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© 2006 Plant Management Network.
Accepted for publication 7 August 2006. Published 27 November 2006.


Detection and Identification of Botrytis Species Associated with Neck Rot, Scape Blight, and Umbel Blight of Onion


Martin I. Chilvers, Postdoctoral Research Associate, PO Box 646430, Department of Plant Pathology, Washington State University, Pullman 99164-6430; and Lindsey J. du Toit, Associate Scientist/Vegetable Seed Pathologist, 16650 State Route 536, Washington State University Mount Vernon NWREC, Mount Vernon 98273-4768


Corresponding author: Lindsey du Toit. dutoit@wsu.edu


Chilvers, M. I., and du Toit, L. J. 2006. Detection and identification of Botrytis species associated with neck rot, scape blight, and umbel blight of onion. Online. Plant Health Progress doi:10.1094/PHP-2006-1127-01-DG.



Primary Economic Host: Onion (Allium cepa L.)


Diseases: Neck rot, scape blight, and umbel blight


Pathogens: Botrytis allii (Munn) Yohalem, B. aclada (Fresen.) Yohalem, B. byssoidea J.C. Walker, and B. squamosa J.C. Walker


Taxonomy

Seven Botrytis spp. have been associated with diseases of Allium crops (Table 1) (19,34). Four of these species have known teleomorphs in the genus Botryotinia, family Sclerotiniaceae, order Helotiales, and class Discomycetes (21). Three species are considered the primary causal agents of neck rot of onion, namely B. aclada, B. allii, and B. byssoidea (22,55). Although B. squamosa and B. tulipae have also been associated with neck rot, these species are not typically primary causes of neck rot (34,45,46).


Table 1. Species of Botrytis associated with onion crops.

Anamorph Teleomorph Onion diseasesx
B. aclada (Fresen.) Yohalem Unknown Neck rot, bulb rot, scape blight, and umbel blight
B. allii (Munn) Yohalem Unknown Neck rot, bulb rot, scape blight, and umbel blight
B. byssoidea J.C. Walker Botryotinia allii (Sawada) Yamamoto Mycelial neck rot
B. cinerea Pers.:Fr. Botryotinia fuckeliana (de Barry) Whetzel Botrytis brown stain (cosmetic), Botrytis leaf blight, umbel blight
B. porri Buchw. Botryotinia porri (van Beyma Thoe Kingma) Whetzel Neck rot of leek and garlic, but recently detected in onion seed crops and onion seedy
B. squamosa J.C. Walker Botryotinia squamosa Vien.-Bourg. Botrytis leaf blight, neck rot, umbel blight
B. tulipae (Lib.) Lind Unknown Neck rot, bulb rot

 x Adapted from Mohan and Schwartz (34).

 y Although primarily a pathogen of garlic and leek, B. porri has been isolated from naturally-infected onion seed crops and onion seed (11,12).


Until the recent taxonomic and nomenclatural clarification by Yohalem et al. (55) of the primary onion neck rot species of Botrytis, B. aclada (Fresen.) Yohalem was considered by many to be synonymous with B. allii (Munn) Yohalem (21), while B. byssoidea J.C. Walker was regarded by some as conspecific with B. aclada (25). The lack of distinction among these species was due, in part, to limitations at differentiating these species using classical morphological and cultural methods (41,42). Nonetheless, Owen et al. (39) demonstrated that B. byssoidea and B. aclada are valid species. Two subgroups within B. aclada (AI and AII) have been distinguished based on chromosome number and conidial dimensions (20,47). More recently, polymerase chain reaction (PCR) alleles and internal transcribed spacer restriction fragment length polymorphisms (ITS-RFLPs) have been used to demonstrate that isolates in subgroups AI and AII are distinct from B. byssoidea (36,37,38). Nielsen and Yohalem (37) concluded that the larger-spored subgroup (AII) developed as a result of a hybridization event between a small-spored isolate of B. aclada and an isolate of B. byssoidea. Yohalem et al. (55) proposed that B. aclada be reserved for the small-spored subgroup (AI), and B. allii for the larger-spored subgroup (AII) of B. aclada. A recent evaluation of the molecular phylogeny of the genus Botrytis, based on three nuclear protein-coding genes (RPB2, G3PDH, and HSP60), supports this proposed hybrid status (48). Molecular markers were used in that study to revise the phylogenetic relationships within the whole Botrytis genus, and may aid in identifying Botrytis species (48).


Symptoms and Signs

Onion bulb crops. Symptoms of neck rot typically develop only after onion bulbs have been harvested and placed in storage, even though infection occurs in the field (28,29). The predominantly latent nature of field infections (12,25) complicates efforts to manage this disease and to predict losses in storage. Under very moist field conditions, Botrytis conidiophores and conidia may be visible on senescent plant tissues. However, differentiating Botrytis species associated with onion tissues requires culturing the fungi on agar media, and/or measurement of conidial dimensions using a compound microscope. During storage, infected onion bulbs develop a semi-watery decay which usually begins inside the neck with no external symptoms visible (Fig. 1A). Infection may spread through the entire bulb (Fig. 1B). Infection can occur at wound sites on other areas of the bulb, in which case the disease may be referred to as shoulder rot, side rot, or bottom rot (Fig. 1C to 1D). White mycelium develops between bulb scales, producing compact gray masses of conidiophores and conidia (Fig. 1D). Dark brown to black, typically flattened sclerotia may form between scales in the neck of the bulb or at other sites of infection (Fig. 1B) (25). Brown stain, caused by B. cinerea, differs from neck rot in that infection is typically limited to the outermost dry scales of the bulb, resulting in a cosmetic brown blemish on the surface of the bulb (Fig. 2) (7). In addition to post-harvest losses from neck rot, B. aclada and B. allii can reduce plant stands and vigor (8), particularly when plants are injured (e.g., by windblown sand, hail, or chemicals) during cool and moist conditions (Fig. 1E to 1G).


