Survey for Botrytis species associated with onion bulb ...

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Aug 6, 1999 - Australasian Plant Pathology Society 2004. 10.1071/AP04046. 0815-3191/04/030419 www.publish.csiro.au/journals/app. Australasian Plant ...
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Australasian Plant Pathology, 2004, 33, 419–422

Survey for Botrytis species associated with onion bulb rot in northern Tasmania, Australia Martin I. ChilversA, F. S. HayA,C and C. R. WilsonB A

Tasmanian Institute of Agricultural Research, University of Tasmania North West Centre, PO Box 3523, Burnie, Tas. 7320, Australia. B Tasmanian Institute of Agricultural Research, New Town Research Laboratories, 13 St Johns Avenue, New Town, Tas. 7008, Australia. C Corresponding author; email: [email protected]

Abstract. Botrytis allii (including B. aclada and B. allii sensu stricto) was detected in latent form in leaf samples in 6 of 16 onion crops sampled early to mid-season in Tasmania, Australia during 1999/2000. Botrytis neck rot was detected in bulb samples from eight of eight crops sampled after harvest at an incidence of 0.4–16.3%, with a mean of 5.3%, and was correlated (r = 0.83, P < 0.05) with leaf infection. AP0406 MeB.taoIrl.Chytis lvpersciesandonino ublbort

Introduction Neck rot of onion (Allium cepa), caused by a complex of Botrytis spp., is an important fungal disease of onion worldwide (Lacy and Lorbeer 1995). In Tasmania, Australia, Botrytis neck rot has caused considerable losses to the onion industry in some seasons (Dennis 1996). However, good control is currently achieved by rotation, using seed tested for the fungus, fungicidal seed treatment and application of benzimidazole fungicides during the season. Recent studies (Nielsen et al. 2001, 2002; Nielsen and Yohalem 2001) of the ribosomal internal transcribed spacer (ITS) region of the genome of Botrytis spp. associated with neck rot of onion have confirmed the existence of three distinct groups. These include a smaller-spored group with 16 mitotic chromosomes, (B. aclada AI), a larger-spored group with 16 mitotic chromosomes (B. byssoidea) and a group with intermediate-sized spores with 32 mitotic chromosomes (B. aclada AII). Yohalem et al. (2003) proposed that B. aclada AI and B aclada AII be referred to as B. aclada and B. allii, respectively. Our study was conducted prior to the work of Yohalem et al. (2003) and did not differentiate B. aclada from B. allii. A limited study of 23 isolates from Australia identified one isolate as B. aclada and 22 isolates as B. allii, with no B. byssoidea detected (Chilvers et al. 2004). As B. allii was the most common, the fungus is referred to by this name in this study. B. allii infects the onion plant in the field through transmission from seed to cotyledons or from external sources of airborne conidia infecting leaves during the season. Maude and Presly (1977a) reported B. allii was © Australasian Plant Pathology Society 2004

transmitted from seed to seedling by the fungus invading the tip of the cotyledon leaf from the attached seed coat and growing downwards into the cotyledon leaf, only producing conidiophores when the leaf tissue became senescent. Occasionally, the fungus moved systemically from the moribund cotyledon up into the first true leaves. However, more often, conidia were produced on the cotyledon, leading to subsequent infections from the tops of the true leaves down (Maude and Presly 1977a). Stewart and Franicevic (1994) also found that B. allii infected the cotyledon before it moved down the plant to infect the true leaves. B. allii usually remains symptomless in leaves and grows from the leaves into the bulb during curing, leading to rots of bulbs in storage. However, conidiophores and sclerotia, and neck and basal rots on bulbs are occasionally noted in the field. The purpose of this study was to characterise the prevalence and incidence of B. allii in foliage of Tasmanian onion crops in the field and to determine the incidence of neck rot of bulbs following storage. Methods Survey of crops for B. allii Early- and late-season onion crops of cv. ‘Cream Gold’ at a range of locations in northern Tasmania were surveyed between late October and late December 1999 (Table 1). Samples were collected from a corner of each field comprising an area 100 m long by 50 beds of onions wide (approximately 80 m). Five leaf samples were collected from each bed, with each bed crossed four times along a ‘W’ pattern. The oldest (most basal) green leaf on each plant was collected, as it would potentially have been exposed to infections for a longer period than younger leaves. Alderman and Lacy (1984) reported that older leaves were more likely 10.1071/AP04046

