Link to home

​Bacterial Streak and Bulb Rot of Onion​

     
     
   

Mei Zhao, Brian Kvitko, Ron Gitaitis, and Bhabesh Dutta 2021. Bacterial Streak and Bulb Rot of Onion. The Plant Health Instructor. DOI: 10.1094/PHI-E​​-2021-0421-01​

Mei Zhao1, Brian Kvitko2, Ron Gitaitis1, and Bhabesh Dutta1

1Department of Plant Pathology, Coastal Plain Experiment Station, University of Georgia, Tifton, GA 31793;

2Department of Plant Pathology, University of Georgia, Athens, GA 30602

Corresponding author: Bhabesh Dutta, email: bhabesh@uga.edu

DISEASE: Bacterial Streak and Bulb Rot of Onion

PATHOGEN:Pseudomonas viridiflava (Burkholder) Dowson

HOSTS: Onion (Allium cepa L).​


Bacterial streak and bulb rot of onion has great destructive potential to cause significant economic losses to onion production due to foliage damage and bulb decay in the field and during storage.

Symptoms

In onion, leaf symptoms initially appear as oval lesions or streaks resulting in the total collapse of the entire leaf (Fig 1). Initially, streaks are water-soaked but later turn brown or dark green to almost black lesions if near the base of infected leaves. If not rotted, at later stages of disease development, collapsed leaf areas can also have a distinctly ribbed or corrugated appearance due to protruding veins (Fig 2). Infection may gradually progress down to the base of the leaf and into the bulb. Infected inner scales are initially lemon yellow that can quickly turn from reddish-brown to dark brown in harvested bulbs (Fig 3). On rare occasions, infected tissues can be blue-green.


Fig 1. Foliar symptoms of bacterial streak and bulb
rot caused by
Pseudomonas viridiflava with
oval-shaped water-soaked lesions.


Fig 2. Foliar symptoms of bacterial streak and bulb rot caused by Pseudomonas viridiflava with water-soaked lesion and collapsed leaf tissues. Corrugated appearance due to protruding veins in leaves.
​​
Fig 3. Onion bulb symptoms caused by Pseudomonas viridiflava under natural light (A) and UV light (B). 

Host range and geographic distribution

Pseudomonas viridiflava (Latin viridis green, flava yellow) is pathogenic to numerous cultivated crops and weeds. Burkholder first described P. viridiflava as a pathogen of bean (Phaseolus vulgaris) (Burkholder 1930). Since then, it has been reported around the world on many plants, both as a disease-causing agent in the field and also as a post-harvest pathogen (Liao and Wells 1987). The hosts include:

  • ​​Acanthus mollis (Sarris et al. 2012)
  • alfalfa (Medicago sativa) (Lukezic et al. 1983)
  • apple (Malus domestica) (Alimi et al. 2011)
  • apricot (Prunus armeniaca) (Bradbury 1986)
  • Arabidopsis thaliana (Jakob et al. 2002)
  • artichoke (Cynara scolymus) (Sarris et al. 2012)
  • basil (Ocimum basilicum) (Little et al. 1994)
  • birdsfoot trefoil (Lotus corniculatus) (Lukezic et al. 1983)
  • blite (Amaranthus blitum) (Goumans and Chatzaki 1998)
  • broccoli (Brassica oleracea var. italica) (Bradbury 1986)
  • brussels sprout (Brassica oleracea var. gemmifera) (Bradbury 1986)
  • cabbage (Brassica oleracea var. capitata) (Bradbury 1986)
  • calla lily (Zantedeschia sp.) (Basavand et al. 2019)
  • cauliflower (Brassica oleracea var. botrytis) (Wilkie et al. 1973)
  • celery (Apium graveolens) (Bradbury 1986)
  • cherry (Prunus sp.) (Billing 1970)
  • chicory (Cichorium intybus) (Caruso and Catara 1996)
  • Chinese gooseberry (Actinidia chinensis) (Wilkie et al. 1973)
  • chrysanthemum (Chrysanthemum morifolium) (Billing 1970)
  • Chrysanthemum coccineum (Bradbury 1986)
  • cucumber (Cucumis sativus) (Bradbury 1986)
  • Delphinium cultorum (Bradbury 1986)
  • dill (Anethum graveolens) (Bradbury 1986)
  • eggplant (Solanum melongena) (Goumans and Chatzaki 1998)
  • garlic (Allium sativum) (Pérez et al. 2004)
  • grape (Vitis vinifera) (Wilkie et al. 1973)
  • hellebore (Helleborus × hybridus) (Taylor et al. 2011)
  • Hydrangea sp. (Bradbury 1986)
  • lettuce (Lactuca sativa) (Bradbury 1986)
  • lupin (Lupinus angustifolius) (Wilkie et al. 1973)
  • marigold (Calendula officinalis) (Moretti et al. 2012)
  • melon (Cucumis melo) (Goumans and Chatzaki 1998)
  • Mexican heather (Cuphea hyssopifolia) (Albu et al. 2018)
  • parsley (Petroselinum crispum)
  • parsnip (Pastinaca sativa) (Hunter and Cigna 1981)
  • passion fruit (Passiflora edulis) (Wilkie et al. 1973)
  • pea (Pisum sativum) (Wilkie et al. 1973)
  • peach (Prunus persica) (Scortichini and Morone 1997)
  • pear (Pyrus sp.) (Billing 1970)
  • pepper (Capsicum annuum) (Bradbury 1986)
  • poinsettia (Euphorbia pulcherrima) (Suslow and McCain 1981)
  • poppy (Papaver somniferum) (Wilkie et al. 1973)
  • pumpkin (Cucurbita maxima) (Wilkie et al. 1973)
  • radish (Raphanus sativus) (Shakya and Vinther 1989)
  • rape (Brassica napus var. napus) (Bradbury 1986)
  • sleeping hibiscus (Malvaviscus penduliflorus) (Rouhrazi and Rahimian 2012)
  • tomato (Solanum lycopersicum) (Jones et al. 1984;  Wilkie et al. 1973)
  • watermelon (Citrullus lanatus) (Mirik et al. 2004)

