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USDA-Agricultural Research Service, Systematic Botany and Mycology Lab, 10300 Baltimore Ave., Beltsville, Maryland 20705
W. Phillips-Mora
Cacao Breeding Program, CATIE 7170, Turrialba, Costa Rica
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONTAXONOMYLITERATURE CITED |
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The two most devastating diseases of cacao (Theobroma cacao)—the source of chocolate—in tropical America are caused by the fungi Crinipellis perniciosa (witches’ broom disease) and Moniliophthora roreri (frosty pod rot or moniliasis disease). Despite the agricultural, socio-economic and environmental impact of these fungi, most aspects of their life cycles are unknown, and the phylogenetic relationships of M. roreri have yet to be conclusively established. In this paper, extensive phylogenetic analyses of five nuclear gene regions (28S rDNA, 18S rDNA, ITS, RPB1, and EF1-
) confirm that C. perniciosa and M. roreri are sister taxa that belong in the Marasmiaceae (euagarics). Furthermore, these taxa form part of a separate and distinct lineage within the family. This lineage includes the biotrophic fungi Moniliophthora perniciosa comb. nov. and M. roreri, as well as one undescribed endophytic species. The sister genera to Moniliophthora are Marasmius, Crinipellis and Chaetocalathus, which consist mainly of saprotrophic litter fungi.
Key words: anamorphic basidiomycetes, cacao pathogens, cocoa, fungal taxonomy, molecular systematics
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONTAXONOMYLITERATURE CITED |
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). Both fungi are indigenous to tropical South America and attack species of Theobroma L. and the closely-related genus Herrania Goudot, in particular, T. cacao, the source of chocolate. Both are able to disrupt host hormonal balance, provoking hypertrophy and hyperplasia followed by tissue necrosis (Holliday 1980). Moreover, the intercellular mycelium of both fungi is typically swollen and convoluted in marked contrast to mycelium growing intracellularly or in standard culture (Evans 1981). Together these diseases have caused relatively recent devastation to major cacao-farming regions in South and Central America (Pereira 1999, Phillips-Mora 2003), and are responsible for the plummet in tropical American cocoa production. Despite the socio-economic and environmental impact of these fungi, aspects of their life cycles, including mode of reproduction, have yet to be resolved, and phylogenetic relationships remain uncertain.
Initial infection by C. perniciosa basidiospores occurs in actively growing cacao meristems, causing a characteristic disorganized proliferation of new shoots in the host that are termed "witches’ brooms" (Isaac et al 1993) (FIG. 1a). Potential crops are lost when clusters of flowers produced on "cushions" on the main trunk and older branches are infected, thus producing seedless strawberry- or carrot-shaped fruits (Pereira 1999). Additionally, C. perniciosa attacks cacao pods in the early stages of development, penetrating the husk and destroying the seeds from which chocolate is derived (FIG. 1b). After death of the broom tissue, fructifications of small pink agarics occur, which Stahel (1915) identified as the causal agent of the disease and named Marasmius perniciosa Stahel. At that time the disease had been known for approximately 20 years in northern South America (Stahel 1915).
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). By 1980 the pathogen had continued to spread into Colombia, Peru, and Venezuela (Holliday 1980). The cocoa production of Bahia, Brazil, once a major world producer and exporter of chocolate, suffered a 95% crop loss since the introduction of C. perniciosa in 1989 (Pereira et al 1990, Isaac et al 1993).
In contrast, M. roreri infection is limited to the fruits of Theobroma and Herrania species, causing internal and external pod damage that results in total loss of the pods (FIGS. 1c, d
). The devastating effects and loss of cacao crops due to this pathogen have been dramatic. However, until the 1950s, frosty pod rot was confined to the northwestern part of South America (Colombia, Ecuador and western Venezuela) and therefore not of widespread interest to other cocoa-producing countries. However, its appearance in Panama in 1956 and Costa Rica in 1978 increased the geographic confines of the pathogen and changed perceptions of the disease (Evans 1986).