 

Fig. 1. Common postharvest symptoms (A to D), and less common preharvest (E to G) symptoms, caused by Botrytis aclada and B. allii on onion bulbs. (A) Internal symptoms of neck rot, including watersoaking and discoloration of the inner scales with white to gray mycelium in the neck. (B) Severe neck rot showing decay of the fleshy scales, shrinkage of the bulb, and black sclerotia (arrow) forming in the neck and shoulders of the bulb. (C) Soil line rot resulting from infection of the bulb at the soil surface (photo courtesy of G.Q. Pelter). (D) Dense sporulation of B. aclada or B. allii on the dry outer scales of a bulb from a cull pile. (E) Sporulation of B. aclada or B. allii on the surface of an onion plant under moist field conditions. (F) Neck rot developing on a plant under particularly moist field conditions close to harvest (healthy plant on the left). (G) Sporulation of B. allii or B. aclada revealed by pulling back the outermost fleshy scale of an infected plant after very moist field conditions.

 

 

Fig. 2. Brown stain, also known as coffee stain, on the outer dry scales of an onion bulb caused by Botrytis cinerea (photo courtesy of Field Fresh Tasmania, Australia).

 

Onion seed crops. In onion seed crops, blighting and girdling of the scapes (seed stalks), and umbel and flower blight (Fig. 3) have been associated primarily with B. allii (usually not differentiated from B. aclada), but also with B. byssoidea, B. cinerea, and B. squamosa (1,9,12,13,17,23,31,35, 44,45,46,54,55). Lesions may develop at any point along the scape, typically girdling the scape and causing the scape to lodge, particularly during windy conditions. Scape blight lesions are initially elongated to diamond-shaped and chlorotic, becoming necrotic as they mature. Gray concentric rings may become evident as the fungus produces spores in concentric rings beneath the outer cuticle and wax layers of the scape (Fig. 3B and 3C). Scape and umbel blights reduce seed yield and quality, and may result in infected seed. Seedborne inoculum of B. allii and B. aclada can be a source of infection resulting in outbreaks of neck rot (28,29,51).


 

Fig. 3. Symptoms of umbel (A), scape (A to C), and flower blight (D) caused by Botrytis aclada and B. allii. Lesions can occur at the top (A), middle (B), or lower end (C) of the scape, often resulting in lodging once the lesion has girdled the scape, which affects seed set. Concentric grey rings may form in the girdling lesions as a result of sporulation of the fungus beneath the outer cuticle. The fungi sporulate on senescing tissues, including the spadix (membranous sheath that covers the immature umbel), anthers, and petals (D).

 

Host Range

The host range of Botrytis spp. associated with neck rot, scape blight, and umbel blight appears to be limited to most Allium spp., i.e., bulb onion (A. cepa), garlic (A. sativum), leek (A. porrum), shallot (A. cepa var. ascalonicum), potato or multiplier onions (A. cepa var. aggregatum), and possibly Allium weeds (25). However, Tichelaar (50) observed B. allii growing saprophytically on decaying plant material of cereals, flax, lucerne, bean, and pea. Chilvers (3) demonstrated that, in the absence of competition, B. allii was capable of colonizing and sporulating profusely on sterilized poppy (Papaver somniferum) debris.


Geographic Distribution

Onions are an important food crop worldwide. In 2005, approximately 65,000 hectares of storage and non-storage onions were harvested in the US, representing a farmgate value of $922 million (USDA National Agricultural Statistics Service). Losses to the onion industry from neck rot occur worldwide, but are variable and sporadic with some seasons bringing greater than 50% losses to individual crops (10,28,29,31). Neck rot species of Botrytis are distributed in all areas of the world where onions are produced, but greatest losses have been reported from temperate regions (4,5,10,12,28,29,31,36,38). Although B. allii and B. aclada are the predominant species reported to cause neck rot of onion, these species are difficult to distinguish morphologically because of similar growth patterns on agar media, and overlapping spore sizes (55). Although B. byssoidea is only occasionally reported as causing neck rot (42,53), the distribution and significance of B. byssoidea as a neck rot pathogen may be underestimated compared to B. allii and B. aclada because of the difficulty of identifying B. byssoidea, which sporulates sparsely on most media used to isolate fungi from plant tissues (41,42).