In addition to the above list, the bacterium also infects the following upon artificial inoculation (Bradbury 1986):

  • ​​chard (Beta vulgaris) ​
  • Calendula officinalis
  • safflower (Carthamus tinctorius)
  • cockscomb (Celosia cristata)
  • Eschscholzia californica
  • Euchlaena mexicana, buckwheat (Fagopyrum esculentum)
  • soybean (Glycine max)
  • hyacinth bean (Lablab purpureus)
  • Lactuca dracoglossa
  • Lagenaria siceria var. hispida
  • Papaver nudicaule
  • Petunia hybrida
  • runner bean (Phaseolus coccineus)
  • lima bean (Phaseolus lunatus)
  • Pueraria hirsute
  • Japanese radish (Raphanus sativus var.acanthiformis)
  • Sorghum sudanense
  • Sorghum vulgare
  • spinach (Spinacia oleracea)
  • red clover (Trifolium pratense)
  • Tropaeolum majus
  • broad bean (Vicia faba)
  • adzuki bean (Vigna angularis)
  • cowpea (Vigna unguiculata)
  • maize (Zea mays)
  • zinnia (Zinnia elegans)

Finally, the bacterium has been recovered from several weed species in the onion-growing area in Georgia, USA as a resident epiphyte. These weeds include cutleaf evening primrose (Oenothera laciniata), dandelion (Taraxacum officinale), common fumitory (Fumaria officinalis), purple cudweed (Gnaphalium purpureum), spiny sowthistle (Sonchus asper), narrowleaf vetch (Vicia sativa), Virginia pepperweed (Lepidium virginicum), and wild radish (Raphanus raphanistrum) (Gitaitis et al. 1998). ​

Bacterial streak and bulb rot caused by P. viridiflava was first reported on onion in Georgia USA in 1990 (Gitaitis et al. 1991), and since then, has been reported elsewhere in the USA including Colorado (Schwartz and Otto 1998), Florida (Mark et al. 2002), and Pennsylvania (Pfeufer 2014). The disease has also been reported in many onion producing countries around the world, including New Zealand (Wright and Grant 1998), Venezuela (Mark et al. 2002), Uruguay on onion and garlic (Pérez et al. 2004), Korea (Lee et al. 2015), and Australia (Gambley 2018).

Pathogen biology and molecular identification

P. viridiflava is a Gram-negative, aerobic, rod-shaped bacterium (Gitaitis et al. 1991). It forms cream colonies that turn yellow with age and produces a diffusible pigment that appears yellowish-green under visible light and fluorescent blue under UV light (Sarris et al. 2012) on King's medium B agar (KMB) (King et al. 1954). Many P.viridiflava strains from onion were copper-tolerant (Gitaitis et al. 1991). Besides nutrient agar and KMB as a routine isolation medium, a semi-selective agar medium (T5) was developed and used to sample the environment and plant material as a habitat for P. viridiflava (Gitaitis et al. 1997).

Initial steps in identification are by differentiating phenotypic characteristics. Typically, P. viridiflava is levan negative, oxidase negative, potato rot positive, arginine dihydrolase negative, positive for a hypersensitive response (HR) on tobacco (LOPAT), does not utilize sucrose, and is ice nucleation active (Gitaitis et al. 1991). Although LOPAT tests are the most widely used protocol for differentiating plant pathogenic pseudomonads, a study examining P. viridiflava strains (n = 59) from diverse origins found that only 37% showed the typical P. viridiflava LOPAT profile (Bartoli et al. 2014). Thus, phenotypic traits may not be strictly diagnostic. In addition to the phenotypic diversity among strains, more than 10 different P. viridiflava strains were observed to produce two phenotypic variants (mucoid and transparent colonies) (Bartoli et al. 2014, 2015). Bartoli et al. (2015) demonstrated that transparent colony variants evolved antibiotic resistance (amoxicillin, ampicillin, kanamycin, and rifamycin) at a higher rate and failed to induce disease on bean.