Evans et al (2003) consider that M. roreri is still in an invasive phase—having reached Nicaragua in 1980, Peru in 1988, Honduras in 1997, Guatemala in 2002, Belize in 2004, and Mexico in 2005 (Phillips-Mora 2003 and unpublished)—and is poised to devastate already crippled production in Bolivia and Brazil, once it arrives in those countries. Because this fungus has never been observed to form any type of fruiting body and the only propagules produced were considered to be conidia, the causal agent of frosty pod rot was originally described as an anamorphic ascomycete, Monilia roreri Cif. (Ciferri and Parodi 1933). Not until 1978 did a critical re-evaluation of M. roreri reveal the presence of dolipore septa (Evans et al 1978), a feature found only in basidiomycete hyphae. A new genus of anamorphic basidiomycete, Moniliophthora H.C. Evans, Stalpers, Samson & Benny, was erected to accommodate this taxon, whose placement within the basidiomycetes was unknown (Evans et al 1978). Recently, several studies have hypothesized that M. roreri might belong to the Tricholomataceae sensu lato (s.l.) (Phillips-Mora 2003, Phillips-Mora et al 2003), and might be closely related to C. perniciosa (Evans 1981, Evans et al 2002, Phillips-Mora 2003, Griffith et al 2003).
The purpose of the present study was to determine the phylogenetic placement and relatives of M. roreri, and to evaluate the hypothesis that M. roreri and C. perniciosa are closely related. This was determined by phylogenetic analyses of DNA sequences from five different nuclear gene regions: large subunit ribosomal DNA (LSU); small subunit ribosomal DNA (SSU); internal transcribed spacer regions 1 & 2 and the intervening 5.8S ribosomal subunit (ITS); elongation factor 1- (EF1-); and the largest subunit of RNA polymerase II (RPB1). As a result of the molecular study, a new combination, Moniliophthora perniciosa Aime & Phillips-Mora comb. nov. is proposed for the causal agent of witches’ broom, and a complete taxonomic discussion of the cacao pathogens and related agarics is provided.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONTAXONOMYLITERATURE CITED |
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). Field collections were described in the fresh condition and then preserved by drying in silica gel in individual containers; prior to drying, a piece (approximately 1–4 mm3) of tissue was preserved in 500 µL of 2x CTAB (hexadecyltrimethylammonium bromide) extraction buffer (Hillis et al 1996) for later DNA extraction. Vouchers of all field collections used for DNA sequences have been deposited in the U.S. National Fungus Collections (BPI) in Beltsville, Maryland.
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Polymerase chain reactions (PCR) and cycle sequencing.— –
Unless otherwise noted, all PCRs were performed in 25 µL reaction volumes with 12.5 µL of PCR Master Mix (Promega Corp., Madison, Wisconsin), 1.25 µL each of 10 µM primers (upstream and downstream), and 10 µL of diluted (10- to 100-fold) DNA template. PCR products were cleaned by one of two methods: (i) the majority were cleaned with Montage PCR Centrifugal Filter Devices (Millipore Corp., Billerica, Massachusetts) according to the manufacturer’s protocol; (ii) if more than one PCR product was produced during amplification, then the band of the correct size was excised from a 1% agarose gel and cleaned with the MinElute PCR Gel Extraction Kit (Qiagen, Inc., Valencia, California). Cleaned PCR products were sequenced with BigDye Terminator sequencing enzyme v.3.1 (Applied Biosystems, Foster City, California) in the reaction: 2 µL of diluted BigDye in a 1 : 3 dilution of BigDye:dilution buffer (400 mM Tris pH8.0, 10 mM MgCl2); 0.3 µL of 10 µM primer; 10–20 ng of cleaned PCR template; and H2O to 5 µL total reaction volume. Cycle sequencing parameters consisted of a 2 min denaturation step at 94 C, then 35 cycles of 94 C for 39 s, 50 C for 15 s and 60 C for 4 min. Sequencing reactions were cleaned by ethanol precipitation and sequenced on an ABI 3100 Genetic Analyzer (Applied Biosystems, Foster City, California). All DNA sequences have been deposited in GenBank, Accessions AY 916668–AY 916757 (TABLE I). Specific primers and PCR amplification parameters for each gene region are provided below.