Previous reports on neck rot typically have not differentiated between B. allii and B. aclada, making it difficult to discern the relative distribution and significance of these two species as neck rot pathogens. However, since description of a PCR-RFLP assay by Nielsen et al. (38) for differentiation of the primary neck rot Botrytis spp., B. allii and B. aclada can be differentiated more readily. Nielsen et al. (36) reported that of 29 isolates collected from Belgium, Canada, Denmark, England, Germany, Japan, the Netherlands, Norway, Scotland, and the US, and identified morphologically as B. allii or B. aclada, 14 and 15 of the isolates were identified as B. aclada and B. allii, respectively, using the PCR-RFLP assay. An additional seven isolates were identified morphologically as B. byssoidea, but PCR-RFLP analysis determined one isolate to be B. allii and three to be B. squamosa, highlighting the difficulty of morphological differentiation of these species (36). Using the PCR-RFLP procedure, Chilvers et al. (5) examined 23 isolates from onion crops in Australia that had been identified morphologically as B. allii or B. aclada. The results differentiated 22 of the isolates as B. allii and 1 as B. aclada. du Toit et al. (12) collected 469 isolates identified morphologically as B. allii (undifferentiated from B. aclada) from onion seed crops and onion seed in central Washington. Using PCR-RFLP analysis, 60 and 40% of the isolates were identified as B. aclada and B. allii, respectively (Chilvers and du Toit, unpublished). These studies demonstrate that both B. allii and B. aclada appear to be distributed worldwide. Further research is needed to evaluate B. aclada, B. allii, and B. byssoidea for differences in epidemiological significance as neck rot pathogens.

B. squamosa is primarily a leaf blight pathogen of onion in temperate regions of bulb or seed production during seasons with high relative humidity and rainfall (44,45,46). Although B. squamosa is occasionally associated with neck rot, scape blight, or umbel blight in these regions, this species does not appear to be associated with neck rot, scape blight, or umbel blight in the semi-arid regions of onion bulb and seed production in the western US (12,17,24).


Pathogen Isolation

Plants. An effective method of inducing sporulation of Botrytis spp. on onion plants with latent infections by these fungi was described by du Toit et al. (12). Whole plant samples were placed in plastic bags in a cool (5 to 10°C) dark environment for 4 to 7 weeks. The plants were then removed from cold storage, cut longitudinally, and placed into clear plastic containers lined with moist paper towel to create moist chambers. The plants were incubated in the moist chambers at ambient temperature (24 to 28°C) on approximately a 12-h/12-h day/night cycle for 5 to 7 days, and then examined macroscopically and microscopically for development of Botrytis conidiophores and conidia (Fig. 4). Surface-sterilizing the plants with 0.5% NaOCl for 60 s, then rinsing the plants with tap water and incubating them in a moist chamber, did not affect the efficacy of this protocol for triggering latent infections to become active and promote sporulation of Botrytis spp. (12). The duration of cold storage needed to trigger latent infections has not been determined, but Kritzman (23) demonstrated that storing onion bulbs for 2 days at 4°C, followed by 16 days at 15°C, activated latent infections by B. allii (undifferentiated from B. aclada). This, in turn, altered the pH of the infected bulb tissue. The change in pH was then visualized by cutting the bulbs longitudinally and staining them with an alcoholic solution of bromocresol green and methyl red. Healthy bulbs stained green but infected bulbs stained red. Alternatively, detached onion leaves can be placed directly into plastic containers lined with moist paper towel, and incubated at room temperature for 7 days. Botrytis conidiophores and conidia develop as infected tissues senesce (4,28).


 

Fig. 4. Detection of latent infection by Botrytis aclada and B. allii on asymptomatic onion plants sampled from an onion seed crop, placed in cold storage (5 to 10°C) for four weeks, and then incubated in a moist chamber for 5 to 7 days. (A) Dense sporulation on an onion scape (noninfected scape on the right). (B) White mycelium and gray sporulation on a scape. (C) Conidiophores and conidia (48× magnification) of B. aclada or B. allii. (D) Sclerotia and sporulation of B. aclada or B. allii on an infected scape.

 

Pure cultures of Botrytis spp. can readily be established by transferring conidia onto agar media from the host material on which the fungi are sporulating. Potato dextrose agar (PDA) or similar general media support growth of all Botrytis spp. associated with Allium crops, although media differ in the propensity to promote sporulation. Agar media may be amended with antibiotics (e.g., streptomycin at 100 ppm) to reduce bacterial contamination. Single-spore isolates can be obtained by streaking conidia, or by dilution plating (18).

Seed. Various procedures can be used to isolate Botrytis spp. from onion seed, including plating seed onto agar media or incubating seed on damp blotters. However, there is currently no certified, internationally-accepted protocol for assaying onion seed for neck rot fungi. This compounds the controversy over the relative significance of seedborne inoculum vs. alternative sources of inoculum (infested onion residues, culled onions, infected volunteers, and soilborne sclerotia) of B. allii or B. aclada in development of neck rot in storage (12,25,26,28,29,31,51). The controversy is not addressed by this paper. The most common difficulty when assaying onion seed for Botrytis spp. is the presence of fast-growing fungi that outgrow Botrytis spp. present in or on the seed. In particular, species of Rhizopus and Aspergillus may prevent, or significantly limit, detection of Botrytis on some onion seed lots, unless selective agar media are used (Table 2). Two versions of an agar media assay for detecting Botrytis spp. associated with onion seed were described by du Toit et al. (12):

(i) Surface (non-sterilized) seed assay: To assay onion seed for infestation by Botrytis spp., onion seed was rinsed under running deionized water for 60 min to remove spores of fast-growing contaminant fungi present on the seed surface, and to assist with removal of fungicides on the surface of treated seed. The rinsed seed was then dried in a laminar flow hood on sterile paper towel, and plated onto Kritzman’s agar (22) with 20 seeds spaced uniformly in each 10-cm-diameter Petri plate (Fig. 5). The plates were then sealed with Parafilm and incubated at 20°C with a 12-h/12-h day/night cycle (cool white fluorescent light and near ultraviolet light by day) for 14 days. The seed were examined microscopically (up to 100× magnification) at approximately 4-day intervals for up to 14 days after plating, for development of Botrytis spp. (12).

