RESUMEN
A survey to identify pathogens causing peony diseases in southern Chile (37°95' to 40°58'S) was conducted from 2008 to 2011. A noticeable symptom consisted of small red spots on leaves, stems, sepals, and first petals of the bud. The central part of the lesions (1 cm) enlarged, darkened, and became necrotic, coincident with rainy and cold weather, including frosts. Lesions grew but never coalesced completely. At the edges of leaves, the infected tissues cracked and produced a twisted and corrugated appearance. Rhizome symptoms were also observed and consisted of black spots, which enlarged to elongated lesions. Small pieces (~1 × 1 mm) of symptomatic leaves, previously washed with 2% NaOCl for 2 min were cultured on potato dextrose agar (PDA) and V8 juice agar (20%) media. The cultures were incubated in growth chambers at 20°C (dark) and 5°C (dark)/20°C (light) for 12/12 h (dark/light). Sporulation was obtained only with V8 under the dark and light regime. Mycelia growth was superficial and immersed in the media; hyphae were septate with thick black walls. Conidiophores were flexuous, not branched, and produced a single spore. Spores were elongate, multiseptate, with a long, strongly curved beak. After one week, sporulation decreased and thick-walled, round black clamydospores formed in the media. Spores ranged from 125 to 235 µm long and most frequently were 170 to 220 µm. The number of septa ranged from 5 to 12. Morphological and cultural characteristics fit the description of Mycocentrospora acerina (Harting) Deighton (1,2). DNA was obtained from fungal culture. The internal transcribed spacer (ITS) region was amplified using ITS1/ITS4 primers (3), and part of the amplicon (502 of 550 pb) was sequenced. The sequence was deposited in GeneBank (Accession No. KF015599) and showed 100% identity values with sequences of similar regions from M. acerina (strain ATCC 16259, Accession No. KF278454). Healthy leaves and rhizome pieces were washed with sterile distilled water and placed on sterile moist paper towel. A small agar disc with mycelium was placed on the leaves and rhizomes with and without wounding before inoculation. The inoculated materials were kept in closed boxes with high humidity at 5 to 20°C. Tests were positive in leaves and rhizomes with and without wounds. Seven days after leaf inoculations, necrotic symptoms developed. A second inoculation test in peony leaves of plants bagged with plastic under field conditions corroborated the in vitro test. Inoculated rhizomes developed dark orange lesions initially, then turned black with a watery consistency, similarly to affected rhizomes in the field. In both tests, controls showed no symptoms. To our knowledge, this is the first report of M. acerina on P. lactiflora globally (4). In an evaluation at Carillanca (38°41'S, 72°25'W), using a scale from 0 (no symptoms) to 7 (necrotic lesions covering >60% of foliage, with death of stems), 11 out of 31 peony varieties scored 0-1. As a reference, a score of 3 is the maximum damage allowed on peonies for export. These varieties might help to keep Chilean peonies in the market without an economic impact. References: (1) M. B. Ellis. Dematiaceous Hyphomycetes. CMI. Surrey, England, 1971. (2) B. C. Sutton and I. A. S Gibson. M. acerina. CMI. Descriptions of Pathogenic Fungi and Bacteria, No. 537.1977. (3) T. J. White et al. PCR Protocols. Academic Press, San Diego, CA, 1990. (4) D. F. Farr and A. Y. Rossman. Fungal Databases. Syst. Mycol. Microbiol. Lab., ARS, USDA. Retrieved April 16, 2014.
