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Ceratocystis pirilliformis, a new species from Eucalyptus nitens in Australia

     Department of Microbiology and Plant Pathology, Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria, Pretoria, South Africa, 0002

B. D. Wingfield

     Department of Genetics, Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria, Pretoria, South Africa, 0002

M. J. Dudzinski
K. M. Old

     CSIRO Forestry and Forest Products, Canberra, ACT, Australia 2604

M. J. Wingfield

     Department of Microbiology and Plant Pathology, Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria, Pretoria, South Africa, 0002

	ABSTRACT

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Several species of Ceratocystis have been recorded on Eucalyptus. These include C. fimbriata, C. eucalypti, C. moniliformis and C. moniliformopsis. Of these, only C. fimbriata is known as a pathogen; it recently has been found causing serious wilt diseases in Uganda, Congo and Brazil. This study was undertaken to collect Ceratocystis species, including C. eucalypti, from artificially induced wounds on Eucalyptus nitens near Canberra in southeastern Australia. Trees were wounded in October 2000, and wounds were examined approximately one month later. Ascomata characteristic of a Ceratocystis species were found covering the wounds, and this fungus also was isolated from the wood using carrot baiting. This species of Ceratocystis has hat-shaped ascospores similar to those of C. fimbriata, but it differs from C. fimbriata and all other species of Ceratocystis in that it possesses ascomata with a pyriform base. Comparison of DNA sequences from the ITS and 5.8S rRNA operon confirmed that the fungus from E. nitens in Australia is unique, and we describe it here as a new species, C. pirilliformis.

Key words: Ascomycetes, Ceratocystis fimbriata, ITS rRNA, phylogeny, systematics

	INTRODUCTION

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Species of Ceratocystis are well known for their association with insects and, in general, are vectored non-specifically by flies (Diptera) and beetles (Coleoptera: Nitidulidae) (Crone and Bachelder 1961, Moller and DeVay 1968b, Hinds 1972, Juzwik and French 1983). These fungi require the host plant to be wounded to cause infections (Kile 1993) because fresh wounds attract sap-feeding insects that carry the fungi to these substrates. Sticky droplets of ascospores, produced at the apices of ostioles, adhere to the bodies of the insects. The insects then frequent wound sites on other plants and disperse ascospores from diseased to healthy hosts (Moller and DeVay 1968b, Upadhyay 1981). This insect-fungus relationship is facilitated by the fact that most species of Ceratocystis produce aromatics attractive to insects (Kile 1993).

Kile et al (1996) conducted a wounding study on Eucalyptus in Australia, and this resulted in the discovery of a new Ceratocystis species, C. eucalypti Z. Q. Yuan & Kile. This species is native to Australia, where it is found on E. sieberi L. Johnson and E. globoidea Blakely in Victoria and on E. regnans F. Muell. in Tasmania. Ceratocystis eucalypti appears to be a non-pathogenic colonist of fresh wounds on Eucalyptus (Kile et al 1996). The only other member of the genus known to infect living Eucalyptus is C. fimbriata, which has been reported to cause rapid wilting of trees in the Democratic Republic of Congo, Brazil (Roux et al 2000) and Uganda (Roux et al 2001). Ceratocystis fimbriata has been reported from Australia, although there are only two records of this fungus on rotting Syngonium in New South Wales, Queensland and Victoria (Walker et al 1988, Vogelzang and Scott 1991).

In an attempt to re-isolate C. eucalypti and other Ceratocystis species on Eucalyptus in Australia, an artificial wounding trial was conducted on planted E. nitens near Canberra and E. globulus near Cann River. Ceratocystis eucalypti was not isolated, but a morphologically distinctive species of Ceratocystis was abundant on the wounds. The aim of this investigation was to characterise this new species and identify its closest relatives, based on a comparison of rRNA sequence data.

	MATERIALS AND METHODS

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Fungal isolates – Artificial wounds were made approximately 1.2 m above the ground on stems of 6-year-old E. nitens near Canberra, Australian Capital Territory (ACT), and on 8-year-old E. globulus near Cann River, Victoria, in southeastern Australia. A brick-cutting chisel was used to remove blocks of bark approximately 10 cm square from the stems and expose the cambium. Thereafter, a second wound approximately 2 cm deep was made at the center of the exposed cambium to expose the xylem. The wounds were made in early October 2000 on 10 trees at each site. Wounds were examined in early November 2000, and slices of wood from all trees were removed from the wound surfaces for laboratory examination.