(ii) Internal (surface-sterilized) seed assay: To differentiate surface-contamination (infestation) from infection of onion seed by Botrytis spp., onion seed can be surface-sterilized using a disinfectant, rinsed thoroughly, and assayed for Botrytis spp. du Toit et al. (12) rinsed onion seed in 0.5% NaOCl with agitation for 60 s, triple-rinsed the seed in sterile deionized water, and then air-dried, plated, and examined the seed as described above for the non-sterilized seed assay. The particular disinfectant used, the concentration of the disinfectant, and the duration of surface-sterilization all affect the results of such seed assays (12,22,28,31,32,51).


 

Fig. 5. Onion seed assayed for Botrytis spp. on Kritzman’s agar (22). (A) Concentric rings of sporulation with darkening of the agar medium resulting from polyphenoloxidase (PPO) activity of B. allii or B. aclada. (B) B. aclada or B. allii sporulating on an onion seed. (C) Fast-growing, flat, white, fan-shaped or ‘ropy’ mycelium of B. porri that sporulates sparsely and also darkens Kritzman’s agar as a result of PPO activity. (D) Sporulation and a brown sclerotium of B. porri developing from an infected onion seed.

 

Table 2. Efficacy of four agar media for determining the percentage of seeds in an onion seed lot that were infected with Botrytis spp. [adapted from Metcalf (32)].

Characteristic assessed Agar mediumx
PDA PLA Kritzman Pectin
Mean percentage of seed with Botrytisy 5.3 5.4 4.4 5.2
Repeatability of nutrient consistency  +z + +++ +++
Freedom from contaminant fungi + + +++ ++
Internationally accepted for Botrytis spp. + +++ +++ +
Ease of preparation +++ + + ++
Ease of identifying Botrytis ++ ++ +++ ++

 x PDA = potato dextrose agar, PLA = prune lactose agar, Kritzman = agar medium developed by Kritzman and Netzer (22), Pectin = pectin agar (33). Refer to the appendix for details of the media.

 y Three replications of 550 seeds were assayed on each medium (standard deviation = 1.5%).

 z + = low, ++ = good, +++ = very good.


Soil. To isolate Botrytis spp. from soil, aliquots of a dilution series of soil samples suspended in water or dilute water agar (approximately 2% to help keep the soil in suspension) can be plated onto selective media such as those described by Kritzman and Netzer (22) or Lorbeer and Tichelaar (27). For the Kritzman and Netzer (22) medium, polyphenoloxidase (PPO) activity of Botrytis spp. converts the tannin in the medium to a dark brown pigment (Fig. 5). This, in combination with the antibiotic and fungicide amendments that inhibit or slow development of other fungi, makes the medium very selective for identification of Botrytis spp. (22). Kritzman and Netzer (22) found a positive correlation between inoculum concentration in soil and the number of brown-pigmented colonies observed on the medium. They were able to identify brown-pigmented colonies within 48 h of incubation at 24°C, and each of the colonies was verified as B. allii (not differentiated from B. aclada) upon sporulation after an additional 4 days of incubation. Kritzman’s agar has been used by others to isolate Botrytis spp. from soil (3,30).


Differentiation of Botrytis spp. Isolated from Onion Plants

Macroscopic identification. Botrytis spp. are readily cultured on various agar media. Differentiation among the species associated with onion based on morphology is possible for most species, except to differentiate B. allii from B. aclada (Table 3, Fig. 6) (41,42). However, accurate species identification based on macroscopic morphology is difficult for non-sporulating cultures. Nielsen et al. (36) found that four of seven non-sporulating isolates initially identified using morphological characteristics as B. byssoidea, were later classified by DNA fingerprinting techniques as B. allii or B. squamosa. Spore production by these species is enhanced by near-ultraviolet light and by adding onion tissue to the medium (42).


Table 3. Morphological characteristics of Botrytis spp. isolated from onion crops when cultured on potato dextrose agar.