Species identification is accomplished by a combination of pathogenicity and molecular tests. Pathogenicity tests of P. viridiflava on onion have been conducted using spray-inoculation with sprayers and stab-inoculation with toothpicks (Gitaitis et al. 1991). Leaf surfaces of 6 to 8-week-old onion seedlings were spray-inoculated with a bacterial suspension (~1 x 108 CFU/ml); onion plants were also stab-inoculated into the base of the leaves immediately above the neck of the bulb with toothpicks that touched a P. viridifalva colony. After inoculation, plants were incubated in a mist chamber for an additional 24 h and then incubated under high relative humidity and a mean temperature of 20-25°C. With both inoculation methods, inoculated plants were observed after 10-14 days for symptom development (foliar leaf spots, streaks, and soft rot of basal areas similar to those observed in the field as described in the Symptom section). Gitaitis et al. (1991) reported that onion plants predisposed by mist treatments or inoculated by toothpick method displayed typical symptoms caused by P. viridiflava while plants with no mist treatment before inoculation showed less severe or no symptoms. For pathogenicity tests, a pathogenic reference strain and sterile water/buffer should be used to inoculate plants as a positive and negative control, respectively.

Two primer sets were developed for P. viridiflava. One primer set targeting the pectate lyase gene amplified 95% of the strains from onions and weeds in Georgia, USA but failed to amplify P. viridiflava strains from bean, bell pepper, parsnip, tomato, and watermelon (Gay et al. 1998). Another primer set (ORF1/2-Fw, ORF1/2-Rv) targeted the monooxygenase and lipoprotein genes unique to the P. viridiflava members belonging to phylogroup 7 (Bartoli et al. 2014). P. viridiflava are found in phylogroup 7 and 8 (based on the similarity of housekeeping genes) (Bartoli et al. 2014;  Parkinson et al. 2011) and genomospecies 6 (based on DNA-DNA hybridization similarity) (Gardan et al. 1999). ORF1/2 primer set showed false-positive results with 3% (6/174) of P. syringae field isolates, which belonged to phylogroup 10 by cts sequences (Borschinger et al. 2016). On the other hand, a false-negative was observed in one out of twelve (8%) field isolates classified as phylogroup 7 by cts sequences (Borschinger et al. 2016). The authors attributed the discrepancy to isolates from different phylogroups (Borschinger et al. 2016).

Multilocus sequence analysis (MLSA) should be used to identify P. viridiflava and compared to the type strain (P. viridiflava ATCC13223T). MLSA or multilocus sequence typing (MLST) was validated in the P. syringae group first with seven housekeeping genes (Sarkar and Guttman 2004), then with four genes, cts, gapA, gyrB, and rpoD (Hwang et al. 2005). These gene loci provide a simple and rapid phylogenetic framework for identifying and classifying potential Pseudomonas spp. The Plant-Associated and Environmental Microbes Database (PAMDB) (http://genome.ppws.vt.edu/cgi-bin/MLST/search_alleles.pl) facilitates the discrimination of plant-associated bacteria based on multilocus sequence data (Almeida et al. 2010). The use of one housekeeping gene, such as partial sequences of rpoD (Parkinson et al. 2011) or cts (Berge et al. 2014), has been proposed to be sufficient to predict the phylogroup affiliation.

Disease cycle and epidemiology

The primary source of inoculum for bacterial streak and bulb rot of onion in Georgia, USA was from weeds (Gitaitis et al. 1998). A survey conducted from 1991-1995 in an onion-growing area in Georgia, USA found that P. viridiflava can survive on several weed species (please see above), especially on cutleaf evening primrose, but found no evidence of P. viridiflava in soil, water, or in association with volunteer soybeans (Gitaitis et al. 1998). However, P. viridiflava was isolated from stream water, snowfall, epilithic biofilm, rain, river water, and irrigation water in France, but has not been isolated from soil so far (Bartoli et al. 2014). In addition, P. viridiflava was detected in 76.6% of onion seed samples by enzyme-linked immunosorbent assay (Rodríguez and Contreras 2006).

In the field, the onions are predisposed to environmental conditions and wounds that are favorable for the infection by this opportunistic pathogen. In Georgia USA, the disease is favored by excessive fertilization and prolonged periods of rain during the cooler winter months of onion production (Gitaitis et al. 1998). Onions are at risk for infections from the first week of January until mid-March, but are vulnerable anytime during the growing season when conditions are favorable. However, this disease occurs in Venezuela and Colorado USA under warmer conditions (Mark et al. 2002). This disease predisposes onion bulbs to other soft rot pathogens and secondary or opportunistic organisms associated with post-harvest decays. If infected bulbs are harvested and not discarded during grading, they will develop a post-harvest rot, which can spread to healthy bulbs in storage. A diagram of a disease cycle depicting the above features is presented in fig. 4.