Chips of wood, approximately 4 cm square and 5 mm thick, were incubated at 25 C in 9 cm diam Petri dishes containing moistened filter paper to induce the production of fungal structures. Duplicate isolations were made by wrapping pieces of wood tightly between two slices of surface-disinfected carrot (5 mm thick) (Moller and DeVay 1968a). Carrot baits were incubated at 25 C for 10 d. Ascomata characteristic of Ceratocystis were found covering the wood chips and the carrot baits collected from five trees at the Canberra site. This fungus was not found on samples from Cann River. Cultures were obtained by transferring masses of ascospores from the apices of ascomatal necks onto 2% malt-extract agar (MEA, Biolab) and incubating them at 25 C. All cultures are maintained in the culture collection (CMW) of the Forestry and Agricultural Biotechnology Institute (FABI), University of Pretoria, South Africa. Dried and living cultures of the holotype and paratypes are deposited in the Plant Pathology Herbarium of NSW Agriculture (DAR), Orange Agricultural Institute, Orange, New South Wales, Australia, and dried cultures are deposited with the National Collection of Fungi, Agricultural Research Council, Pretoria (PREM), South Africa.

Morphology – The growth rates of three isolates (CMW6579, CMW6670 and CMW7569) were determined on 2% MEA. Isolates were cultured 2 wk at 25 C before growth-rate studies. Mycelial plugs were removed from the edges of actively growing cultures with a 4 mm cork borer, and a single plug was placed at the center of 60 mm Petri dishes containing 2% MEA. Four plates for each isolate were incubated at 15, 20, 25 and 30 C, respectively. Colony diameter was assessed after 12 d of incubation by making two measurements at right angles to each other for each culture. This resulted in eight measurements for each isolate at each temperature. Averages were computed for all growth measurements.

Morphological characteristics were described from cultures grown on 2% MEA. Fungal structures were mounted in lactophenol containing cotton blue. Fifty measurements for each taxonomically relevant structure were made, and corresponding ranges, averages and standard deviations calculated. Color descriptions were determined using the color charts of Rayner (1970).

DNA extraction – Single drops containing ascospores from ascomata were transferred from sporulating cultures to 50 mL, 2% malt extract broth using sterile toothpicks. Cultures were incubated at 25 C for 2 wk to obtain thick mycelial mats. These were freeze-dried, crushed in liquid nitrogen and the DNA extracted according to the method described by Barnes et al (2001).

PCR amplification – The two internal transcribed spacer regions (ITS1 and ITS2) and the 5.8S gene of the ribosomal RNA operon were amplified using primers ITS1 and ITS4 (White et al 1990). Polymerase chain reaction (PCR) mixtures consisted of 200 nM of each primer, 200 µM of each dNTP, Expand High Fidelity PCR System enzyme mix (1.75 U) (Roche Molecular Biochemicals), 1x Expand HF buffer containing 1.5 mM MgCl₂ (supplied with the enzyme) and 2–10 ng DNA. Reaction volumes were adjusted to 50 µL with Sabax water. The PCR program was set at 96 C for 2 min, followed by 10 cycles at 94 C for 20 s, 55 C for 48 s and 72 C for 45 s. A further 25 cycles were included with the annealing time altered to 40 s and a 5 s extension after each cycle. A final step of 10 min at 72 C completed the program. PCR amplicons were purified using the Magic PCR Preps, Purification System (Promega).

Sequencing – PCR amplicons were sequenced with the ABI PRISM^TM Big DYE Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems), and the primers ITS1 and ITS4 were used. Sequence reactions were run on an ABI PRISM^TM 377 Autosequencer (Applied Biosystems) and sequence electropherograms were analyzed with Sequence Navigator version 1.0.1 (Applied BioSystems).

The sequences obtained for the Ceratocystis species from E. nitens were compared to ITS sequences of Ceratocystis species obtained from GenBank (Table I). Sequences were aligned manually and analyzed using PAUP version 4.0* (Phylogenetic Analysis Using Parsimony * and other methods) (Swofford 1998). Gaps were treated as "newstate", and trees were obtained via stepwise addition of 1000 replicates. The Mulpar option was in effect. The heuristic search, based on parsimony with tree bisection reconnection, was used to obtain the phylogram. Confidence intervals using 1000 bootstrap replicates were calculated. Outgroup C. moniliformis was treated as a paraphyletic sister group with respect to the ingroup. All sequences derived in this study have been deposited in GenBank (Table I) and the sequence alignments in TreeBASE (5869).