RESUMEN
Oilseed rape (Brassica napus L.) plants with typical club root symptoms were detected on two farms in the La Araucanía region (37°35' to 39°37' S), southern Chile. In 2010, affected plants were found in large areas throughout three fields on a single farm and disease incidence ranged from 30 to 100%. In 2013, plants with club root were found in one field on a different farm. Disease incidence in the affected areas was 30%. In both cases, affected plants showed root swellings or distortions, but no aerial symptoms were evident. Cross-sections from symptomatic roots were observed under light and fluorescent microscope and compared to healthy roots. The presence of plasmodia with resting spores in the root tissue pointed to Plasmodiophora brassicae Woronin as the causal agent. Pathogenicity was evaluated in the greenhouse. Inoculum was prepared by grinding 10 g of fresh galled roots in sterile water. The macerated tissue was filtered through sterile cheesecloth and the spore suspensions were adjusted to 1 × 107 spores/ml. Seeds of oilseed rape cv. Imminent were germinated and 5-day-old seedlings were transplanted in 250-ml pots (4 seedlings per pot). The soil surrounding the base of each seedling was inoculated with 1 ml of spore suspension. One pot received no inoculum and was used as a control. Pots were watered regularly. After 45 days, the inoculated plants showed root swelling similar to that observed in the field, whereas no symptoms were observed on the roots of the control plants. Specific PCR detection for P. brassicae was performed with pairs of primers TC1F/TC1R and TC2F/TC2R, according to the protocol described by Cao et al. (1). Total DNA was obtained from old galled roots collected in the field and galled roots from plants of the pathogenicity test, using the E.Z.N.A HP Plant DNA mini kit (Omega Biotek). Amplicon sizes of 548 and 519 bp, respectively, were obtained for each primer set, which is consistent with that reported by Cao et al. (1). Seed contamination with P. brassicae in the same seed lot used to sow the commercial field of 2013 was evaluated using the PCR method described above. Washing protocols to collect resting spores from seed was based on Rennie et al. (2). Total DNA was extracted from the resting spores pellet that had been ground in liquid nitrogen, using E.Z.N.A HP Plant DNA mini kit. PCR was performed on undiluted and diluted (1/10, 1/100, 1/1000, and 1/10000) DNA. Total DNA from a plant with gall roots where plasmodia were observed and a plant with healthy roots were used as positive and negative control, respectively. A 548-bp amplicon was amplified in the 1/10 and 1/100 dilutions with the TC1F/TC1R primers only indicating that the pathogen may have been present in the seed lot. In Chile, club root symptoms on B. napus were described in 2008 (3), though no indication of location or incidence was given. To our knowledge, this is the first confirmed case of club root disease in an oilseed rape field. This finding could prelude new cases and possibly an outbreak of club root disease on oilseed rape, jeopardizing this important crop of southern Chile. Oilseed rape production in Chile relies on imported seed of hybrids and parental materials. Although seed contamination with P. brassicae is thought to play a minor role in the epidemiology of the disease, we cannot ignore the possibility that the occurrence of this disease in 2013 may have been associated with the use of contaminated seed. References: (1) T. Cao et al. Plant Dis. 91:80, 2007. (2) D. C. Rennie et al. Plant Pathol. 60:811, 2011. (3) Rina Acuña P. Compendio de Fitopatógenos de Cultivos Agrícolas en Chile. Servicio Agrícola y Ganadero, Santiago, Chile, 2008.