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Table 1. Crop

A B C CA D DA E F G GA H J K L M N O P Q

M. I. Chilvers et al.

Incidence of Botrytis allii in leaf and bulb samples from commercial onion crops in Tasmania during the 1999/2000 season Location – nearest town

Sowing date

Sample date

Burnie Kindred Forth Forth Forth Forth Forth Forth Forth Forth Kindred Forth Forth Penguin Ulverstone Ulverstone Scottsdale Scottsdale Latrobe

6/8/99 15/7/99 2/7/99 2/7/99 9/7/99 9/7/99 17/8/99 2/7/99 3/6/99 3/6/99 – 4/6/99 5/8/99 1/6/99 5/6/99 5/8/99 24/4/99 27/7/99 –

25/10/99 28/10/99 2/11/99 18/1/00 2/11/99 18/1/00 3/11/99 3/11/99 3/11/99 28/1/00 5/11/99 17/11/99 19/11/99 19/11/99 24/11/99 24/11/99 2/12/99 2/12/99 22/12/99

Crop age (days)

Growth stage

Leaf sample size

080 105 123 200 116 193 078 124 153 239 – 166 106 171 172 111 222 128 –

2–3 leaf 3 leaf 4 leaf 70% tops downB 4 leaf 5% tops down 3–4 leaf 4 leaf 4–5 leaf > 70% tops down 3–4 leaf stage Bulbing Bulbing Bulbing Bulbing Bulbing Bulbing Bulbing 1-week from lifting

0880 1000 1000 1000 1000 1000 1000 1000 1000 1000 1000 1000 0970 1000 1000 1000 1000 1000 1000

Incidence (%) Incidence (%) of of B. allii in B. allii in bulbs (and bulb leaves sample size) 0.0 0.0 0.1 3.2 0.0 0.5 0.1 0.0 0.3 5.8 0.0 0.2 0.0 0.0 0.3 0.0 0.2 0.0 0.0

0.4 (524)0 – 2.9 (547)0 2.6 (537)0 4.8 (440)0 2.5 (636)0 15.6 (900)0 – – – 8.5 (1122) – – 16.3 (1132) – –

A B

Second harvest. 70% of plants with leaves fallen over as a result of maturity.

to harbour B. squamosa. Senescent leaves were avoided because of the potential for saprophytes to mask the presence of B. allii, as demonstrated by Kohl et al. (1997). Additional leaf samples were collected from three crops (C, D and G) in January of 2000. Leaf samples were processed in the laboratory on the day of collection. Five leaves from each onion bed were placed into a non-sterile 1 L Genfac plastic food tray (length 16 cm, breadth 10 cm and height 6 cm), lined with a moistened tissue and sealed with a lid. The trays were incubated at room temperature (15–20°C) on the laboratory bench for 7 days during which time leaves senesced. Leaves were observed using a dissecting microscope (40× magnification) for the presence of Botrytis conidiophores. Under laboratory conditions, B. allii produced short (< 1 mm) conidiophores that were rarely branched. By comparison, B. cinerea produced long (> 2 mm) branched conidiophores. To confirm identification, 110 isolations were made from conidiophores in a laminar flow cabinet under a dissecting microscope (40× magnification). Conidia were removed from conidiophores with a sterile needle and placed onto pectin agar in a Petri plate under sterile conditions. Pectin agar was composed of NH4H2PO4 (0.9 g/L), (NH4)2HPO4 (2.0 g/L), MgSO4.7H2O (0.1 g/L), KCl (0.5 g/L), citrus pectin (10 g/L) and agar (30 g/L). Ingredients in distilled water were mixed in a blender until the pectin dissolved (2 min), adjusted to pH 4.0 with HCl, autoclaved and poured into Petri plates. Plates were incubated in the dark at 20°C. Botrytis isolates were identified to species by their characteristic morphology on Pectin agar. Conidiophores of B. allii formed as a mass across the surface of Pectin agar plates whereas B. cinerea produced tufts of conidiophores in sparse patches. B. allii occasionally formed sclerotia on agar after a long period, but B. cinerea developed sclerotia rapidly (within 7 days). Conidiophores and conidia were also examined using a compound microscope (400–1000× magnification) and were compared with published accounts of Botrytis (Hancock and Lorbeer 1963; Lacy and Lorbeer 1995). To compare the number of infected leaves with the incidence of bulb rot caused by B. allii, bulb samples (440 to 1122 bulbs per crop) were collected from each of eight crops (Table 1). The bulb samples were

collected from bins after harvest and before grading. Bulbs were stored for 4 months in half-tonne wooden crates in an insulated shed at 10–15°C and relative humidity of 65–98%. Bulbs were cut in half longitudinally and the incidence of neck rot recorded. Neck rot was differentiated from other rots by the characteristic slight water-soaked appearance and the presence of B. allii conidiophores and/or sclerotia. In contrast, bacterial rots produced a more water soaked lesion and had a characteristic strong odour.