Botrytis
species
Morphological characteristicsw
Optimum temp.
(°C)
for sporula-
tion
Production of conidiophores
& conidia
Production of sclerotia Colony color & morphology Relative rate of growthx
B. aclada 20 +
(abundant short conidiophores, about 1 mm long)
Tan (dense mass of conidiophores) Intermediate
B. allii 20 +
(abundant short conidiophores, about 1 mm long)
Tan (dense mass of conidiophores) Intermediate
B.
byssoidea
15 y White to off-white mycelium Intermediate
B. cinerea 15 +z
(long, frequently 2 mm or more, dematiaceous, branched conidiophores)
+
(medium to large, smooth-surfaced, light to dark brown)
White mycelium, gray to brown colony as conidiophores mature Fast
B. porri + +
(medium to large, brown, often cerebriform)
White, flat, ‘ropy’ mycelium Fast
B.
squamosa
5 y +
(abundant, small, black)
White mycelium that collapses onto agar surface Intermediate

 w Adapted from Presly (41,42), Ellis (14), and from observation of cultures on potato dextrose agar (PDA) (Fig. 6).

 x Intermediate vs. fast growth rates = 6 to 10 vs. 3 to 6 days, respectively, for the fungus to cover the surface of a 10-cm diameter plate of PDA from a colonized 5 mm-diameter plug placed in the center of the plate (du Toit, unpublished).

 y B. byssoidea and B. squamosa produce spores sparsely when exposed to near-ultraviolet light, but do not produce spores when cultured in the dark. B. squamosa produces a very marked concertina-like collapse of conidiophores.

 z B. cinerea produces abundant spores when exposed to near-ultraviolet light, but very few spores in the dark. Sealing plates with Parafilm also may inhibit sporulation of B. cinerea.


 

Fig. 6. Appearance of isolates of Botrytis aclada, B. allii, B. byssoidea, B. cinerea, B. porri, and B. squamosa on potato dextrose agar (PDA) after 9 or 13 days of growth at ambient temperature (25 ± 3°C).

 

Microscopic identification. Botrytis spp. associated with onion plants can be differentiated to some degree based on conidial dimensions. However, spore dimensions of isolates of the three predominant neck rot species overlap, complicating efforts to differentiate these species morphologically (Table 4, Fig. 7).


Table 4. Dimensions of conidia, average number of nuclei/conidium, and number of chromosomes/nucleus for Botrytis spp. isolated from onion or leek crops.

Botrytis species Length of
conidia
(µm)
Width of
conidia
(µm)
Average number of nuclei/
conidium
w
Number of chromo-
somes
/
nucleus
w
B. aclada 8.4 - 12.1u
6.5 - (8.6) - 11.0v
4.6 - 5.6u
4.0 - (4.6) - 6.0v
1.4 16
B. allii 9.7 - 14.1u
9.0 - (10.2) -15.0v
5.3 - 7.3u
5.0 - (5.7) - 6.5v
1.5 32
B. byssoidea 12.0 - 14.0w
12.0 - 15.0x
8.0 - 9.0w
6.5 - 8.5x
5.1 16
B. cinerea 9.0 - 11.0w
6.0 - 18.0y
6.0 - 8.0w
4.0 - 11.0y
4.6 16
B. porri 14.5 - 16.5x 8.5 - 11.0x z
B. squamosa 21.0 - 22.5x 16.5 - 17.0x 18.4 16

 u Nielsen et al. (36).

 v Yohalem et al. (55), with the mean dimension reported in parentheses.

 w Shirane et al. (47).

 x Presly (41).

 y Ellis and Waller (15).

 z “—” = unknown.


 

Fig. 7. Microscope photos (200x magnification) of conidia of each of six Botrytis species isolated from onion plants and grown on potato dextrose agar (PDA). Cultures were maintained for 7 to 14 days at ambient temperature (25 ± 3°C) with a 12 h/12 h day/night cycle (cool white fluorescent light), except for B. byssoidea and B. squamosa which were maintained at 15 ± 2°C under constant near-ultraviolet light for 7 days.

 

Key to Botrytis species on Allium based on macroscopic and microscopic characteristics. Use of this key, together with matching the Botrytis characteristicsx observed with those shown in Table 3, Table 4, Fig. 6 and Fig. 7 will enable identification of most isolates:


1. Sclerotia readily produced on PDA . . . .  (B. cinerea, B. porri, B. squamosa)

 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

1'. Sclerotia not produced on PDA . . . .  (B. aclada, B. allii, B. byssoidea)

  . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

2. Conidia longer than 20 μm and wider than 16 μm; sclerotia black and 1 to 3 mm in diameter

 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. squamosay

2'. Conidia shorter than 20 μm and narrower than 16 μm; sclerotia medium to dark brown and variable in size . . . .  (B. cinerea, B. porri)

 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

3. Conidia have a minimum length of 12 μm; very sparse sporulation on PDA; mycelium white to off-white on PDA

  . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. byssoidea

3'. Most conidia shorter than 12 μm; dense gray mass of conidiophores and conidia on PDA; sparse mycelium on PDA . . . .  (B. aclada, B. allii)

 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

4. Conidia have a minimum length of 14 μm and a minimum width of 8 μm; conidiophores about 1 mm long; sclerotia smooth to cerebriform

 . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . B. porri

4'. Conidia often shorter than 14 μm and often narrower than 8 μm; conidiophores typically 2 mm or longer; sclerotia smooth-surfaced

  . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. cinerea

5. Conidia have a maximum length of 14 to 15 μm, and mean dimensions of 10.2 × 5.7 μm

  . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. allii

5'. Conidia have a maximum length of 11 to 12 μm, and mean dimensions of 8.6 × 4.6 μm

 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. aclada

 x Excluding B. globosa, which has spherical to sub-spherical conidia, commonly 12 to 18 μm long, and which occurs on wild garlic, Allium ursinum; excluding B. sphaerosperma, which has spherical conidia commonly 20 to 26 μm long, and which occurs on three-cornered leek, Allium triquetrum; and excluding B. tulipae (records on Allium are extremely infrequent).

 y In Ellis (14), conidia of B. squamosa are more highly variable in length than indicated in Table 4, but are up to 26 μm long; and also highly variable in width, but up to 18 μm wide, i.e., in spite of variability, the maximum length and maximum width of conidia exceed those of other species.