The first 1250 bp of the large ribosomal subunit (LSU) were amplified with primers LSU4-B (5'-CTGGACCGTGTACAAGTCTCCTG, a basidiomycete-specific primer designed as the reverse-complement of the Gardes & Bruns [1993] primer ITS4-B) and LR6 and sequenced with LSU4-B, LR6, LR3 and LR3R (primer sequences for LR6, LR3 and LR3R are available on-line from the mycology lab of Duke University, http://www.biology.duke.edu/fungi/mycolab/primers.htm). Amplification was achieved with an initial denaturation step of 5 min at 94 C; 35 cycles of 30 s at 94 C, 45 s at 50 C, and 1 min at 72 C; and a final extension of 7 min at 72 C.
The complete small ribosomal subunit (SSU) of approximately 1800 bp was amplified with primers NS1 and NS8 and sequenced with NS1, NS2, NS3, NS4, NS5, NS6, NS7 and NS8 (http://www.biology.duke.edu/fungi/mycolab/primers.htm). Amplification and sequencing of the SSU from older herbarium specimens was achieved by two separate PCR reactions: (i) NS1 to NS4, sequenced with NS1, NS2, NS3 and NS4 and (ii) NS3 to NS8 sequenced with NS5, NS6, NS7 and NS8. In all cases, SSU amplification was carried out under the same cycling program as for the LSU, except the primer annealing step was carried out at 55 C, and final extension was at 72 C for 10 min.
Primers ITS1-F and ITS4-B (Gardes and Bruns 1993) were used to amplify and sequence about 800 bp that make up the internal transcribed spacer (ITS) region consisting of ITS-1, 5.8S ribosomal DNA, and ITS-2. Amplification parameters were the same as for the LSU, except the cyclic extension step was shortened to 45 s.
To sequence the elongation factor 1- region (EF1-), initially two degenerate primer pairs, (i) EF1-526F and 1567R and (ii) 983F and 2218R (Rehner and Buckley 2005), were used to amplify overlapping regions of the gene from a select group of exemplar taxa from the Marasmiaceae (Marasmius sp., Crinipellis sp. and Campanella sp.). The PCR products were ligated into pGEM-T Easy vectors and cloned in JM109 competent cells using the manufacturer’s protocols (Promega Corp., Madison, Wisconsin). Transformed colonies were amplified directly with the M13/pUC 17-mer forward and reverse primers (MBI Fermentas, Hanover, Maryland) in the following manner: individual transformed colonies were selected with a sterile toothpick that was then agitated in the PCR cocktail described above except that 10 µL of sterile H2O replaced the DNA template. The same cycling parameters as for the LSU were used, except 25 PCR amplification cycles were performed. PCR products of the correct size were cleaned and sequenced with the M13 primers. Sequences from exemplar taxa were aligned and several sets of primers were designed from nucleotide regions conserved across the exemplar taxa. After testing of primer pairs and PCR optimization, all additional taxa were amplified with primers RAS.EF1-F2 (5'-AGGARGCTGCTGAGYTSG) and RAS.EF1-R2 (5'-GCARGMATCRCCVGACTTGACR), with the PCR cocktail initially described except primer concentration was 5 µM instead of 10 µM, and with these cycling parameters: initial denaturation for 5 min at 94 C; 40 cycles of 94 C for 30 s, 57 C for 1 min, and 72 C for 2 min; 72 C final extension for 7 min. The approximately 1200 bp products were sequenced with the amplification primers and two internal primers, 983F (Rehner and Buckley 2005) and EFgr (Steve Rehner pers comm).