	RESULTS

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View larger version (25K): [in this window] [in a new window]   FIG. 1. Phylogram inferred from analysis of ITS 1, 5.8s and ITS 2 rDNA sequences (L = 471, CI = 0.745, RI = 0.848, g1 = -0.566). Ceratocystis moniliformis was used as outgroup. Bootstrap confidence values are indicated above the branches

	TAXONOMY

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Ceratocystis pirilliformis I. Barnes & M. J. Wingfield sp. nov. Figs. 2 and 3

View larger version (51K): [in this window] [in a new window]   FIG. 2. Cultural characteristics of Ceratocystis species most closely related to C. pirilliformis. A) C. albofundus from Acacia mearnsii (CMW2475) in South Africa produces cultures that are cream-colored (21''f). B) C. pirilliformis (CMW6579) has light gray aerial mycelium (21'''''d) with greenish to brownish submerged mycelium (21''''b). C) C. fimbriata from Eucalyptus grandis in Uruguay (CMW7383) produces cultures that are brownish (21''K). All cultures were grown on 2% MEA at 25 C for ca 3 wk

View larger version (94K): [in this window] [in a new window]   FIG. 3.  Ceratocystis pirilliformis (PREM 57323). a. Ascomata with a pyriform venter. b. Ostiolar hyphae with ascospores emerging through the opening of the neck. c. Hat-shaped ascospores in side and top view. d. Conidiophores with cylindrical conidia being released from the conidiogenous cells. e. Barrel-shaped conidia. f. Cylindrical conidia. g. Chlamydospore. All scale bars = 10 µm

Coloniae tarde crescentiae, usque ad 22 mm diametro post 12 dies; effusae, mycelio aerio laete olivaceo-griseo, mycelio immerso griseo-olivaceo, pagina reversa coloniae griseo-olivacea. Ascomata nigra, venter pyriformis, basin versus globosa, 115.2–186.8(–205.5) µm diametro, parte superiori late papillata, 35.8–76.5(–86.8) x (23.9–42.4(–75.2) µm. Colla ascomatarum erecta vel curvata, basin versus nigra, apicem versus subhylainescens, parietibus laevis vel crenulatis, basin versus 18.8–32.7(–39.8) µm lata, apicem versus 11.9–21.0(–25.3) µm lata; 372.4–683.0(–777.5) µm longa, hyphis ostiolaribus inclusis. Ascosporae in guttulis in collis ascomatarum crescentes, unicellulares, hyalinae, e supra visae ellipticae, 4.3–5.7(–6.4) x 3.2–4.4(4.8) µm, vagina hyalina aspectu laterali petasiformi circumcincta, 2.9–5.2(–6.5) x 2.0–3.8(–5.4) µm, margine 4.7–6.4(–7.5) µm longa, diffluentes. Conidiophorae chalariformes, erectae, ramosae vel non ramosae, hyalinae, multiseptatae, laeves, 62–147(–216) µm longae, cellula terminali conidiogena inclusa. Cellulae conidiogenae tubulares, determinatae, cylindricae vel lageniformes, 31.9–63.6(–72.4) µm longae. Conidia enteroblastica phialidosa, formis duabus: i) doliiformia, basi truncata, parietibus laevis, non septata, mononemata, hyalina, 4.1–5.6(–6.4) x 3.2–4.3(–4.8) µm, solitaria vel in catenis portata, vel ii) cylindrica, apicibus rotundatis, laevia, non septata, hyalina, 11.6–25.2(–33.0) x 2.4–4.0(–4.7) µm, in catenis portata.