Molecular Identification and Botrytis Detection

PCR detection and RFLP differentiation of Botrytis spp. causing neck rot.  Nielsen et al. (38) developed the following DNA primers for amplification and subsequent differentiation of five Botrytis spp. associated with neck rot, using sequence characterized amplified regions (SCARS):

BA2f: 5’-GTGGGGGTAGGATGAGATGATG-3’
BA1r: 5’-TGAGTGCTGGCGGAAACAAA-3’

Using a polymerase chain reaction (PCR) assay, the primers amplify a 413 bp DNA product from all five neck rot species. Restriction fragment length polymorphism (RFLP) analysis of the PCR amplicon using the restriction enzyme ApoI distinguishes the following diagnostic DNA bands (Fig. 8) (38):

B. aclada AI = 413 bp,
B. allii (B. aclada AII) = 413 + 298 bp,
B. byssoidea = 298 bp,
B. cinerea = 250 bp,
B. squamosa = 269 bp.


 

Fig. 8. PCR-RFLP analysis using the protocol described by Nielsen et al. (38) for six Botrytis spp. isolated from onion plants or seed. 100 bp ladder (New England Biolabs, Beverly, MA). Arrows indicate 300 and 400 bp DNA ladder fragments.

 

Although Nielsen et al. (38) apparently did not amplify DNA of B. porri using these primers, we detected an amplicon of similar size which showed the same restriction pattern as B. cinerea for isolates of B. porri obtained from onion plants and seed in Washington (Fig. 8) (Chilvers and du Toit, unpublished).

Magnetic capture hybridization (MCH)-PCR. Although the PCR-RFLP assay described by Nielsen et al. (38) facilitates more rapid detection and differentiation of neck rot species of Botrytis compared to traditional methods of isolating the fungi onto selective media, onion seeds contain compounds that inhibit the PCR and can yield false-negative results (52). To overcome this problem, Walcott et al. (52) developed a magnetic capture hybridization (MCH)-PCR protocol for seedborne Botrytis spp. In summary, magnetic beads were coated with DNA probes based on the SCAR described by Nielsen et al. (38). The probes captured the fungal DNA from crude seed extracts, and PCR inhibitors were washed off the captured DNA, which was then amplified by PCR using the primers described by Nielsen et al. (38). The MCH-PCR assay reduced the time required to assay onion seeds from 10 to 14 days when using selective agar media, to less than 24 h. However, the agar assay yielded higher detection frequencies of B. aclada for seed lots with low infection levels (2.1%) compared to the MCH-PCR assay, and the MCH-PCR assay lacked adequate reproducibility or the ability to quantify the incidence of infection in individual seed lots. With further refinement, and in combination with quantitative real-time PCR, the MCH-PCR assay has potential as a rapid commercial method of screening onion seed lots for infection by Botrytis spp. (52).

Real-time PCR assay.  A real-time fluorescent PCR assay based on SYBR green chemistry has been developed to quantify the amount of neck rot Botrytis spp. (B. allii, B. aclada, and B. byssoidea) present in onion seed (6). The nuclear ribosomal intergenic spacer (IGS) region of target and non-target Botrytis spp. was sequenced, aligned, and used to design primers specific to B. allii, B. aclada, and B. byssoidea. Primers and amplification parameters were optimized to avoid amplifying the related species B. cinerea, B. porri, B. squamosa, and Sclerotinia sclerotiorum, as well as 15 other fungal species commonly found on onion seed. The primers were capable of detecting 10 fg genomic DNA extracted from pure cultures of B. allii and B. aclada (6). The real-time PCR assay appears to be more sensitive than plating onion seed onto selective agar media, as the PCR assay enabled DNA of Botrytis neck rot species to be detected from seed lots that tested negative using conventional seed plating assays (6).

ELISA assays. Linfield et al. (26) developed polyclonal antisera for an indirect ELISA assay to detect latent B. allii (undifferentiated from B. aclada) infection in stored onion bulbs. The ELISA enabled earlier detection of B. allii in bulb tissue compared with conventional isolations onto agar media. The antisera did not cross-react with B. byssoidea, B. porri, or B. squamosa, but showed some cross-reaction with B. cinerea. Antiserum developed to cell wall extracts of B. allii displayed less cross-reaction with B. cinerea than antiserum developed from surface washings of B. allii cultures, as the latter contained whole spores and mycelial fragments. Linfield et al. (26) also demonstrated that non-uniform infection of onion neck tissue by B. allii necessitates sampling whole slices across the neck of each bulb for more accurate detection of the incidence of bulbs with latent infection. Neogen Europe Ltd. (Auchincruive, UK) offers a commercial, non-quantitative ELISA IDENTIKIT for B. allii (undifferentiated from B. aclada). To our knowledge, this is the only commercial ELISA kit available for onion neck rot fungi.