Results Survey of crops for B. allii B. allii was detected in leaf samples collected from 6 of 16 crops surveyed from October to December 1999 (Table 1). The incidence of B. allii ranged from 0–0.3% with a mean of 0.2%. In the three crops that were re-sampled in January 2000, incidence increased from 0.1 to 3.2% in crop C, 0 to 0.5% in crop D and 0.3 to 5.8% in crop G (Table 1). Infections in the field were symptomless except for crop O at Scottsdale, where bulbs occurred with mycelium and conidiophores characteristic of B. allii around the base of the bulb just below the soil line. Isolations onto Pectin agar confirmed the presence of B. allii. Botrytis neck rot occurred in bulbs from all eight crops sampled at an incidence of 0.4–16.3% (Table l), with an average incidence of 5.3%. In four of the eight crops, B. allii was detected in the earliest leaf sampling and in bulbs. In the other four crops, B. allii was not detected in the earliest leaf sampling but was subsequently detected in bulbs (Table 1). For the eight crops from which leaf samples and bulb samples were assessed, the correlation coefficient between

Botrytis species and onion bulb rot

the incidence of B. allii in first leaf samples and the incidence of bulb rot was r = 0.83 (P < 0.05). A linear regression analysis gave an adjusted R2 value of 0.63 (P = 0.01) for the regression equation y = 2.74 + 45.3x, where y = the incidence of neck rot in bulbs (%) and x = the incidence of B. allii in the early-season leaf sample (%). Discussion This is one of the first studies to attempt to detect latent infections of B. allii in onion foliage during the growing season. B. allii was common in commercial Tasmanian onion crops, occurring in leaf samples from 6 of 16 crops sampled. B. allii was detected in bulb samples from all eight crops from which bulb samples were subsequently sampled, but was detected only in early to mid-season leaf samples from four of these crops. This indicates the difficulty in detecting B. allii reliably in crops from leaf samples taken from crops pre-bulb initiation to bulbing. There would be approximately 520000 plants in the 0.8 ha sample area from which 1000 leaves were collected. At this sampling intensity, if no infected leaves were found, the incidence of infection would be estimated to be no more than 0.53% (Johnson and Kotz 1969). Maude (1983) reported a 1:1 relationship between the incidence of B. allii in seed at planting and bulbs at harvest in dry environments not conducive to secondary spread. However, in Tasmania, Australia, regular rainfall events through the season are conducive to secondary spread, and the incidence of plants infected with B. allii can increase exponentially during crop growth (Chilvers et al. 2001). Weather conditions during and after lifting can also have a major influence on the transmission rate of B. allii from infected leaves to bulbs. Rapid curing can prevent infections residing in leaves from entering the necks of the bulbs (Maude et al. 1984). Therefore, the incidence of B. allii detected early in the crop life is unlikely to be always well correlated with the incidence of bulb rot at harvest. The incidence of B. allii in foliar samples in three crops which were taken just prior to harvest gave a better agreement with the incidence of B. allii in bulbs in comparison with samples taken earlier in the crop life. B. allii was symptomless in all crops except for crop O, where B. allii conidiophores were found below the soil on immature onion bulbs that did not have any rot symptoms. B. allii was not found in a neighbouring crop (P), located approximately 50 m away that had been established with a different seed line. The source of infection in crop O may have been the seed line that was used to establish the crop, but a seed sample was not available for testing. The basal infections observed may have occurred as a result of conidia being washed from infected leaves down into the soil. Alternatively, as infected outer leaves senesce, the fungus may have moved downward into the outer scales and produced conidiophores under conducive environmental conditions. Infection of the base of bulbs may also result