Colonies slow growing with optimal growth at 25 C on 2% MEA, reaching 22 mm diam in 12 d. No growth below 15 C or above 30 C. Colonies effuse, aerial mycelium "pale olivaceous grey" (21'''''d), submerged mycelium "grey olivaceous" (21''''b). Reverse side of colony "grey olivaceous" (21''''b). Submerged mycelium darkening as the ascomata develop forming fine, radiating fibrils. Ascomata developing within 8 d and mature within 12 d, superficial or partly embedded in agar, black (7'''''k). Venter pyriform with basal part globose 115.2–186.8(–205.5) µm diam and upper part broadly papillate, 35.8–76.5(–86.8) x 23.9–42.4(–75.2) µm. Ascomatal necks erect, occasionally curved, black at bases becoming subhyaline toward the apex, smooth to crenulate, tapering slightly from 18.8–32.7(–39.8) µm wide at base to 11.9–21.0(–25.3) µm wide at apex, 372.4–683.0(–777.5) µm long including ostiolar hyphae. Ostiolar hyphae extending from the outer layer of the neck cells, hyaline, straight or flexuous, subulate, mostly convergent, non-septate. Asci evanescent, not seen. Ascospores accumulating in droplets at tips of ascomatal necks, single-celled, hyaline, elliptical in top view, 4.3–5.7(–6.4) x 3.2–4.4(–4.8) µm, surrounded by a hyaline sheath; sheath appearing hat-shaped in side view, 2.9–5.2(–6.5) x 2.0–3.8(–5.4) µm, dissolving at maturity. Conidiophores, thielaviopsis-like, erect, branched or unbranched, hyaline, multiseptate, smooth walled, 62–147(–216) µm long including the integrated, terminal conidiogenous cell. Conidiogenous cells tubular, determinate, cylindrical to lageniform, 31.9–63.6(–77.4) µm long. Conidia of two types emerging from different conidiogenous cells: i) barrel-shaped, doliiform with truncate bases, smooth-walled, non-septate, hyaline, 4.1–5.6(–6.4) x 3.2–4.3(–4.8) µm, borne in chains, ii) cylindrical with apices rounded, smooth, non-septate, hyaline, 11.6–25.2(–33.0) x 2.4–4(–4.7) µm, borne in chains. Chlamydospores oval, thick walled, smooth, isabelline (17''i) to olivaceous (21''m), 7.8–12(–13.0) x 5.4–8.3(–9.5) µm, embedded in agar, formed singly or in short chains.

HOLOTYPE: AUSTRALIA. ACT: Uriarra, near Canberra, isolated from wounds on Eucalyptus nitens, Nov 2000, M. J. Wingfield [PREM57323. ISOTYPE: DAR 75996 (culture CMW6579)].

PARATYPES: AUSTRALIA. ACT: Uriarra, near Canberra, isolated from wounds on Eucalyptus nitens, Nov 2000, M. J. Wingfield [PREM57322, DAR 75993 (culture CMW6569)]; same collecting data [PREM57320, DAR 75994 (culture CMW6574)]; same collecting data [PREM57325, DAR75997 (culture CMW6583)]; same collecting data [PREM57321 (culture CMW6566)]; same collecting data [PREM57324, DAR 75995 (culture CMW6577)].

Etymology. pirilliformis, Latin = shaped like a little pear; referring to the characteristic pear-shaped venter of the ascomata in this species.

	DISCUSSION

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Ceratocystis pirilliformis can be distinguished easily from all other species of Ceratocystis. Although, morphologically, it most resembles C. fimbriata and C. albofundus, it is distinctive in possessing ascomata with pyriform bases. Comparisons of ITS rRNA sequences support the treatment of C. pirilliformis as a new species. It is one of the five species of Ceratocystis known to occur on Eucalyptus and only the third isolated from living Eucalyptus tissue.

There is considerable overlap in the morphological characteristics of Ceratocystis species. The sizes of ascomata, ascospores and conidia of C. pirilliformis, for example, fall within the ranges observed for C. fimbriata and C. albofundus (Wingfield et al 1996). However, notable differences separate these three species. Ceratocystis albofundus has light-colored ascomatal bases, divergent ostiolar hyphae and lacks chlamydospores (Wingfield et al 1996). Ceratocystis pirilliformis and C. fimbriata are almost indistinguishable from each other. Both have dark ascomatal bases, convergent ostiolar hyphae and chlamydospores. The only obvious difference between these two species is the shape of the bases of their ascomata. Ceratocystis fimbriata has globose ascomatal bases, whereas the bases of C. pirilliformis are distinctly pyriform.

Respective culture morphologies of C. fimbriata, C. albofundus, and C. pirilliformis differ. Colonies of C. albofundus are pale, almost creamy, while C. fimbriata varies from greenish to brown. Ceratocystis pirilliformis forms colonies with a grayish aerial mycelium and a submerged mycelium that is green, sometimes turning brown in older cultures.