Storage of isolates.  Isolates of Botrytis spp. can readily be stored using a number of methods, e.g., on colonized PDA slants held at 4°C (16), as spore suspensions in 15% glycerol stored at –80°C (36), or on colonized and dried filter paper stored at –20°C (40) (du Toit and Chilvers, unpublished).


Pathogenicity Tests

Inoculum production. Spores of B. allii and B. aclada can be produced readily on a variety of media, including PDA and onion leaf extract agar (37), or on infected onion tissue (senescing leaves or infected bulbs) maintained under high relative humidity and at moderate temperatures (15 to 25°C). A conidial suspension can then be prepared in water. Adding a small amount of surfactant (e.g., Tween 80 at 0.01% v/v) counteracts the hydrophobic nature of Botrytis spores in suspension. Alternatively, naturally infected, decaying onion bulbs with symptoms of neck rot can be used as inoculum, particularly as a source of sclerotia. Production of sclerotia in culture can be achieved by autoclaving 10 g unprocessed natural bran and 250 g sand (<500-μm-diameter particles) in a flask, and inoculating the contents of the flask with a conidial suspension of the pathogen. The inoculated bran-sand mixture is incubated for 6 weeks in the dark at approximately 20°C, and then the sclerotia are separated by wet-sieving the mix through a 500-μm mesh sieve (3).

Inoculum application. To inoculate onion bulbs with B. allii or B. aclada, a spore suspension of the pathogen can be injected into the neck of each bulb using a hypodermic needle and syringe, or the necks can be cut horizontally and a spore suspension or a colonized disk of agar medium applied to the freshly-cut surface of each bulb (29,42). Inoculated bulbs should be stored at ambient temperature (15 to 25°C) for 2 to 4 weeks, and then cut longitudinally and incubated at high relative humidity for about 7 days to promote sporulation of the pathogen.

To inoculate onion plants with neck rot species of Botrytis, a spore suspension of the pathogen can be sprayed onto plants. Inoculated plants are maintained under high humidity for 2 to 3 days at moderate temperatures (15 to 25°C), where possible, to promote spore germination and establishment of infection. In field trials, plants should be inoculated during cool and moist conditions that promote extended periods of leaf wetness (3,33). Alternatively, neck rot-infected bulbs can be placed within the field as a source of inoculum (3,33).

Pathogenicity tests. Yohalem et al. (55) tested the virulence of Botrytis isolates by placing an agar disk with mycelium of the appropriate isolate onto the cut neck of each onion bulb. The depth of maceration of each inoculated bulb was assessed after incubating the bulbs in a dark moist chamber at 18°C for 14 days. Stewart and Mansfield (49) placed drops of a conidial suspension of each Botrytis isolate on detached onion leaves or onion scales to test for pathogenicity.


Detecting Latent Botrytis Infection in Onion Bulbs

Latent Botrytis infections in onion bulbs are typically detected by storing harvested bulbs for 8 to 16 weeks, and then cutting the bulbs vertically to assess for internal symptoms of neck rot (4). However, the need exists for a rapid predictive test at, or prior to, harvest to determine the incidence of bulbs with latent infection. This would facilitate decisions regarding storage, processing, and sale of the bulbs. As described above, Kritzman (23) developed a method of detecting latent bulb infections by B. allii that was based on triggering latent infections to become active by changing the temperature during incubation of the bulbs. The pH in infected bulb tissues dropped to 4.4 when infection by B. allii was activated, compared to pH 5.2 for healthy tissue. The pH change was visualized using bromocresol green and methyl red, which stained infected tissue red and healthy tissue green. Kritzman (23) stated that, although culturing bulb tissue onto selective agar media provided more accurate diagnosis than the staining method, the latter enables large numbers of bulbs to be tested at very low cost without the need for laboratory facilities. However, other factors can cause a change in the pH of bulb tissues, which could result in false positives when using this staining procedure (33).

Chilvers et al. (2) attempted to predict the incidence of neck rot in onion bulbs by removing a core of neck tissue from each bulb using an apple corer, and incubating the core for 7 days in humid conditions. They were unable to detect a consistent relationship between the number of bulbs from which the sampled cores developed Botrytis spp. and the number of bulbs that developed neck rot in storage (2,3). In contrast, the indirect ELISA developed by Linfield et al. (26) gave a "good indication" at harvest of the probable incidence of neck rot in storage, for both non-inoculated plants and plants inoculated in the field with B. allii. Isolations from bulbs onto agar media consistently showed a lower incidence of latent infection detected than the ELISA, until the bulbs had been stored for several months, by which time the ELISA and conventional isolation methods gave similar results. The researchers noted the necessity of sampling whole slices across the neck of each bulb for accurate detection of latent infection, because of non-uniform infection of onion neck tissue by B. allii. This might explain the poor results for the core sampling method evaluated by Chilvers et al. (2,3).

Prithiviraj et al. (43) demonstrated that detecting volatile metabolites can be used to discriminate among onion bulbs infected by Erwinia carotovora subsp. carotovora, Fusarium oxysporum, and B. allii, and to differentiate healthy bulbs from infected bulbs. Onions inoculated with B. allii released 1-ethenyl-4-ethyl-benzene, whereas healthy bulbs and bulbs inoculated with the other two pathogens did not release this metabolite (43). Although sampling for volatile metabolites may facilitate determination of the incidence of bulbs infected with Botrytis in a bulb lot, the procedure may be limited to detection after an incubation period, combined with a protocol that triggers latent infections to become active, as described by Kritzman (23).