Fig. 4. A proposed disease cycle of bacterial streak and bulb rot in onion caused by Pseudomonas viridiflava.

Disease management options

Field trials in Georgia demonstrated a three-fold reduction in disease incidence when weeds were effectively controlled in the field (Gitaitis et al. 2003). It may be advisable to control weeds along field borders and in irrigation strips if using fixed irrigation. Crop rotation and the sanitation of onion crop debris, culls, and volunteer plants may also reduce sources of inoculum in the fields (Gitaitis et al. 1991). It is advisable not to enter fields when leaf wetness is present. Many onion strains of P. viridiflava were copper tolerant, but sensitive to copper and mancozeb mixtures (Gitaitis et al. 1991). Preventive application of fixed copper materials tank-mixed with EBDC fungicides (Mankocide) significantly reduces the incidence and spread of this disease (Gitaitis et al. 2003). Avoiding over-fertilization with N during winter months may reduce bacterial streak losses (Gitaitis et al. 2003). Also, practices that reduce post-harvest rots such as harvesting onions at a proper stage of maturity, field-curing onions for a minimum of 48 hours under dry conditions before clipping, and avoiding bruising or wounding at all times will help reduce disease problems (Gitaitis et al. 2003). Post-harvest rot due to this disease was significantly reduced (7% rot vs. 55% rot) in onions that were stored under controlled atmospheric conditions vs. typical cold storage (4°C) (Gitaitis et al. 2003).

Genome resources and population structure of P. viridiflava

Whole-genome sequence data (n = 1,530 genomes) are available in Genbank under the name for P. viridiflava (https://www.ncbi.nlm.nih.gov/genome/browse/#!/prokaryotes/11288/). The majority of draft genome sequences were from a large-scale survey of wild A. thaliana in Germany (Karasov et al. 2018). The authors found strains of P. viridiflava to be the most abundant operational taxonomical unit in A. thaliana but genetically diverse, diverged from one another by approximately 300,000 years (Karasov et al. 2018). Other sequenced P. viridiflava strains available were isolated from plant hosts cherry, bean, radish, Primula officinalis, Lepidium draba, and Rhododendron sp. as well as from precipitation and epilithon (underwater rock biofilm). Although P. viridiflava isolates from onion have not yet been sequenced, the currently available genome sequences might be used to design improved diagnostic PCR primers.

P.viridiflava from diverse sources was shown to display a high level of genetic variation worldwide (Goss et al. 2005;  Sarris et al. 2012;  Bartoli et al. 2014). Goss et al. (2005) divided P. viridiflava strains from A. thaliana worldwide into two distinct clades (A and B), with evidence of frequent recombination within clades. Sarris et al. (2012) also found high genetic variation among local P. viridiflava strains (Crete, Greece) and showed that the local celery strains clustered separately from other local and global P. viridiflava strains isolated from different hosts. Bartoli et al. (2014) characterized P. viridiflava strains from different hosts in the Mediterranean region. The authors observed two distinct phylogroups (7 and 8) in P. viridiflava with wide genetic variability.

Virulence factors and plant-pathogen interaction

P. viridiflava uses multiple virulence factors during the interactions with host plants. At different infection stages, P. viridiflava requires functional motility to move to and in host tissues, pectate lyase enzyme(s) and type II secretion system (T2SS) to break down plant cell walls, and the type III secretion system (T3SS) to colonize plant tissues.

Czelleng et al. (2006) used an IVET (in vivo expression technique) method based on plasmid pIviGK to identify virulence-associated genes of P. viridiflava during infection of green pepper fruits. The authors identified the following genes as strongly induced in plants: pel (pectate lyase), mviN (lipid II flippase), deoxyguanosine-triphosphate triphosphohydrolase (dGTPase), an ABC transporter, and a K-dependent Na+/Ca2+ exchange related-protein (Czelleng et al. 2006). The marker-exchange mutant of 'mviN' showed a non-motile phenotype, and reduced growth in A. thaliana by dip inoculation compared to the wild-type strain. However, direct inoculation of the apoplast via blunt syringe was not associated with virulence defects. The results suggested that the mviN contributes to P. viridiflava invasion of plant tissues.

A single alkaline extracellular pectate lyase of P. viridiflava strain SF312A was responsible for the maceration of plant tissues (Liao et al. 1988). Mutation of 'pel' and the 'out' (T2SS) loci that direct the production and secretion of pectate lyase in strain SF312A resulted in the loss of the soft rot phenotype in five different plant hosts (potato tubers, celery petioles, fruits of bell pepper, tomato, and squash) (Liao et al. 1988). Pectate lyase from P. viridiflava strain RMX23.1a also contributed to bacterial virulence and symptom development in A. thaliana (Jakob et al. 2007). Moreover, pectate lyase activities varied among P. viridiflava strains. The P. viridiflava strains from clade B showed significantly higher pectate lyase activity on agar medium and induced severer symptoms in A. thaliana than clade A strains (Goss and Bergelson 2006).