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from B. allii residing in the soil from previously infected crops in the form of sclerotia, or perhaps mycelium. This is less likely as onions had not been grown at the site for over 4 years. Maude et al. (1982) found that a proportion of sclerotia of B. allii remained viable in sterile soil for more than 3 years, but lost viability within 6 months in non-sterile soil. However, at an uncultivated site under Tasmanian conditions, sclerotia of B. allii remained viable in field soil for more than 29 months (M. I. Chilvers, unpublished data). Similarly for B. squamosa, Ellerbrock and Lorbeer (1977) found that after 21 months, 83 and 43% of sclerotia buried at 15 cm and 3 cm, respectively, were viable. Seed is a potential source of infection into Tasmanian onion crops. Seed testing and fungicide treatment to control damping-off fungi and B. allii are routinely used to reduce the likelihood of infection. However, current seed testing protocols often detect only down to approximately 1% incidence at 99% confidence. Treatment of seed with fungicide may reduce the incidence of initial inoculum of B. allii either by eradication or reducing the rate of seed to seedling transmission. However, fungicide treatment may not result in complete control. For example, Maude and Presly (1977b) showed that seed with an incidence of 43% B. allii that was treated with fungicide gave rise to 0.1% infected seedlings shortly after emergence. No alternative hosts for B. allii outside the Allium genus have been reported. However, other Allium crop and weed species are a potential source of inoculum for commercial crops. B. allii has been reported to infect common onion (A. cepa), garlic (A. sativum), leek (A. porrum), shallot (A. cepa var. ascalonicum), and potato or multiplier onions (A. cepa var. aggregatum) (Lacy and Lorbeer 1995). B. allii also infects weed species of Allium, including A. vineale (crow garlic) and A. ursinum (ramson) (Maude and Presly 1977b). A. vineale is recorded as a weed in Tasmania. Two other Allium weeds, A. triquetrum (three-cornered garlic) and A. neapolitanum occur in Tasmania (Curtis and Morris 1994), but their status as hosts of B. allii is not known. Tichelaar (1967) reported B. allii to survive as a saprophyte on decaying plant materials (cereals, lucerne, bean and pea). However, this report has not been further substantiated, and other authors (Kohl et al. 1997) have shown B. allii to be out-competed by other organisms on senescent plant tissue. Acknowledgements The authors gratefully acknowledge funding by the Tasmanian Onion Industry Panel (Field Fresh Tasmania Pty Ltd, Forth Farm Produce Pty Ltd, Perfecta Produce Pty Ltd, Tasmanian Farmers and Graziers Onion Commodity Group) and Australian Research Council Australian Postgraduate Award (Industry). Special thanks to the field officers of the above companies and Tasmanian onion growers for access to their crops.

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Maude RB, Presly AH (1977a) Neck rot (Botrytis allii) of bulb onions. I. Seed-borne infection and its relationship to the disease in the onion crop. The Annals of Applied Biology 86, 163–180. Maude RB, Presly AH (1977b) Neck rot (Botrytis allii) of bulb onions. II. Seed-borne infection in relationship to the disease in store and the effect of seed treatment. The Annals of Applied Biology 86, 181–188. Maude RB, Shipway MR, Presly AH, O’Connor D (1984) The effects of direct harvesting and drying systems on the incidence and control of neck rot (Botrytis allii) in onions. Plant Pathology 33, 263–268. Nielsen K, Justesen AF, Funck Jensen D, Yohalem DS (2001) Universally primed polymerase chain reaction alleles and internal transcribed spacer restriction fragment length polymorphisms distinguish two subgroups in Botrytis aclada distinct from B. byssoidea. Phytopathology 91, 527–533. Nielsen K, Yohalem DS (2001) Origin of a polyploid Botrytis pathogen through interspecific hybridization between Botrytis aclada and B. byssoidea. Mycologia 93, 1064–1071. Nielsen K, Yohalem DS, Jensen DF (2002) PCR detection and RFLP differentiation of Botrytis species associated with neck rot of onion. Plant Disease 86, 682–686. Stewart A, Franicevic SC (1994) Infected seed as a source of inoculum for Botrytis infection of onion bulbs in store. Australasian Plant Pathology 23, 36–40. Tichelaar GM (1967) Studies on the biology of Botrytis allii on Allium cepa. Netherlands Journal of Plant Pathology 73, 157–160. Yohalem DS, Nielsen K, Nicolaisen M (2003) Taxonomic and nomenclatural clarification of the onion neck rotting Botrytis species. Mycotaxon 85, 175–182.

Received 24 October 2003, accepted 2 April 2004

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