Ceratocystis pirilliformis and C. eucalypti both are found on Eucalyptus in southeastern Australia. Ceratocystis eucalypti probably is endemic to this area (Kile et al 1996), and we suspect that the same is true of C. pirilliformis. These species, however, are distinct morphologically. The most obvious difference is the shape of the ascospores; C. eucalypti has elongated, sheathed ascospores, whereas C. pirilliformis has hat-shaped, sheathed ascospores. Only five other species of Ceratocystis have hat-shaped ascospores, i.e., C. moniliformis (Hedgec.) C. Moreau, C. moniliformopsis Z.Q. Yuan & C. Mohammed, C. acericola Griffin, C. fimbriata and C. albofundus. Ceratocystis moniliformis and C. moniliformopsis, however, differ from other species with hat-shaped ascospores in that they have short conical spines on the ascomatal bases (Upadhyay 1981, Yuan and Mohammed 2002).

Phylogenetic comparisons, based on sequences of the ITS regions, strongly support the separation of species of Ceratocystis based on observed morphological differences. Species within Ceratocystis can be divided into two major groups. One of these includes C. coerulescens (Witthuhn et al 1999) and other species that do not have hat-shaped ascospores. Ceratocystis eucalypti is most closely allied to species in the C. coerulescens group, a relationship that confirms that ascospore shape is a reasonably strong indicator of phylogenetic relationships within Ceratocystis. The second major group in Ceratocystis is typified by C. fimbriata. It is not surprising, therefore, that C. pirilliformis, with a morphology similar to C. fimbriata, resides in a strongly resolved clade within this subgroup.

Eucalyptus nitens, also known as shining gum, occurs naturally in Australia, where it mainly is restricted to the southeast (Poynton 1979). Large artificial plantations have been established in Tasmania and, to a lesser extent, in New South Wales (Beadle 1999). The tree also is planted widely in South Africa for pulp production, and it increasingly is being used as a hybrid partner with other species to increase cold tolerance. Although we know little concerning the pathogenicity of C. pirilliformis, it certainly is able to cause wood discoloration. Given the economic importance of E. nitens and the fact that many species of Ceratocystis are pathogenic, it will be important to test this characteristic of C. pirilliformis.

	ACKNOWLEDGMENTS

 
We are grateful to members of the Tree Protection Co-operative Program (TPCP), National Research Foundation (NRF), the THRIP initiative of the Department of Trade and Industry, South Africa, and CSIRO Forestry and Forest Products for financial support. Ruth Gibbs ably assisted with fieldwork. We also thank Dr. Hugh Glen for preparing the Latin diagnosis and Prof. J.P. van der Walt for suggesting a name for the new species.

	FOOTNOTES

 
¹ Corresponding author, Email: irene.barnes{at}fabi.up.ac.za

Accepted for publication February 3, 2003.

	LITERATURE CITED

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS TAXONOMY DISCUSSION LITERATURE CITED

 
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Beadle C., 1999 Pruning a knotty problem for plantation timber. CSIRO Media Release 99/57

Crone LJ, Bachelder S., 1961 Insect transmission of canker stain fungus, Ceratocystis fimbriata f. platani. Phytopathology 51:576

Hinds TE., 1972 Ceratocystis canker of aspen. Phytopathology 62:213-220

Juzwik J, French DW., 1983 Ceratocystis fagacearum and C. piceae on the surfaces of free-flying and fungus-mat-inhabiting nitidulids. Phytopathology 73:1164-1168

Kile GA., 1993 Plant diseases caused by species of Ceratocystis sensu stricto and Chalara. In: Wingfield MJ, Seifert KA, Webber JF, eds. Ceratocystis and Ophiostoma: taxonomy, ecology and pathogenicity. St. Paul, Minnesota: American Phytopathological Society. p 173–183

Kile GA., Harrington TC, Yuan ZQ, Dudzinski MJ, Old KM., 1996 Ceratocystis eucalypti sp. nov., a vascular stain fungus from eucalypts in Australia. Mycol Res 100:571-579

Moller WJ, DeVay JE., 1968a Carrot as a species-selective isolation medium for Ceratocystis fimbriata. Phytopathology 58:123-126

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Poynton RJ., 1979 Tree planting in Southern Africa. The Pines. South African Forestry Research Institute 1:576

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White TJ, Bruns T, Lee S, Taylor J., 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 sequencing guide to methods and applications. San Diego: Academic Press. p 315–322

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This Article Abstract Full Text (PDF) Services Similar articles in this journal Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Barnes, I. Articles by Wingfield, M. J. Search for Related Content PubMed Articles by Barnes, I. Articles by Wingfield, M. J. Agricola Articles by Barnes, I. Articles by Wingfield, M. J.