The histidine kinase LemA/RepA and the response regulator GacA/RepB were both observed to be required for the production of extracellular pectate lyase, proteases, and the exopolysaccharide alginate by P. viridiflava strains PJ-08-6A and SF312A (Liao et al. 1994;  Liao et al. 1996). In addition, gacA/repB but not lemA/repA of P. viridiflava strain PJ-08-6A also regulated the production of fluorescent siderophores (Liao et al. 1996).

In most Gram-negative plant pathogenic bacteria, the T3SS is a major virulence factor that translocates bacterial effector virulence proteins directly into plant cells. The T3SS is typically encoded within a pathogenicity island (PAI). Strains of P. viridiflava encode a Hrp1 class T3SS gene cluster in one of two alternate PAIs, either an S-PAI (for single) or T-PAI (for tripartite) based on the arrangement of the T3SS structural and conserved effector genes relative to one another. None of the 286 P. viridiflava strains tested carried both types of PAIs (Araki et al. 2006). Both S-PAI and T-PAI strains produced T3SS pili when grown in T3SS inducing medium (Jakob et al. 2007). Mutation of  P. viridiflava strain LP23.1a hrpL ECF (extracytoplasmic function) sigma factor gene driving expression of the Hrp1 T3SS structural genes and T3SS effectors resulted in complete loss of the HR on tobacco and reduced bacterial population during A. thaliana infection (Jakob et al. 2007). However, pathogenicity-related traits of P. viridiflava are not strictly linked to the two T3SS configurations (S/T-PAI), but were suggested to correlate with the presence or absence of the type 3 secreted effector avrE (Bartoli et al. 2014). The direct contribution of AvrE to P. viridiflava virulence awaits experimental validation.

Studies on host responses to P. viridiflava are limited. Since P. viridiflava is a natural pathogen and shows extensive genetic variation of resistance on the model plant species A.thaliana (Goss and Bergelson 2006), many studies on P. viridiflava-host interactions have been focused on A. thaliana. P. viridiflava induced strong jasmonic acid-mediated and relatively weak salicylic acid-mediated defense responses in A. thaliana (Jakob et al. 2007). In addition, apoplastic accumulation and further oxidation of tetraamine spermine were shown to protect tobacco (Marina et al. 2008) and A.thaliana against P. viridiflava strain Pvalb8 (Gonzalez et al. 2011). The contributions of specific P. viridiflava virulence factors have yet to be investigated in the onion pathosystem. Nevertheless, evaluation of Allium varieties and accessions against P. viridiflava has been conducted. In 1998, 85 accessions of 33 species of Allium were screened for susceptibility to foliar bacterial soft rot caused by P. marginalis and P. viridiflava by co-inoculation after wounding (Wright and Grant 1998). The highly resistant Allium accession was A. sativum 'TNL' (Wright and Grant 1998). After combining the data for each species' accessions, A. vavilovii and A. cepa were highly susceptible, while A. rotundum and A. cernuum were highly resistant (Wright and Grant 1998). Identifying and introducing genetic resistance against P. viridiflava from other resistant Allium species into A. cepa cultivars may provide another critical tool for managing this disease.

Significance

Bacterial streak and bulb rot of onion has been a problem in onion-growing regions of Georgia, USA since 1990. This disease can be a major limiting factor in onion production in other parts of the world. Weed species have been shown to serve as important inoculum sources for P. viridiflava (Gitaitis et al. 1998). Although P. viridiflava was detected from onion seeds (Rodríguez and Contreras 2006), no further studies were conducted to investigate the role of seeds in pathogen survival and transmission. Moreover, little is known about the population structure and virulence mechanisms of P. viridiflava from onions. Understanding the population diversity of P. viridiflava strains and how they infect onion foliage and bulbs are critical to developing integrated disease management options. Future studies on these topics will shed light on breeding for host resistance and improved disease management approaches.​

Funding information

This work is supported by the Specialty Crops Research Initiative Award 2019-51181-30013 from the USDA National Institute of Food and Agriculture. Any opinions, findings, conclusions, or recommendations expressed in this publication are those of the author(s) and do not necessarily reflect the view of the U.S. Department of Agriculture.

References

Albu, S., Lai, M., Woods, P., and Kumagai, L. 2018. First Report of Pseudomonas viridiflava as the Causal Agent of Bacterial Leaf Spot of Mexican Heather in California. Plant Dis. 102:2633.

Alimi, M., Rahimian, H., Hassanzadeh, N., Darzi, M. T., Ahmadikhah, A., Heydari, A., and Balestra, G. M. 2011. First Detection of Pseudomonas viridiflava, the Causal Agent of Blossom Blight in Apple by Using Specific Designed Primers. Afr. J. Microbiol. Res. 5:4708-4713.4702.