Amplification and sequencing of the largest subunit of RNA polymerase II gene (RPB1) were achieved in a similar manner as for the EF1-. Initially amplification and sequencing of exemplar taxa was done with primers RPB1-Af and RPB1-Cr (Matheny et al 2002), and internal sequencing primers RPB1-INT2F or RPB1-INT2.1F (P. Brandon Matheny, Clark University, Worcester, Massachusetts, pers comm). Optimized primers were designed and tested for enhanced specificity to marasmioid taxa, yielding a product of about 900 bp. All additional taxa were amplified and sequenced with primers RAS.RPB1-F2 (5'-CACCCCCACMACCCAATTTTCTGGGGG) and RAS.RPB1-R2 (5'-TCRTCYTCACTKCGCATYGCKCCWCCATCR), with one additional internal sequencing primer, RPB1-Br (Matheny et al 2002). Primer concentration in the PCR cocktail was 5 µM instead of 10 µM, and PCR cycling occurred as an initial denaturation step of 5 min at 94 C, followed by 40 cycles of 30 s at 94 C, 1 min at 55 C and 2 min at 72 C and a final extension of 7 min at 72 C.
Sampling strategy and sequence analyses.— –
Preliminary phylogenetic analysis was conducted by analyzing LSU sequences of Crinipellis perniciosa and Moniliophthora roreri within the 877 taxa dataset of Moncalvo et al (2002). This dataset contains representatives of all major euagaric lineages, as well as exemplar taxa from the other major homobasidiomycete clades using a heterobasidiomycete as outgroup (Moncalvo et al 2002). Bootstrapping analyses using maximum parsimony were conducted in PAUP* 4.0b10 (Swofford 2002) as described in Moncalvo et al (2002). Because these initial analyses could not resolve the generic placement of C. perniciosa and M. roreri within /marasmioid, additional taxon sampling focused on obtaining material and LSU sequences from taxa suspected to be members of the /marasmioid and /tetrapyrgoid clades, including the type species and other representatives from all genera proposed to be members of /marasmioid as inferred from the classification of Singer (1976, 1986). Material from South America was preferentially sampled because both pathogens are found only on that continent. A total of 45 (39 /marasmioid, and 6 /tetrapyrgoid as outgroup) LSU sequences were assembled and analyzed (Datamatrix A). Because phylogenetic placement of M. roreri and C. perniciosa could not be determined with confidence by the LSU dataset alone, Datamatrix B was constructed. Datamatrix B consisted of sequences of five gene regions (LSU, SSU, ITS, RPB1 and EF1-) for each of 15 taxa; these were selected as a cross sample representing each major clade uncovered with Datamatrix A.
Sequencing reactions were edited and contiguous sequences were assembled in Sequencher 4.1.4 (Gene Codes Corp., Ann Arbor, Michigan). Sequence alignments were constructed by eye in Se-Al v2.0a11 (Andrew Rambaut, Dept. Zoology, University of Oxford, UK; http://evolve.zoo.ox.ac.uk/). Sequences for Datamatrix A were trimmed to 1041 bp each, alignable across all bases. Datamatrix B consisted of a total of 5171 bp: 1769 bp of the SSU, alignable across all bases; 1041 bp from the LSU, alignable across all bases; ITS sequences were trimmed to 503 bp, alignable across all bases; RPB1 sequences were trimmed to 765 bp, alignable across all bases; and the EF1- was trimmed to 1093 bp, of which 120 bp that were too variable to confidently align were excluded from the final analysis.
