospores of Peronospora tabacina Pl Dis Reporter 39:887-888
Picard C, Ponsonnet C, Paget E, Nesme X, Simonet P., 1992 Detection and enumeration of bacteria in soil by direct DNA extraction and polymerase chain reaction Appl Environ Microbiol 58:2717-2722
Powell MJ., 1993 Looking at mycology with a janus face: a glimpse at Chytridiomycetes active in the environment Mycologia 85:1-20
Ross JP, Ruttencutter R., 1977 Population dynamics of two vesicular-arbuscular endomycorrhizal fungi and the role of hyperparasitic fungi Phytopathology 67:490-496
Schwieger F, Tebbe CC., 1998 A new approach to utilize PCR-single-strand-conformation polymorphism for 16S rRNA gene-based microbial community analysis Appl Environ Microbiol 64:4870-4876
Thompson JD, Higgins DG, Gibson TJ., 1994 CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice Nucl Acids Res 22:4673-4680
Tooley PW, Bunyard BA, Carras MM, Hatziloukas E., 1997 Development of PCR primers from Internal Transcribed Spacer Region 2 for detection of Phytopthora species infecting potatoes Appl Environ Microbiol 63:1467-1475[Abstract]
Tzean SS, Chu CL, Su HJ., 1983 Spiroplasma-like organisms in a vesicular-arbuscular mycorrhizal fungus and its mycoparasite Phytopathology 73:989-991
Ward E, Adams MJ., 1998 Analysis of ribosomal DNA sequences of Polymyxa species and related fungi and the development of genus- and species- specific PCR primers Mycol Res 102:965-974
White TJ, Bruns TD, Lee SB, Taylor JW., 1990 Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics In Innis MA, Gelfand DH, Sninsky JJ, White TJ, eds. PCR protocols: a guide to methods and applications. New York: Academic Press. p 315–322
Willoughby LG., 1962 The occurrence and distribution of reproductive spores of Saprolegniales in fresh water J Ecol 50:733-759
Willoughby LG., 1998 A quantitative ecological study on the monocentric soil chytrid, Rhizophlyctis rosea, in Provence Mycol Res 102:1338-1342
Zuker M, Mathews DH, Turner DH., 1999 Algorithms and thermodynamics for RNA secondary structure prediction: a practical guide In: Barciszewski J, Clark BFC, eds. RNA biochemistry and biotechnology. NATO ASI Series 3. High Technology. Vol. 70. Norwell, Massachusetts: Kluwer Academic Publishers. p. 11–43
This article has been cited by other articles:
 |
|
 |
|
  A. J Scupham, L. L. Presley, B. Wei, E. Bent, N. Griffith, M. McPherson, F. Zhu, O. Oluwadara, N. Rao, J. Braun, et al. Abundant and Diverse Fungal Microbiota in the Murine Intestine Appl. Envir. Microbiol., January 1, 2006; 72(1): 793 - 801. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