RESUMEN
Branched broomrape is a holoparasitic weed present in some areas of central and southern Chile (33°S to 38°S), which is often found parasitizing tomato and tobacco crops. During an extensive survey conducted in different tomato-growing areas during the summer of 2010, branched broomrape plants with stem rot symptoms were detected in a commercial tomato crop located in the central zone (34°14'S, 71°1'W). Rotten stems were observed with white mycelia and approximately 1-mm-diameter spherical sclerotia on affected tissue below the soil surface. Parasitized tomato plants showed no symptoms. Sclerotia were taken directly from affected stems with a dissecting needle under a stereoscopic microscope in a flow chamber and placed on potato dextrose agar (PDA) medium. Germinating sclerotia consistently produced colonies similar to Sclerotium rolfsii with new sclerotia formed within 6 to 7 days. Mycelia produced hyphal clamp connections under the light microscope (2). DNA was extracted from one fungal culture. The ITS1 region, 5.8S rRNA gene, and the ITS2 region of the nuclear-encoded ribosomal RNA gene were amplified with primers ITS1 and ITS4 (4). The sequence was deposited in GenBank (Accession No. HM222638) and showed ≥99% identity values with sequences of similar regions from Athelia rolfsii (anamorph S. rolfsii; Accession Nos. AB075304, DQ0595578, AF499018, and AB075305). Different pathogenicity tests were performed. Inoculum was prepared by placing mycelia plugs from a PDA-grown, 6-day-old colony in a flask with sterilized wheat seeds and incubated for 2 weeks. Three Orobanche plants, each one with 10 to 15 shoots at different aerial stages (starting emergence, flowering, and formed capsules) were gently planted separately in 35-cm pots. Inoculum (10 g per pot) was placed in the soil surrounding the plants. One pot was used as a control. Forty-five-day-old tomato and tobacco plants were additionally inoculated by a similar procedure. After 12 days, inoculated Orobanche plants showed reduced vigor and stem decay. After 9 days, tomato and tobacco plants showed wilt. In all cases, the fungus was reisolated on PDA from all inoculated plants. To our knowledge, this is the first report of S. rolfsii on O. ramosa (1). A previous report of S. rolfsii parasitizing O. cernua has also been made (3). The high susceptibility of tomato and tobacco plants to this isolate of S. rolfsii precludes the use of this pathogen as a biological control agent against broomrape. References: (1) D. F. Farr and A. Y. Rossman. Fungal Databases. Systematic Mycology and Microbiology Laboratory, ARS, USDA. Retrieved May 14, 2010, from http://nt.ars-grin.gov/fungaldatabases . (2) Z. K. Punja and A. Damiani. Mycologia 88:694, 1996. (3) C. A. Raju et al. Phytoparasitica 23:235, 1995. (4) T. J. White et al. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. Page 315 in: PCR Protocols: A Guide to Methods and Applications. M. A. Innis et al., eds. Academic Press, New York, 1990.
RESUMEN
Chickpea (Cicer arietinum L.) plants with foliar and stem lesions fitting the description of Ascochyta blight were observed in October 2002 in four chickpea crops located in the La Araucania Region (38°S, 72°24'W) in southern Chile. Large, circular foliar and stem lesions containing pycnidia arranged in concentric circles were observed (1). Stem breakage also was observed. Isolates were obtained from mature pycnidia developed on stems by culturing a spore suspension on potato dextrose agar (PDA) and chickpea seed meal agar. A pathogenicity test was performed by inoculating 25 plants with a suspension of 1.2 × 105 conidia ml-1 and incubating at 22°C and 75% relative humidity. Foliar and stem lesions were observed 5 and 7 days after inoculation, respectively. Four check plants sprayed with sterile distilled water showed no symptoms. Fungal colonies obtained from inoculated plants showed the same cultural characteristics as the original isolates. Cultural morphology was consistent with the description of Ascochyta rabiei (Pass.) Labrousse (teleomorph Didymella rabiei (Kovacheski) v. Arx (= Mycosphaerella rabiei Kovacheski)) (3). Conidia produced on PDA were predominantly aseptate, 3.90 to 5.85 µm wide, and 9.75 to 11.7 µm long. Affected plants (cv. Kaniva) originated from seed introduced at commercial volumes (69 ton) from Victoria, Australia in August 2002. A. Rabiei can be disseminated via infected seed (1). Ascochyta blight symptoms also have been observed in small patches in several crops near Temuco, the capital of the La Araucania Region. Chickpea production is currently, relatively small in southern Chile, however, plans to promote its cultivation may be hindered by this outbreak. Previously, the only other country to report Ascochyta blight of chickpea in South America was Bolivia (2). References: (1) W. J. Kaiser. Epidemiology of Ascochyta rabiei. Pages 117-134 in: Disease-resistance Breeding in Chickpea. K. B. Singh and M. C. Saxena, eds. ICARDA, Aleppo, Syria, 1992. (2) W. J. Kaiser et al. Plant Dis. 84:102, 2000. (3) E. Punithalingam and P. Holliday. No. 337 in: CMI Descriptions of Pathogenic Fungi and Bacteria. CMI, Kew, Surrey, UK, 1972.