Maximum parsimony (MP) analyses were conducted in PAUP* v4.0b10 as heuristic searches with 100 random addition replicates and TBR branch swapping. Support for the branching topologies was evaluated by bootstrap analysis derived from 1000 replicates with 10 random addition replicates each. Maximum likelihood (ML) analyses were conducted by the quartet puzzling method (Strimmer and von Haeseler 1996) in PAUP* with 10 000 puzzling steps; transition/transversion ratio = 2. For both datamatrices, the /tetrapyrgoid fungi were selected as outgroups for rooting purposes.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONTAXONOMYLITERATURE CITED |
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, placed M. roreri and C. perniciosa within the /marasmiaceae sensu Moncalvo et al (2002) (results not shown, but available from the lead author upon request); within the /marasmiaceae these taxa are strongly supported (67% booststrap support) members of the /marasmioid with /tetrapyrgoid as the sister clade. In these analyses, M. roreri and C. perniciosa appear as sister taxa, which together are sister to the genus Chaetocalathus, and are more distantly related to the genera Crinipellis and Marasmius.
Combined analyses of Datamatrix A, based on sequence data from the nLSU rDNA region alone, are shown (FIG. 2). Of 1041 included characters, 137 were parsimony-informative; 35 variable characters were parsimony-uninformative. Twenty-eight equally parsimonious trees of length 405 were found by MP; CI = 0.56; RI = 0.80. The same major clades were uncovered by both MP and ML, i.e. six monophyletic groups consisting of: (i) Campanella spp. and (ii) Tetrapyrgos spp. as outgroups; and (iii) Crinipellis spp. excepting C. perniciosa, hereinafter referred to as Crinipellis sensu stricto (s.s.); (iv) Chaetocalathus spp.; (v) Marasmius spp. (including Amyloflagellula inflata); and (vi) a group consisting of M. roreri, C. perniciosa, and an unnamed grass endophyte from New Mexico, hereinafter referred to as the Moniliophthora clade (FIG. 2). However, overall support for several of these groupings was low: only the genera Marasmius, Crinipellis s.s. and the two outgroup genera (Campanella and Tetrapyrgos) receive modest support with both methods; the Moniliophthora clade is not supported by bootstrapping methods, and weakly supported (45%) by quartet puzzling (FIG. 2).
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indicates exemplar taxa selected for additional sequencing to construct Datamatrix B. Results of analyses with Datamatrix B are presented (FIG. 3). Of 5049 included characters, 630 were parsimony-informative and 318 variable characters were not parsimony-informative. A single MP tree was found of length = 2044, CI = 0.62, and RI = 0.54. Both MP and ML uncovered the same clades in the analyses of Datamatrix B as had been uncovered with Datamatrix A. Significant support for each of the four ingroup taxa (Crinipellis s.s., Marasmius, Chaetocalathus, and the Moniliophthora clade) was found by both bootstrapping and quartet puzzling with Data-matrix B (FIG. 3).
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DISCUSSION |
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, 1986). Genera allied in the Crinipellinae are Crinipellis, Chaetocalathus, Amyloflagellula Singer, Lachnella Fr., and Flagelloscypha Donk apud Singer (Singer 1986). The majority of these species are saprotrophic litter fungi, and as a group considered to be sister to Marasmius and eight other reduced marasmioid genera (Singer 1986). Prior work has shown that Lachnella (Moncalvo et al 2002) and Flagelloscypha (Bodensteiner et al 2004) do not share a common ancestry with the core Crinipellinae. Our results (FIG. 2) and those of Bodensteiner et al (2004) have shown that at least some elements of Amyloflagellula are congeneric with Marasmius s.l. in /marasmioid. Furthermore, Moncalvo et al (2002) showed strong support for a monophyletic group consisting of Marasmius, Chaetocalathus and Crinipellis, termed /marasmioid, that is sister to Tetrapyrgos E. Horak and Campanella Henn. within a larger /marasmiaceae. Presently the Marasmiaceae Kühner is being redefined, and will incorporate those fungi in /marasmioid, i.e. Marasmius, Crinipellis, and Chaetocalathus (Dennis Desjardin, San Francisco State University and Andy Wilson, Clark University, pers comm). The present work shows that the two cacao pathogens also belong to the Marasmiaceae s.s. (FIGS. 2 and 3).