Almeida, N. F., Yan, S., Cai, R., Clarke, C. R., Morris, C. E., Schaad, N. W., Schuenzel, E. L., Lacy, G. H., Sun, X., and Jones, J. B. 2010. Pamdb, a Multilocus Sequence Typing and Analysis Database and Website for Plant-Associated Microbes. Phytopathology 100:208-215.

Araki, H., Tian, D., Goss, E. M., Jakob, K., Halldorsdottir, S. S., Kreitman, M., and Bergelson, J. 2006. Presence/Absence Polymorphism for Alternative Pathogenicity Islands in Pseudomonas viridiflava, a Pathogen of Arabidopsis. Proc. Natl. Acad. Sci. U. S. A. 103:5887-5892.

Bartoli, C., Lamichhane, J. R., Berge, O., Varvaro, L., and Morris, C. E. 2015. Mutability in Pseudomonas viridiflava as a Programmed Balance between Antibiotic Resistance and Pathogenicity. Mol. Plant Pathol. 16:860-869.

Bartoli, C., Berge, O., Monteil, C. L., Guilbaud, C., Balestra, G. M., Varvaro, L., Jones, C., Dangl, J. L., Baltrus, D. A., and Sands, D. C. 2014. The Pseudomonas viridiflava Phylogroups in the P. syringae Species Complex Are Characterized by Genetic Variability and Phenotypic Plasticity of Pathogenicity‐Related Traits. Environ. Microbiol. 16:2301-2315.

Basavand, E., Khodaygan, P., and Rahimian, H. 2019. First Report of Bacterial Leaf Spot on Calla Lily (Zantedeschia Spp.) Caused by Pseudomonas viridiflava in Iran. J. Plant Pathol. 101:393-393.

Berge, O., Monteil, C. L., Bartoli, C., Chandeysson, C., Guilbaud, C., Sands, D. C., and Morris, C. E. 2014. A User's Guide to a Data Base of the Diversity of Pseudomonas syringae and Its Application to Classifying Strains in This Phylogenetic Complex. PloS One 9.

Billing, E. 1970. Pseudomonas viridiflava (Burkholder, 1930; Clara 1934). J. Appl. Microbiol 33:492-500.

Borschinger, B., Bartoli, C., Chandeysson, C., Guilbaud, C., Parisi, L., Bourgeay, J., Buisson, E., and Morris, C. 2016. A Set of PCRs for Rapid Identification and Characterization of Pseudomonas syringae Phylogroups. J. Appl. Microbiol. 120:714-723.

Bradbury, J. F. 1986. Description of Pseudomonas viridiflava. Pages 183-184 in: Guide to Plant Pathogenic Bacteria. CAB international.

Burkholder, W. H. 1930. The Bacterial Diseases of the Bean. A Comparative Study. Memoir. Cornell Agricultural Experiment Station 127.

Caruso, P., and Catara, V. 1996. First Report of Pseudomonas viridiflava Leaf Spot of Red-Leaved Chicory. Plant Dis. 80.

Czelleng, A., Bozso, Z., Ott, P., Besenyei, E., Varga, G., Szatmari, A., Kiraly, L., and Klement, Z. 2006. Identification of Virulence-Associated Genes of Pseudomonas viridiflava Activated During Infection by Use of a Novel Ivet Promoter Probing Plasmid. Curr. Microbiol. 52:282-286.

Gambley, C. 2018. Detection and Management of Bacterial Diseases in Australian Allium Crops.

Gardan, L., Shafik, H., Belouin, S., Broch, R., Grimont, F., and Grimont, P. 1999. DNA Relatedness among the Pathovars of Pseudomonas syringae and Description of Pseudomonas tremae sp. Nov. And Pseudomonas cannabina Sp. Nov.(Ex Sutic and Dowson 1959). Int. J. Syst. Evol. Microbiol. 49:469-478.

Gay, J. D., Gitaitis, R. D., Maw, B. W., Sumner, P. E., Sumner, D. R., Torrance, R. L., Randle, B., Tollner, E. W., Schmidt, N. E., and Batal, K. D. 1998. Georgia Onion Res.-Ext. Report 1996-97.

Gitaitis, R.D., Sanders, F.H., Diaz-Perez, J.C., and Walcott. R.R. 2003. Integrated management of bacterial streak and bulb rot of onion. Pages 443-449 in: Pseudomonas syringae and related pathogens Biology and Genetics. N.S. Iacobellis, A. Collmer, S.W. Hutcheson, J.W. Mansfield, C.E. Morris, J. Murillo, N.W. Schaad, D. E. Stead, G. Surico and M.S. Ullrich, editors. Kluwer Academic Publishers, Dordrecht, The Netherlands. 708 pp.

Gitaitis, R., Baird, R., Beaver, R., Sumner, D., Gay, J., and Smittle, D. 1991. Bacterial Blight of Sweet Onion Caused by Pseudomonas viridiflava in Vidalia, Georgia. Plant Dis. 75:1180-1182.