With any method of assessing the incidence of latent infection in onion bulbs for predicting the incidence of neck rot in storage, research is needed to determine the optimum sampling time and sample size for accurate prediction of the incidence of neck rot in storage, as noted by Linfield et al. (26). Although morphological and molecular tools are available for accurately differentiating among Botrytis spp. that infect onion plants, the predominantly latent nature of these infections remains a hurdle to managing the pathogens effectively.


Conclusions

Although traditional methods of isolating fungi onto standard agar media, followed by classic morphological and microscopic techniques, provide relatively easy and accessible methods to differentiate B. porri, B. cinerea, and B. squamosa from B. aclada and B. allii in association with onion tissues, accurate differentiation of B. allii and B. aclada is difficult using these techniques because of very similar morphological characteristics of the latter two species. Furthermore, identification of B. byssoidea is complicated by the fact that this species typically does not sporulate or form sclerotia on agar media commonly used to isolate fungi from plant tissues. Therefore, combining classic morphological and microscropic examination of cultures with newer molecular techniques, such as PCR-RFLP (38) or real-time PCR (6), will enable accurate differentiation of these species. Real-time PCR assays may prove valuable as a rapid means of detecting latent Botrytis infections in onion bulbs or in plant samples. The choice of which method(s) to use for detecting and identifying Botrytis species associated with onion samples essentially depends upon the resources available and the requirements for identification, i.e., for extension or diagnostic purposes, identifying the causal organism as one of the "neck rot species" (B. allii, B. aclada, or B. byssoidea) using isolations onto standard agar media, followed by microscopic examination of the cultures, is probably sufficient. However, confident identification to the species level may be required for detailed epidemiological or etiological research, which is possible with the combined use of morphological and molecular techniques.

Complicating this issue further is the lack of epidemiological research that demonstrates the relative significance of B. byssoidea vs. B. aclada vs. B. allii as neck rot pathogens under the diversity of environmental conditions in which onion bulb crops are grown. Until such information is generated, the three species probably should be considered of equal importance as causal agents of neck rot, and differentiation among the three species may not be as important for applied research purposes as differentiating these species from B. porri, B. cinerea, and B. squamosa. Even with the ability to differentiate accurately among all Botrytis spp. associated with onion plants, a suitable sampling strategy remains perhaps the biggest challenge to applying these techniques to estimate the incidence of onion plants or bulbs with latent infection in the field or at harvest as a predictor of the incidence of neck rot in storage.


Acknowledgments

The authors thank David Yohalem and Karsten Nielsen for valuable discussions on neck rot, Tobin Peever for providing laboratory space and support for Martin Chilvers during his postdoctoral appointment for this project, Jim Lorbeer and Gary Chastagner for providing cultures of Botrytis isolates, and Frank Dugan and Jack Rogers for reviewing the manuscript.

This work was supported by the Washington State University (WSU) IMPACT Center, WSU Agricultural Research Center, California Onion & Garlic Research Committee, Nevada Onion & Garlic Advisory Board, Pacific Northwest Vegetable Association, Columbia Basin Vegetable Seed Association, and the onion seed industry.

This paper was submitted as Plant Pathology New Series No. 0419, Department of Plant Pathology, College of Agricultural, Human, and Natural Resource Sciences Research Center, CRIS Project No. WNP00595, Washington State University, Pullman, WA 99164.


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Appendix

Onion leaf extract agar

(i) Place the following ingredients into a 2 liter flask:

  Agar 16 g
  Dried onion leaves, cut or crushed 15 g
  Distilled or deionized water to 1000 ml

(ii) Autoclave and dispense the medium into Petri plates.


Kritzman’s (modified) agar [adapted from Kritzman and Netzer (22)]

(i) Place the following ingredients into a suitable flask:

  NaNO3 1.0 g
  K2HPO4 0.9 g
  MgSO4·7H2O 0.2 g
  KCl 0.15 g
  Glucose 20.0 g
  Agar 25.0 g
  Distilled or deionized water to 1000 ml

(ii) Autoclave the suspension.

(iii) Add the following ingredients to the autoclaved suspension, starting with the tannic acid and ensuring it is dissolved completely before adding the copper sulfate:

  Tannic acid 5.0 g
  CuSO4.hydrated 1.7 g

(iv) Adjust the pH of the medium to 6.0 using 1 N NaOH.

(v) Autoclave, cool the medium to 50°C, then add the following ingredients just prior to pouring:

  PCNB (99%) 0.005 g
  Dithane (8%) 0.005 g
  Chloramphenicol 0.025 g

(vi) Dispense the medium into Petri plates.


Pectin Agar

(i) Mix the following ingredients in a blender for 1 to 2 minutes, until the pectin dissolves:

  NH4H2PO4 0.9 g
  (NH4)2HPO4 2.0 g
  MgSO4·7H2O 0.1 g
  KCl 0.5 g
  Citrus Pectin 10 g
  Agar 30 g
  Distilled or deionized water to 1000 ml

(ii) Adjust the suspension to pH 4.0 using HCl.

(iii) Autoclave and dispense the medium into Petri plates.