Moniliophthora was described as a monotypic, anamorphic genus of basidiomycetes, incertae sedis, although similarities in disease symptomatology between M. roreri and C. perniciosa had been noted (Evans et al 1978). Citing these symptomatological similarities and other similarities in cytology, Evans et al (2002) transferred M. roreri to Crinipellis, although no other species of the genus Crinipellis were examined. The present study shows that Moniliophthora is a member of the Marasmiaceae s.s., and that M. roreri and C. perniciosa are closely related fungi. Within the confines of our sampling, M. roreri and C. perniciosa appear as sister species (FIG. 3). Neither, however, is congeneric with Crinipellis as exemplified by the type species, C. stipitaria (Fr.) Pat., (Singer 1942, 1986; Kühner 1980) (FIG. 2). Rather, the two cacao pathogens form part of a distinct lineage within the Marasmiaceae s.s. that is not congeneric with any other genus previously allied within Subtribus Crinipellinae.
We recognize that changing the names of well-known pathogens is often accepted with reluctance. Singer noted in 1976 that it took more than 30 years for the preferential use of the name C. perniciosa over its prior name of Marasmius perniciosus; the name Monilia roreri still is often used for the causal agent of frosty pod rot even though 25 years have passed since the transfer of that taxon into Moniliophthora (Evans et al 2002). Nevertheless the present study makes necessary a taxonomic re-evaluation of the cacao pathogens.
At present two options are available for naming and identifying the monophyletic clade that contains M. roreri and C. perniciosa: (i) A new generic name is erected; or (ii) the name Moniliophthora, although typified by an anamorphic fungus, is used in essence as a pleomorphic name, and C. perniciosa is transferred into that genus. We have chosen the second option for several reasons. First, we advocate that nomenclature should reflect best phylogenetic estimates, and furthermore, that the proliferation of names in the literature should be avoided where at all possible. If a new generic name is erected to accommodate the agent of witches’ broom, then the current International Code of Botanical Nomenclature (ICBN) (Greuter et al 1999) does not allow the simultaneous transfer of M. roreri into that genus (Paul Kirk, CABI Bioscience, UK, pers comm), resulting in a situation where two sister-species do not share the same generic name. Although it is almost never done, to the best of our understanding nothing within the ICBN prohibits the transfer of a teleomorphic fungus into a genus based on an anamorph. Secondly, Evans et al (2002, 2003) provide cytological evidence that M. roreri is, in fact, a teleomorphic fungus that undergoes meiosis within its "conidia." Therefore the name Moniliophthora could be considered the earliest legitimate name for this genus.
The genus Crinipellis is delimited as those centrally-stipitate agarics of marasmioid stature that posses dextrinoid "hairs" on the pileipellis (Singer 1942). Approximately 75 species are currently recognized in the genus (Kirk et al 2001), the majority of which are saprotrophic litter and wood decomposers (Holliday 1980, Singer 1986). Eleven of these, including C. perniciosa and C. eggersii Pat., were placed in Section Iopodinae Singer (Singer 1976, 1986). We have molecular evidence that C. eggersii also belongs in Moniliophthora (unpublished), and it is likely that, pending molecular and type studies, the entirety of Crinipellis Section Iopodinae will be found to be congeneric with Moniliophthora. Section Iopodinae contains the other parasitic members of Crinipellis; other taxa in this section infect tropical rainforest and greenhouse trees such as species of Siparuna Aubl. and Vitex L (Singer 1976). Additionally, mushrooms in Section Iopodinae share the features of purple, violet or red pigments in the pileus that do not change color in the presence of an alkaloid solution, whereas members of the other sections of Crinipellis are brown pigmented or if not, the pigments turn green in the presence of KOH (Singer 1976, 1986). A few plant pathogenic species currently placed in the genus Marasmiellus Murrill—a genus now known to be synonymous with Gymnopus (Pers.) Roussel (Mata et al 2004)—have setose hairs on the pileus and may also belong to Moniliophthora.