Gitaitis, R., MacDonald, G., Torrance, R., Hartley, R., Sumner, D., Gay, J., and Johnson III, W. 1998. Bacterial Streak and Bulb Rot of Sweet Onion: Ii. Epiphytic Survival of Pseudomonas viridiflava in Association with Multiple Weed Hosts. Plant Dis. 82:935-938.

Gitaitis, R., Sumner, D., Gay, D., Smittle, D., McDonald, G., Maw, B., Johnson III, W., Tollner, B., and Hung, Y. 1997. Bacterial Streak and Bulb Rot of Onion: I. A Diagnostic Medium for the Semiselective Isolation and Enumeration of Pseudomonas viridiflava. Plant Dis. 81:897-900.

Gonzalez, M. E., Marco, F., Minguet, E. G., Carrasco-Sorli, P., Blázquez, M. A., Carbonell, J., Ruiz, O. A., and Pieckenstain, F. L. 2011. Perturbation of Spermine Synthase Gene Expression and Transcript Profiling Provide New Insights on the Role of the Tetraamine Spermine in Arabidopsis Defense against Pseudomonas viridiflava. Plant Physiol. 156:2266-2277.

Goss, E. M., and Bergelson, J. 2006. Variation in Resistance and Virulence in the Interaction between Arabidopsis thaliana and a Bacterial Pathogen. Evolution 60:1562-1573.

Goss, E. M., Kreitman, M., and Bergelson, J. 2005. Genetic Diversity, Recombination and Cryptic Clades in Pseudomonas viridiflava Infecting Natural Populations of Arabidopsis thaliana. Genetics 169:21-35.

Goumans, D., and Chatzaki, A. 1998. Characterization and Host Range Evaluation of Pseudomonas viridiflava from Melon, Blite, Tomato, Chrysanthemum and Eggplant. Eur. J. Plant Pathol. 104:181-188.

Hunter, J., and Cigna, J. 1981. Bacterial Blight Incited in Parsnip by Pseudomonas Marginalis and Pseudomonas viridiflava. Phytopathology 71:1238-1241.

Hwang, M. S., Morgan, R. L., Sarkar, S. F., Wang, P. W., and Guttman, D. S. 2005. Phylogenetic Characterization of Virulence and Resistance Phenotypes of Pseudomonas syringae. Appl. Environ. Microbiol. 71:5182-5191.

Jakob, K., Kniskern, J. M., and Bergelson, J. 2007. The Role of Pectate Lyase and the Jasmonic Acid Defense Response in Pseudomonas viridiflava Virulence. Mol. Plant-Microbe Interact. 20:146-158.

Jakob, K., Goss, E. M., Araki, H., Van, T., Kreitman, M., and Bergelson, J. 2002. Pseudomonas viridiflava and P. syringae—Natural Pathogens of Arabidopsis thaliana. Mol. Plant-Microbe Interact. 15:1195-1203.

Jones, J., Jones, J., McCarter, S., and Stall, R. 1984. Pseudomonas viridiflava: Causal Agent of Bacterial Leaf Blight of Tomato. Plant Dis. 68:341-342.

Karasov, T. L., Almario, J., Friedemann, C., Ding, W., Giolai, M., Heavens, D., Kersten, S., Lundberg, D. S., Neumann, M., and Regalado, J. 2018. Arabidopsis thaliana and Pseudomonas Pathogens Exhibit Stable Associations over Evolutionary Timescales. Cell Host Microbe 24:168-179. e164.

King, E. O., Ward, M. K., and Raney, D. E. 1954. Two Simple Media for the Demonstration of Pyocyanin and Fluorescin. J Lab Clin Med. 44:301-307.

Lee, J., Hwang, S., Ha, I., Min, B., Hwang, H., and Lee, S. 2015. Comparison of Bulb and Leaf Quality, and Antioxidant Compounds of Intermediate-Day Onion from Organic and Conventional Systems. Hortic. Environ. Biotechnol. 56:427-436.

Liao, C., and Wells, J. 1987. Diversity of Pectolytic, Fluorescent Pseudomonads Causing Soft Rots of Fresh Vegetables at Produce Markets. Phytopathology 77:673-677.

Liao, C., McCallus, D., and Fett, W. 1994. Molecular Characterization of Two Gene Loci Required for Production of the Key Pathogenicity Factor Pectate Lyase in Pseudomonas viridiflava. Mol. Plant-Microbe Interact. 7:391-400.

Liao, C.-H., Hung, H.-Y., and Chatterjee, A. K. 1988. An Extracellular Pectate Lyase Is the Pathogenicity Factor of the Soft-Rotting Bacterium Pseudomonas viridiflava. Mol. Plant-Microbe Interact 1:199-206.

Liao, C.-H., McCallus, D., Wells, J., Tzean, S.-S., and Kang, G.-Y. 1996. The Repb Gene Required for Production of Extracellular Enzymes and Fluorescent Siderophores in Pseudomonas viridiflava Is an Analog of the Gaca Gene of Pseudomonas syringae. Can. J. Microbiol. 42:177-182.

Little, E., Gilbertson, R., and Koike, S. 1994. First Report of Pseudomonas viridiflava Causing a Leaf Necrosis on Basil. Plant Dis. 78.

Lukezic, F., Leath, K., and Levine, R. 1983. Pseudomonas viridiflava Associated with Root and Crown Rot of Alfalfa and Wilt of Birdsfoot Trefoil. Plant Dis. 67:808-811.

Marina, M., Maiale, S. J., Rossi, F. R., Romero, M. F., Rivas, E. I., Gárriz, A., Ruiz, O. A., and Pieckenstain, F. L. 2008. Apoplastic Polyamine Oxidation Plays Different Roles in Local Responses of Tobacco to Infection by the Necrotrophic Fungus Sclerotinia sclerotiorum and the Biotrophic Bacterium Pseudomonas viridiflava. Plant Physiol. 147:2164-2178.

Mark, G., Gitaitis , R., and Lorbeer, J. 2002. 11 Bacterial Diseases of Onion. Pages 267-292. In: Allium Crop Science: Recent Advances. (ed) H.D. Rabinowitch and L. Currah. CABI Publishing, New York, NY. 515 pp.

Mirik, M., Aysan, Y., Cetinkaya-Yildiz, R., Sahin, F., and Saygili, H. 2004. Watermelon as a New Host of Pseudomonas viridiflava, Causal Agent of Leaf and Stem Necrosis, Discovered in Turkey. Plant Dis. 88:907-907.

Moretti, C., Fakhr, R., and Buonaurio, R. 2012. Calendula Officinalis: A New Natural Host of Pseudomonas viridiflava in Italy. Plant Dis. 96:285-285.

Parkinson, N., Bryant, R., Bew, J., and Elphinstone, J. 2011. Rapid Phylogenetic Identification of Members of the Pseudomonas syringae Species Complex Using the rpoD Locus. Plant Pathol. 60:338-344.

Pérez, F., Silvera, P., and Gepp, W. 2004. Pseudomonas viridiflava (Burkholder) Dowson: Causal Agent of Necrotic Spots on Onion and Garlic (Allium spp.) Leaves in Uruguay. Agrociencia (Montevideo) 8:33-37.

Pfeufer, E. E. 2014. Sources of Inoculum, Epidemiology, and Integrated Management of Bacterial Rots of Onion (Allium cepa) with a Focus on Center Rot, Caused by Pantoea ananatis and Pantoea agglomerans. Ph.D. diss., Pennsylvania State University, University Park, PA. https://etda.libraries.psu.edu/ files/final_submissions/9951

Rodríguez, C., and Contreras, N. 2006. Elisa Technique for the Detection of Phytopathogenic Bacteria in Onion (Allium cepa L.) Seeds. Pages 21-32 in: Proceedings of the Interamerican Society for Tropical Horticulture Interamerican Society for Tropical Horticulture.

Rouhrazi, K., and Rahimian, H. 2012. Sleeping Hibiscus a New Host for Pseudomonas viridiflava. J. Plant Pathol. 94.

Sarkar, S. F., and Guttman, D. S. 2004. Evolution of the Core Genome of Pseudomonas syringae, a Highly Clonal, Endemic Plant Pathogen. Appl. Environ. Microbiol. 70:1999-2012.

Sarris, P. F., Trantas, E. A., Mpalantinaki, E., Ververidis, F., and Goumas, D. E. 2012. Pseudomonas viridiflava, a Multi Host Plant Pathogen with Significant Genetic Variation at the Molecular Level. PloS One 7:e36090.

Schwartz, H., and Otto, K. 1998. Onion Bacterial Disease Management in Colorado. Pages 214-218 in: Proc. 1998 Natl. Onion (and other Allium) Res. Conf, Sacramento, CA.

Scortichini, M., and Morone, C. 1997. Apoplexy of Peach Trees Caused by Pseudomonas viridiflava. J Phytopathol. 145:397-399.

Shakya, D. D., and Vinther, F. 1989. Occurrence of Pseudomonas viridiflava in Seedlings of Radish. J Phytopathol. 124:123-127.

Suslow, T., and McCain, A. 1981. Greasy Canker of Poinsettia Caused by Pseudomonas viridiflava. Plant Dis. 65:513-514.

Taylor, R. K., Romberg, M. K., and Alexander, B. J. 2011. A Bacterial Disease of Hellebore Caused by Pseudomonas viridiflava in New Zealand. Australas. Plant Dis. Notes 6:28-29.

Wilkie, J. P., Dye, D., and Watson, D. 1973. Further Hosts of Pseudomonas viridiflava. N. Z. J. Agric. Res. 16:315-323.

Wright, P., and Grant, D. 1998. Evaluation of Allium Germplasm for Susceptibility to Foliage Bacterial Soft Rot Caused by Pseudomonas marginalis andPseudomonas viridiflava. N. Z. J. Crop Hortic. Sci. 26:17-21.