It is tempting to speculate on the origins of Moniliophthora, especially given the discovery that an unnamed asymptomatic and presumably symbiotic endophyte of grasses, isolated from New Mexico, is congeneric with the two pathogens (FIG. 3), and it might be that the pathogens have evolved from a common biotrophic ancestor somewhere near the Andes of South America as commonly conjectured (see Evans et al 2003). Certainly, other potentially endophytic, biotrophic species of Moniliophthora await discovery.
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TAXONOMY |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONTAXONOMYLITERATURE CITED |
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FIGS . 1a, b
Marasmius perniciosus Stahel, Dep. Landb. Suriname 33:16. 1915.
Crinipellis perniciosa (Stahel) Singer, Lilloa 8:503. 1943 [1942].
This taxon is completely described in Singer (1942, 1976) and Holliday (1970) and discussed in Evans and Barreto (1996).
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Mention of trade names or commercial products in this article is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. 
1 Corresponding author. E-mail: cathie{at}nt.ars-grin.gov
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LITERATURE CITED |
|---|
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Chapela IH, Rehner SA, Schultz TR, Mueller UG. 1994. Evolutionary history of the symbiosis between fungus-growing ants and their fungi. Science 266:1691–1694.
Ciferri R, Parodi E. 1933. Descrizione del fungo che causa la "Moniliasi" del cacao. Phytopathologische Zeitschrift 5:539–542.
Evans HC. 1981. Pod rot of cacao caused by Moniliophthora (Monilia) roreri. Phytopath Pap 24:1–44.
———. 1986. A reassessment of Moniliophthora (Monilia) pod rot of cocoa. Cocoa Growers’ Bulletin 37:34–43.
———, Barreto RW. 1996. Crinipellis perniciosa: a much investigated but little understood fungus. Mycologist 10:58–61.
———, Holmes KA, Phillips W, Wilkinson MJ. 2002. What’s in a name: Crinipellis, the final resting place for the frosty pod rot pathogen of cocoa? Mycologist 16:148–152.[CrossRef]
———, ———, Reid AP. 2003. Phylogeny of the frosty pod rot pathogen of cocoa. Plant Path 52:476–485.[CrossRef]
———, Stalpers JA, Samson RA, Benny GL. 1978. On the taxonomy of Monilia roreri, an important pathogen of Theobroma cacao in South America. Can J Bot 56:2528–2532.[CrossRef]
Gardes M, Bruns TD. 1993. ITS primers with enhanced specificity for basidiomycetes—application to the identification of mycorrhizae and rusts. Molec Ecol 2:113–118.[Medline]
Greuter W, McNeill J, Barrie FR, Burdet HM, Demoulin V, Filgueiras TS, Nicolson DH, Silva PC, Skog JE, Trehane P, Turland NJ, Hawksworth DL, eds. 1999. International Code of Botanical Nomenclature (St Louis Code). Regnum Vegetabile 138. Königstein, Germany: Koeltz Scientific Books.
Griffith GW, Nicholson J, Nenninger A, Birch RN, Hedger JN. 2003. Witches’ brooms and frosty pods: two major pathogens of cacao. New Zealand J Bot 41:423–435.
Hillis DM, Moritz C, Mable BK. 1996. Molecular Systematics. 2nd ed. Sunderland, Massachusetts: Sinauer Associates. 655 p.
Holliday P. 1970. Crinipellis perniciosa. CMI Descriptions of Pathogenic Fungi and Bacteria No. 223.
———. 1980. Fungus Diseases of Tropical Crops. New York: Dover Publications, Inc. 607 p.
Isaac S, Hardwick K, Collin H. 1993. Interactions between the pathogen Crinipellis perniciosa and cocoa tissue. In: Isaac S, Frankland JC, Watling R, and Whalley AJS, eds. Aspects of tropical mycology. Cambridge, UK: Cambridge University Press. p 219–232.
Kirk PM, Cannon PF, David JC, Stalpers JA. 2001. Dictionary of the fungi. Oxon, UK: CAB International. 655 p.
Kühner R. 1980. Les Hyménomycètes agaricoïdes (Agaricales, Tricholomatales, Pluteales, Russulales) Etude gènèrale et classification. Bulletin de la Société Linnéenne de Lyon, France. 1027 p.
Mata JL, Hughes KW, Petersen RH. 2004. Phylogenetic placement of Marasmiellus juniperinus. Mycoscience 45:214–221.
Matheny PB, Liu YJ, Ammirati JF, Hall BD. 2002. Using RPB1 sequences to improve phylogenetic inference among mushrooms (Inocybe, Agaricales). Amer J Bot 89:688–698.
Moncalvo JM, Lutzoni FM, Rehner SA, Johnson J, Vilgalys R. 2000. Phylogenetic relationships of agaric fungi based on nuclear large subunit ribosomal DNA sequences. Syst Biol 49:278–305.[CrossRef][Medline]
———, Vilgalys R, Redhead SA, Johnson JE, James TY, Aime MC, Hofstetter V, Verduin SJW, Larsson E, Baroni TJ, Thorn RG, Jacobsson S, Clémençon H, Miller OK. 2002. One hundred and seventeen clades of euagarics. Molec Phylo Evol 23:357–400.[CrossRef]
Pereira JL. 1999. Renewed advance of witches’ broom disease of cocoa: 100 years later. In: International Cocoa Research Conference, 12th, Salvador, Bahía, Brasil, 1996. Proceedings. Lagos, Nigeria: Cocoa Producers’ Alliance: 87–91.
———, Ram A, Figueiredo JM, Almeida LCC. 1990. The first occurrence of witches’ broom disease in the principal cocoa growing region of Brazil. Tropical Agriculture 67:188–189.
Phillips-Mora W. 2003. Origin, biogeography, genetic diversity and taxonomic affinities of the cacao fungus Moniliophthora roreri as determined using molecular, phytopathological and morpho-physiological evidence [Doctoral dissertation]. Reading, UK: Univ. Reading. 349 p.
———, Krauss U, Evans H, Wilkinson M. 2003. Genetic diversity of the cacao pathogen Moniliophthora roreri (Cif.) Evans et al. in Tropical America. In: Proceedings of the Fourteenth International Cocoa Research Conference. Accra, Ghana: Cocoa Producers’ Alliance.
Rehner SA, Buckley E. 2005. A Beauveria phylogeny inferred from nuclear ITS and EF1- sequences: evidence for cryptic diversification and links to Cordyceps teleomorphs. Mycologia 97:84–98.
Singer R. 1942. A monographic study of the genera "Crinipellis" and "Chaetocalathus." Lilloa 8:441–534.
———. 1976. Marasmieae (Basidiomycetes-Tricholomataceae). Flora Neotropica No. 17. New York: New York Botanical Garden. 347 p.
———. 1986. The Agaricales in Modern Taxonomy. Koeltz Scientific Books, Germany. 981 p.
Stahel G. 1915. Marasmius perniciosus nov. spec. Dept Landbouw in Suriname Bull 33:1–27.
Strimmer K, von Haeseler A. 1996. Quartet puzzling: a quartet maximum-likelihood method for reconstructing tree topologies. Mol Biol Evol 13:964–969.
Swofford DL. 2002. PAUP*. Phylogenetic analysis using parsimony (*and other methods), version 4. Sunderland, Massachusetts: Sinauer Associates.
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70 % quartet puzzling, 